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Impact of free living protozoa and bacterial interactions on multispecies biofilm
PhD Thesis
Prem Krishnan Raghupathi
Promoters
Prof. Dr. Søren J. Sørensen
University of Copenhagen
Prof. Dr. Kurt Houf
Ghent University
Prof. Dr. Mette Burmølle
University of Copenhagen
Prof. Dr. Koen Sabbe
Ghent University
In collaboration with
Section of Microbiology
Department of Biology
University of Copenhagen
Denmark
&
Department of Veterinary Public Health and Food Safety
Laboratory of Microbiology
Faculty of Veterinary Medicine
Ghent University
Belgium
Raghupathi, PK. (2018). Impact of free living protozoa and bacterial interactions on multispecies biofilm.
Ph.D. Thesis, Ghent University, Belgium & University of Copenhagen, Denmark.
Copyright © 2018 Prem Krishnan Raghupathi
ISBN:
All Rights Reserved.
Printed By: University Press, 9185 Wachtebeke, Belgium |http://www.universitypress.be
This work was funded by the BOF Joint PhD Research Grant, Ghent University, Belgium and Faculty
Scholarships, University of Copenhagen, Denmark.
Publicly defended in Ghent, Belgium, on 29 August 2018.
Members of the Examination Board
Prof. Dr. EDWIN CLAEREBOUT
Department of Virology, Parasitology and Immunology
Faculty of Veterinary Medicine, Ghent University, Ghent, Belgium
Prof. Dr. ANNE WILLEMS
Laboratory of Microbiology
Faculty of Sciences, Ghent Unviersity, Ghent, Belgium
Prof. Dr. HANNE INGMER
Microbial Food Safety and Zoonosis
Department of Veterinary and Animal Sciences
Faculty of Health and Medical Sciences University of Copenhagen, Copenhagen, Denmark
Prof. Dr. TOM COYENE
Laboratory of Pharmaceutical Microbiology
Faculty of Pharmaceutical Sciences, Ghent Unviersity, Ghent, Belgium
Prof.Dr. SØREN AABO
Division for Risk Assessment and Nutrition
Research Group for Microbiology and Hygiene
National Food Institute, Technical University of Denmark, Kgs. Lyngby, Denmark
Prof. Dr. KURT HOUF (Supervisor)
Department of Veterinary Public Health and Food Safety & Laboratory of Microbiology
Faculty of Sciences & Veterinary Medicine, Ghent Unviersity, Ghent, Belgium
Prof. Dr. KOEN SABBE (Supervisor)
Protistology and Aquatic Ecology
Faculty of Sciences, Ghent University, Ghent, Belgium
Prof. Dr. METTE BURMØLLE (Supervisor)
Section of Microbiology,
Department of Biology, University of Copenhagen, Copenhagen, Denmark
Prof. Dr. SØREN JOHANNES SØRENSEN (Supervisor)
Section of Microbiology,
Department of Biology, University of Copenhagen, Copenhagen, Denmark
Acknowledgements
This thesis is the result of my work as a joint Ph.D. at the Section for Microbiology, Institute of
Biology, Faculty of Science, University of Copenhagen (KU) and Faculty of Veterinary medicine,
Department of veterinary public health, Ghent University (UGhent). The research work was supported
by the Faculty Scholarships, Copenhagen University and BOF Special Research Fund, Belgium.
The collaborative work would not exist, if not for the many people across different countries, who
have supported me immensely. First and foremost, I owe my sincere thank you to Prof. Dr. Søren J.
Sørensen and Prof.Dr. Kurt Houf for accepting me in their labs; where I was allowed to develop
myself at various aspects of research and as a professional. I thank Prof. Sørensen for introducing me
to my other promoter Prof. Houf. Post this meeting in Copenahgen, we were able to initiate a
wonderful collaboration that paved way to begin my PhD study. I also owe my gratitude to my co-
supervisors Prof. Dr. Mette Burmølle and Prof. Dr. Koen Sabbe for their timely help, support and
guidance at all times. I thank all of them for offering a workplace where I stayed motivated, for
inspiring me with many ideas and providing me support at times of difficulties. Secondly, I thank
Anette Løth and Karin Vestberg for their excellent technical assistance during my time at the section
for Microbiology, KU. I thank Margo Cnockaert, Bart Hoste and Liesbeth Lebbe for making it so easy
for me to get accustomed to the new lab, when I joined and began my work at Laboratory of
Microbiology, UGhent. Further, I would like to thank Prof. Dr. Nina Gunde-Cimerman and Neja
Zupančič of Ljubljana University for extending their collaboration and for the opportunity to co-
author publications. I thank the department secretaries Trine Madsen, Tim Evison (Section for
Microbiology, KU) and Wendy Lievre (UGhent) for their assistance in official paper works. To my
colleagues both in UGhent and KU, I thank you for your valuable discussions, for your help at hand
and making the work environment incredibly enjoyable.
Finally, I thank all my friends, my brother Prashanth and my parents in India and my heartfelt thanks
to Hanna, Jonathan and my parents- in-law, in Germany, for their invaluable love and support. And
most importantly, a big thank you to my wife, Maren Kühlmann for her love, understanding, care,
advice and her immeasurable support during my tenure as a PhD student and in caring for our twin
daughters, Asta and Svala.
Prem Krishnan Raghupathi, Ghent 2018.
Preface
Interactions between bacteria belonging to different species are vital for the development of complex
microbial communities, including multispecies biofilm. Multispecies biofilms are ubiquitous in most
natural and man-made environments; their presence is now subject to ever-increasing attention.
Several studies have shown that bacterial species living in complex bacterial communities interact,
both intra- and interpecifically, and that these interactions are instrumental in structural establishment
and distribution of bacterial species within multispecies biofilm. These complex interactions often
result in the bacteria developing properties that would not been present when grown alone. These
emergent properties include increased tolerance to antibiotics, host immune responses, and other
stressors, which has proven to provide increased fitness benefits to members of the mixed community.
Co-cultivation studies using in-vitro multispecies settings have shown that bacteria in mixed
communities produce increased biomass, and many studies have documented the formation of
microbial aggregates, microcolonies or biofilm formation in response to the presence of predatory
protozoa. The threat from bactericidal protozoans can affect the physiological state of the bacterial
community and result in bacterial responses at both species and social levels, which is in turn
influenced by the combination of different interactions and parameters.
The purpose of this Ph.D. thesis was to address various aspects of bacterial interactions, all of which
support multispecies biofilm formation, and to investigate the role of biofilms as protective
mechanisms when grazing is widespread. More specifically, the microbial diversity of multispecies
biofilm and selected eukaryotic organisms (protozoa and fungi, reespectively) associated with
toothbrushes (manuscript 1) and dishwashers (manuscripts 2 and 3) were investigated. Multiple
bacterial communities isolated from dishwashers were screened for their ability to produce biofilm;
both individually and in co-cultures. The influence of bacterial interactions on population dynamics in
a model culture with four different bacterial strains exposed to grazing (manuscript 4) was also
studied.
The results presented in this thesis shows that studies conducted under the conditions of multiple
species, even though they are less complex than naturally occurring bacterial communities, allow us
to characterize biofilms representing their natural environments where they most often exist as
multispecies microbial communities. The resulting emerging properties such as increased biomass
production and fitness benefits (protection against grazing) associated within the biofilm architecture,
substantiate the presence of synergistic interactions in multispecies biofilm and further emphasize
their influence on individual bacterial species during biofilm formation.
Table of Contents
Introduction ............................................................................................................................................. 1
1. Insights into Free living protozoa and bacterial interactions .............................................................. 1
1.1 Free living protozoa (FLP) ............................................................................................................ 1
1.2 Occurrence of FLP ........................................................................................................................ 2
1.3 Ecological role of protozoa predation –Bacterivory ..................................................................... 3
1.4 Predator – prey interactions .......................................................................................................... 3
2. Significance of microbial biofilm formation and its properties .......................................................... 5
2.1 Bacterial Biofilms – Life attached to surfaces .............................................................................. 5
2.1.1. The EPS Matrix ......................................................................................................................... 6
2.1.2. Bacterial Communication and Genetic exchanges .................................................................... 7
2.1.3. Phenotypic diversity and adaptive capacity .............................................................................. 8
2.1.4. Increased tolerance to antimicrobials ........................................................................................ 8
2.2 Significance of biofilms at homes and industries ......................................................................... 9
3. Understanding multispecies biofilms .................................................................................................. 9
3.1 Multispecies biofilms – an introduction ........................................................................................ 9
3.2 Microbial Co-aggregation ........................................................................................................... 11
3.3 Spatial organization in multispecies biofilms ............................................................................. 13
3.4 Interactions in multispecies biofilm ............................................................................................ 13
3.5 Synergism and exploring synergistic interactions....................................................................... 14
4. Current understanding on FLP and biofilm interactions ................................................................... 16
4.1 Protozoa and biofilms ................................................................................................................. 16
4.2 Protozoan communities in biofilms ............................................................................................ 17
4.3 Biofilms as response against predation ....................................................................................... 17
4.4 Evolutionary aspects of predation ............................................................................................... 18
Aims ...................................................................................................................................................... 19
Summary of Research ........................................................................................................................... 21
Samenvatting (Summary in Dutch) ....................................................................................................... 24
Resumé (Summary in Danish) .............................................................................................................. 27
General Discussion ............................................................................................................................... 28
Future Perspectives ............................................................................................................................... 34
Bibliography ......................................................................................................................................... 35
MANUSCRIPT 1 .............................................................................................................................. 50
MANUSCRIPT 2 .............................................................................................................................. 73
MANUSCRIPT 3 ............................................................................................................................ 106
MANUSCRIPT 4 ............................................................................................................................ 142
CV and List of publications ................................................................................................................ 167
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Introduction
Microorganisms usually live in dense and diverse communities and their arrangement into biofilm
structures and the interactions within these biofilms are vital to the productivity, stability and
resilience of the community. Microbial biofilms form prevalent communities in many environments,
and are characterized by their resistance to pollutants, desiccation, antimicrobial agents and predation.
Despite their widespread occurrence, there is a growing need to understand and characterize complex
microbial communities and the benefits for microbes living in biofilms. In this thesis, we therefore
determined inter-bacterial interactions that favored biofilm formation and its associated fitness
benefits under grazing pressure. The following sections present an overview of various types of
interactions within bacterial communities during biofilm development as well as predator-prey
relationships between protozoa and bacteria.
1. Insights into Free living protozoa and bacterial interactions
1.1 Free living protozoa (FLP)
Free living protozoa (FLP) are heterotrophic unicellular eukaryotes (Adl et al., 2005), found in a
variety of habitats (Ekelund & Rønn, 1994; Hahn & Hofle, 2001). Most FLP are solitary, but some
species forms colonies (Adl et al., 2005; Patterson & Hedley, 1992). These multicellular consortia
however are never differentiated into tissues and therefore remain fundamentally unicellular (Adl et
al., 2005; Vaerewijck et al., 2014). FLP are either motile (e.g., swimming, crawling or gliding) or live
attached to surfaces (Vaerewijck et al., 2014). They are characterized by their cell shape which forms
the basis of morphological identification (Smirnov & Brown, 2004). They are classified into three
groups namely amoebae, ciliates and flagellates. Amoebae develop pseudopodia, ciliates are
characterized by numerous cilia and flagellates possess one or more flagella; used for locomotion
and/or to acquire food (Fig. 1). FLP feed on algae, fungi or other protozoa and are widely considered
as important predators of bacteria (Matz & Kjelleberg, 2005; Parry, 2004). They also obtain nutrients
by absorption of organic material or molecules and can grow on detritus (Scherwass et al., 2005;
Veira, 1986). Many FLP are found in two life stages namely the trophozoite stage, where the cells
feed and multiply, and a cyst stage, where the cells are dormant or resting (Aguilar-Díaz et al., 2011).
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Figure 1: Morphotypes of free living protozoa; A: Amoeba, B: Flagellate and C: Ciliate (Adapted
from http://pinkava.asu.edu/starcentral/microscope/. Drawings made by Stuart Hedley and David
Patterson, licensed to Marine Biology Laboratory (micro*scope).
1.2 Occurrence of FLP
FLP are found in various environments and are abundant in soil and water (Hoorman, 2011). In
addition to their presence in these habitats, FLP have also been reported to be present in air (
Rodriguez-Zaragoza,1994)(Kingston & Warhurst, 1969), geothermal hot springs (Aguilera et al.,
2010; Badirzadeh et al., 2011; Lekkla et al., 2005), caves (Bastian et al., 2009) and in arid (Robinson
et al., 2002) and colder regions (Brown et al., 1982). In relation to anthropogenic environments, FLP
have been detected in hospital and dental water systems (Hikal et al., 2015; Muchesa et al., 2015), air
conditioning units and humidifiers (Schuster, 2002), cooling towers (Canals et al., 2015), swimming
pools (Bonadonna et al., 2004), household kitchens (Chavatte et al., 2014) and refrigerators
(Vaerewijck et al., 2010). Their presence under a wide range of environmental conditions further
confirms that FLP can be highly tolerant towards abiotic factors such as salinity, temperature and
oxygen enabling them to inhabit natural and extreme habitats (Arndt et al., 2000).
Food related environments like commercial broiler houses (Baré et al., 2009) and meat cutting plants
(Vaerewijck et al., 2008) were also found to harbor FLP. They have been detected on vegetables
including carrots, cauliflower, radishes, mushrooms, tomatoes (Rude et al., 1984; Sharma et al., 2004;
Shukla & Sharma, 2011) and numbers of protozoa were reported to be high on leafy vegetables
(Gourabathini et al., 2008; Vaerewijck et al., 2011). Also, investigations on protozoan occurrence on
different sprout types, tap water and bottled mineral water revealed the presence of FLP (Chavatte et
al., 2016; Hoffmann & Michel, 2001; Maschio et al., 2015). Additionally, FLP are naturally present
in the normal microbial gut flora of various animals and insects (Newbold et al., 2015; Tokuda et al.,
2014). In humans, FLP have been detected in nasal, gut and stool samples and can also be found at
high prevalence in healthy populations (Chabé et al., 2017; Corsaro & Venditti, 2015).
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1.3 Ecological role of protozoa predation –Bacterivory
In ecosystems, predators can strongly control numbers and biodiversity of prey in natural habitats.
Predators reduce the abundance of prey, which in turn affects the abundance of predators resulting in
oscillations between numbers of prey and predator (Paisie et al., 2014). However, in the microbial
world, less is known about the effects of predators on microbial communities (Pernthaler, 2005; Sherr
& Sherr, 2002). Predation by FLP is particularly important in the context of the microbial loop (Azam
et al., 1983) where bacterial growth fuelled by dissolved organic materials are consumed by predatory
protozoa. These protozoa are further consumed by meso- and macro-invertebrates and thus transfer
the nutrients throughout the food web for e.g. in soil as shown in Fig. 2 (Fenchel, 1987a).
Figure 2: Trophic interactions in soil (Picture taken from the Soil Foodweb Institute;
https://www.soilfoodweb.com.au/about-our-organisation/benefits-of-a-healthy-soil-food-web).
Protozoan predation consumes 30-100% of bacterial production (Sherr et al., 1983) and is considered
to be a major source of bacterial mortality in various ecosystems such as soil, marine and freshwater
systems (Fenchel & Blackburn, 1999). Predation depends on the type of protozoa present, their
feeding/grazing rates, the physiological state and growth form (free-living or attached) of the prey.
Depending on the type of predator, feeding on bacteria occurs through phagocytosis where the
bacteria are internalized in food vacuoles (phagosomes) and further digested.
1.4 Predator – prey interactions
FLP-bacterial interactions are complex and dynamic, and depend on various conditions including
bacterial species identity and (in the case of pathogenic species) virulence, the type and number of
FLP species present, environmental conditions and abiotic factors such as light, water availability and
temperature. In addition, not all bacteria are susceptible to grazing. Moreover, predator-prey
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interactions are also thought to introduce different traits in bacteria to prevent themselves from being
ingested (Matz & Kjelleberg, 2005). The fate of an ingested bacterium within the protozoa was
suggested to having three possible outcomes by Barker & Brown, 1994:
1) Bacteria survive without multiplication
2) Bacteria multiply without lysis of the protozoan cell
3) Bacteria multiply and cause lysis of the protozoan cell.
During the last decades, adaptations to prevent ingestion have been reported in many studies. Such
pre-ingestional adaptations include alterations in cell size and morphology (e.g. elongated filamentous
cells) (Hahn et al., 1999), increased bacterial motility (Matz & Jürgens, 2005), modification of cell
surface molecules (Wildschutte et al., 2004), microcolony formation and secretion of toxins (Erken et
al., 2011; Matz et al., 2004). However, once the bacteria were ingested by grazing protozoa, various
post-ingestional adaptations have shown to favor prey fitness by resisting predation. These include:
a) FLP serve as a reservoir for various pathogenic bacteria, e.g. Salmonella and Campylobacter (Baré
et al., 2009; Hadas et al., 2004) by bacterial resistance to protozoan digestion.
b) The internalized bacteria are released back into the environments and/or carried to new
environments where FLP could serve as a vector or Trojan horse of bacteria (Barker & Brown, 1994).
c) FLP serve as gene melting pots enhancing lateral gene transfer between bacteria and host, resulting
in increased adaptation to intracellular life within the host. E.g. in Legionella pneumophila, several
genes encode eukaryotic-like proteins that are likely to be interfering with eukaryotic cellular
functions, enabling the bacterium to invade eukaryotic hosts (Lurie-Weinberger et al., 2010). Studies
have also reported that FLP can influence the transfer of antibiotic resistance plasmids between
bacteria (Oguri et al., 2011).
d) FLP also serves to protect internalized bacteria against antimicrobial agents. Examples include
ingested Campylobacter, Salmonella and Yersinia strains showed increased tolerance against
disinfectants while free living bacterial strains were more susceptible (King et al., 1988; Snelling et
al., 2005). Pathogenic bacteria like Listeria monocytogenes, Mycobacterium avium and L.
pneumophila were found to be resistant against antibiotics when internalized within FLP (Barker et
al., 1995; Miltner & Bermudez, 2000; Raghu Nadhanan & Thomas, 2014).
In addition, FLP-bacterial interactions were shown to be influenced by the predator’s feeding
preferences. Amoeba grazing appeared to be non-size selective, whereas flagellates and ciliates
preferred medium to small sized bacterial cells (Hahn & Hofle, 2001; Jürgens & Güde, 1994).
However, this could be due to the fact that they take up prey at specific locations on the cell surface,
which constrains the size of the prey. It was also reported that protozoa preferred to graze on gram
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negative cells due to the lower digestion efficiency when feeding on gram positive cells (Iriberri et al.,
1994; Rønn et al., 2002a). However, this is not always the case, as Gram positive bacteria were found
to be consumed upon (Weekers et al., 1993) and in absence of any choice, FLP have been observed to
consume any available bacteria (Khan et al., 2014). Furthermore, though production of toxins was
shown to negatively influence predation, production of metabolites by protozoa was also reported to
positively influence prey survival (Laskowski-Arce & Orth, 2008), underlining the existence of
complex interactions between these organisms.
In parallel to the above mentioned studies focused on the many ways in which individual cells avoid
predation, investigations on prey-predator relationships tend to be more complex when involving
bacterial aggregates and biofilms. Currently, there is little information on prey-predator interactions
within microbial biofilms. The following sections address the importance and benefits of biofilms,
interactions within biofilms, multispecies settings and current insights into the role of protozoa in
biofilms.
2. Significance of microbial biofilm formation and its properties
2.1 Bacterial Biofilms – Life attached to surfaces
Bacteria can present distinct lifestyles during their growth, either in planktonic form or as biofilm
(Hernandez-Jimenez et al., 2013). Planktonic bacteria are free-floating in suspension. Biofilms are
assemblages of microbial cells clumped together as multicellular aggregates (Kragh et al., 2016),
enclosed within a matrix of extracellular polymeric substances (Donlan, 2002), or attached to solid
surfaces, soft tissues of living organisms or at the liquid-air interface (Jain et al., 2007). Biofilm
growth can be divided into a series of different stages often depicted in a biofilm life cycle (Stoodley
et al., 2002). When individual cells come in contact with a surface, they attach themselves by
extracellular pili or through secretion factors (Klausen et al., 2003; Latasa et al., 2005). After surface
attachment, these bacterial cells divide and grow to form micro-colonies, leading to a succession to a
structurally mature biofilm. Afterwards, under right conditions, the mature biofilm can experiences a
dispersal phase to colonize new niches, thus repeating the above stages of biofilm development. After
attachment and maturation, a bacterium detaches from the surface and returns to a planktonic lifestyle
(Fig. 3) (Stoodley et al., 2002), or shifts to a more permanent bonding, leading to an irreversible
attachment, also influenced by the surface properties (Pasmore et al., 2002; Yoda et al., 2014).
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Figure 3: Stages in biofilm formation (Adapted from Chung & Toh, 2014).
Compared to planktonic cells, biofilm formation is now increasingly considered as a survival strategy
for bacterial cells, as it offers a number of advantages, listed below.
2.1.1. The EPS Matrix
The microorganisms in biofilms live within a self-produced matrix of extracellular polymeric
substances (EPS) that offers structural stability to biofilms. EPS constitute up to 90% of dry weight in
most biofilms (Sutherland, 2001) and are mainly composed of water, exopolysaccharides, proteins,
nucleic acids, lipids and extracellular DNA. EPS provide the three-dimensional architecture (dense
areas, pores and water channels) in biofilms and are responsible for biofilm adhesion to a surface and
cohesion during the biofilm development.
Polysaccharides are the major fraction of the EPS matrix (Frølund et al., 1996; Wingender et al.,
2001), consisting of different homopolysaccharides (sucrose-derived glucans and fructans; cellulose)
and heteropolysaccharides (alginate, xanthan). Wide ranges of species, from a variety of
environments, have been reported to produce exopolysaccharides (Sutherland, 2007), and the
production is also variable between strains of a single species, for e.g. the production of alginate and
two polysaccharide-encoding genes, pel (pellicle) and psl (polysaccharide synthesis locus) by
different Pseudomonas aeruginosa strains (Ryder et al., 2007).
Proteins are present in considerable amounts within the matrix and are mainly involved in the
degradation of polymeric substances (Kaplan, 2014). During biofilm development or during
starvation, many enzymes degrade complex EPS components to simple low molecular mass products,
which can be taken up and utilized as carbon and energy sources (Tielen et al., 2013). In addition,
structural proteins were reported to be involved in the formation and stabilization of the matrix
network. Examples includes the expression of peptidase M7 (TasA) required for the structural
integrity of Bacillus subtilis during biofilm formation (Romero et al., 2014) and biofilm-associated
protein (Bap) in Salmonella enterica serovar Enteritidis (Latasa et al., 2005) provide structural
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integrity and promote biofilm formation. Proteinaceous appendages like Type IV pili, fimbriae and
flagella also act as structural elements by interacting with other EPS components of the biofilm matrix
(van Schaik et al., 2005; Zogaj et al., 2001).
Extracellular DNA (eDNA) serves as a structural component in biofilms. eDNA forms grid-like
structures in P. aeruginosa, filamentous networks in Rheinheimera texasensis and as a dense network
of fine strands in Haemophilus influenzae biofilms (Allesen-Holm et al., 2006; Bockelmann et al.,
2006; Jurcisek & Bakaletz, 2007; Merchant et al., 2007). Lipids found in the matrix are crucial for
adherence and have been proposed to act during initial microcolony formation, facilitating surface-
associated bacterial migration, and playing a role during biofilm dispersion events (Boles et al., 2005;
Conrad et al., 2003). Water within the matrix provides a hydrated environment. Water buffers the
biofilm cells against fluctuations in water potential, during rapid wetting or drying events, thus
protecting the biofilm-embedded bacteria (Flemming & Wingender, 2010; Or et al., 2007). Also, an
higher proportion of EPS matrix and its water retention capacities within biofilms have shown to
confer tolerance against desiccation (Flemming & Wingender, 2010; Potts, 1994; Roberson &
Firestone, 1992).
2.1.2. Bacterial Communication and Genetic exchanges
The close proximity of cells within the EPS matrix allows for intense interactions, enabling the
bacteria to communicate with each other using chemical signals or by transferring genetic material.
Quorum sensing (QS) is an important phenomenon that occurs via chemical signals, helping cells to
communicate and maintain cell-population density and regulate gene expression (Miller & Bassler,
2001). Cell-to-cell signaling in P. aeruginosa and its role in biofilm formation was reported by
Davies et al., 1998. The close proximity of cells, spatio-chemical conditions and the compound-
retaining matrix provide optimal conditions for QS-mediated gene regulation. Thus, QS is an integral
component of bacterial global gene regulatory networks responsible for bacterial communication (von
Bodman et al., 2008) that could play a role during biofilm development. Genetic exchanges through
horizontal gene transfer (HGT) between biofilm residents have major consequences for the
physiology of biofilms, as well as evolutionary outcomes (Madsen et al., 2012; Molin & Tolker-
Nielsen, 2003). It is now known that microbes can take up external genetic material such as
transposons, plasmids and viruses through transduction or transformation (Frost et al., 2005; Sørensen
et al., 2005). Various conjugative plasmids for encoding adhesive structures (like fimbriae) were
characterized, and their presence induced biofilm formation (Burmølle et al., 2008; Reisner et al.,
2006). The large amounts of eDNA in biofilms also facilitate the transfer of genetic elements to
members of the biofilm community (Das et al., 2013).
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2.1.3. Phenotypic diversity and adaptive capacity
Sessile bacteria in biofilms exhibit a phenotype that differs from bacteria grown in suspensions
(planktonic), sometimes expressing genes that are never expressed in their planktonic life stage.
Biofilms represent heterogeneous aggregations of microbial phenotypes and genetic variants (Chia et
al., 2008). Examples include P. aeruginosa variants that hyper-adhere to solid surfaces and which
showed an increased expression of the psl and pel loci (Kirisits et al., 2005). In Serratia marcescens
biofilms, variants with sticky colony textures produced biofilm biomass two- to threefold more than
the wild type (Koh et al., 2007) and mature biofilm of Flavobacterium psychrophilum produced
phenotypes with altered global transcriptional activity that differed significantly from cells in
suspension (Levipan & Avendaño-Herrera, 2017). Phenotypic diversity contributes to phase variation,
experienced under different environmental and nutritional conditions, affecting the biofilm
development (Chia et al., 2008). Under favorable conditions, biofilms maintain a balance between
growth and maintenance of its structure. When faced with unfavorable conditions, biofilms regress to
earlier stages, while maintaining their surface adhesion, and develop again when conditions improve.
Thus, altered phenotypes enhance the adaptive capacity of biofilm members by altering cellular
processes, which in turn, lead to the overall success of the biofilm community.
2.1.4. Increased tolerance to antimicrobials
Bacteria residing in biofilms have enhanced tolerance to antimicrobial agents and antibiotics
compared to free-living cells (Luppens et al., 2002). In biofilms, this enhanced tolerance is multi-
factorial, through combinations of different mechanisms such as restricted penetration of antibiotics
though the EPS matrix (Anderl et al., 2000; Campanac et al., 2002; Drenkard, 2003); acquisition of
random mutations or plasmid uptake (Donlan & Costerton, 2002; Drenkard, 2003; Gilbert et al.,
2002; Lewis, 2001; Stewart & Costerton, 2001); reduced metabolic rates, altered micro-environments,
slow growth rates of biofilm cells (Anderl et al., 2003; Gilbert et al., 1990; Leid et al., 2002) and
formation of dormant persister cells that are not killed by exposure to antibiotics (Keren et al., 2004;
Shapiro et al., 2011). Recent evidence suggests contributions of biofilm physiology to tolerance, as
biofilm bacteria were reported to express specific protective factors such as multidrug efflux pumps
and stress response regulons (Allegrucci et al., 2006; Amaral et al., 2014; Liao & Sauer, 2012; Mah et
al., 2003; Stewart & Costerton, 2001). In addition to biofilm mediated enhanced protection reported
for antimicrobial agents (Fux et al., 2005), biofilms also offer protection against various chemical
stresses generated by diverse environment and metabolic processes (Cappitelli et al., 2014; Gambino
& Cappitelli, 2016).
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2.2 Significance of biofilms at homes and industries
Considering the substantial physiological advantages of surface colonization and biofilm formation, it
is not surprising that the surface associated lifestyle plays a vital role in microbial adaptation to
different environments (Dang & Lovell, 2016), and 99% of bacteria found in nature can live attached
to surfaces (Dalton & March, 1998). In this perspective, numerous efforts are underway to
characterize the microbial diversity in home environments (Flores et al., 2013; Hamada & Abe, 2009)
and various man-made ecosystems such as trash bins (Naegele et al., 2015), tap water pipes (Ren et
al., 2015b), automated teller machines (Bik et al., 2016), coffee-machines (Vilanova et al., 2015),
washing machines (Callewaert et al., 2015) and dishwashers (Zalar et al., 2011; Zupančič et al.,
2016), as problems including food spoilage and the spread of infectious diseases continue to persist.
Data on the incidence of infectious disease caused by emerging and re-emerging pathogens has led to
initiatives to improve home hygiene (Gorman et al., 2002), for example, the removal of bacteria from
domestic surfaces. Also, a survey in England showed that approximately 16% of food poisoning
outbreaks were associated with meals prepared in private houses (Mattick et al., 2003). Studies have
confirmed the ubiquity of biofilms in household surfaces (Rayner et al., 2004) and transfer of bacteria
from biofilms formed on food ingredients, food handlers or food contact surfaces (Pérez-Rodríguez et
al., 2008) could contribute to a certain fraction of food related outbreaks.
In food industries, biofilms cause serious problems with microbial clogging and its effects result in
impeding the flow of heat across surfaces, increases in frictional resistance and increased corrosion
rates of surfaces (Verran, 2002). Pathogenic microbiota growing on food surfaces or associated with
biofilms in processing environments could cross-contaminate and cause post-processing
contamination (Kumar & Anand, 1998; Wingender & Flemming, 2011). If these biofilm-associated
microorganisms from food-contact surfaces are not completely removed, they can lead to mature
biofilm formation and increase cross-contamination risks (Garrett et al., 2008). The genetic,
physiological and ecological processes of, and the mechanisms and functions associated with the
surface-associated microbiota, make their study fascinating. Increasing attention is now being given to
the area of multispecies biofilms in order to understand their significance in microbiology.
3. Understanding multispecies biofilms
3.1 Multispecies biofilms – an introduction
Biofilm formation is an integral part of the bacterial lifestyle and involves the formation of complex
communities with other microbial species (Fig. 4) (Hall-Stoodley et al., 2004). Bacteria, algae, fungi
and protozoa are all common inhabitants of multispecies biofilms in natural environments (Burmølle
et al., 2011). One proposed model in the development of multispecies biofilms proceeds with
successions of adhesion and multiplication events. The primary (early) colonizers initiate the
[INTRODUCTION]
10
attachment of individual cells on to a surface, mediated through specific or non-specific physico-
chemical interactions. Under suitable conditions, the primary colonizers multiply on the surface to
form micro-colonies. As conditions change within the young biofilm, secondary (late) colonizers are
also able to attach to the primary colonizers and the biofilm begins to develop into a multi-species
community (Rickard et al., 2003).
Figure 4: Development of mixed biofilm with different bacterial species (Picture taken from
http://www.cresa.cat/blogs/sociedad/en/espanol-biofilms-bacterianos-por-que-deberia-importarnos/).
The defined structure within biofilms allows different microbes to adhere to surfaces and presents a
network of complex interactions between different members that we have only begun to understand.
Interspecies dynamics within mixed biofilms such as communication and/or competition for nutrients
and physical resources represent those of a community. This constitutes a layer of complexity when
addressing mixed-species biofilms (Elias & Banin, 2012). However, while the above-mentioned
biofilm properties were put forward using mono-species conditions, similar emergent properties were
also reported in multispecies biofilm communities.
Extracellular polymeric substances (EPS) serves as a biological ‘glue’ enabling different microbes to
adhere to, colonize and establish on a wide range of surfaces (Limoli et al., 2015). As the composition
of the EPS matrix varies greatly depending on the bacterial species and environmental conditions
(Flemming & Wingender, 2010), the composition of EPS is different in mono-and multi-species
biofilms. The matrix thus represents a complex mixture resulting from the fusion of different EPS
produced by many members of the microbial community (Decho et al., 2005). This further adds
complexity when characterization of EPS in mixed species biofilms is to be performed. However,
there is an agreed consensus that in multispecies biofilms, the EPS matrix serves an interspecies
public good (Sanchez-Vizuete et al., 2015). EPS confers protection against biocides by limiting
diffusion rates (Andersson et al., 2011) or using matrix associated enzymes produced by one species
[INTRODUCTION]
11
that benefit the whole population, for e.g., the production of hydrolase by P. aeruginosa and cross-
seeding of functional bacterial amyloids aid in multispecies biofilm development (Lee et al., 2014;
Zhou et al., 2012). In addition, BslA amphiphilic protein from B. subtilis have been shown to prevent
penetration of biocides, thus protecting other inhabitants (Kobayashi & Iwano, 2012).
Multispecies biofilms are likely to provide cell-to-cell contact and HGT events (Stalder & Top, 2016).
Studies have shown that quorum sensing signals such as autoinducer -2 (AI-2), autoinducer peptides
and N-acylhomoserine lactone (AHL) promoted mixed-species biofilm formation in different
bacterial combinations, e.g. in Porphyromonas gingivalis and Streptococcus gordonii; in Actinomyces
naeslundii and Streptococcus oralis; and in Pseudomonas aeruginosa and Burkholderia cepacia
(McNab et al., 2003; Rickard et al., 2008; Riedel et al., 2001). Multispecies biofilms are associated
with increased matrix production and the emergence of genetic variants within specific species under
mixed conditions (Sanchez-Vizuete et al., 2015). A study on dual-species biofilms reported the
occurrence of different phenotypic variants of Pseudomonas putida contributing to a more stable and
productive community in the presence of Acinetobacter sp. (Hansen et al., 2007).
It is also important to address the presence of microbial members belonging to other kingdoms like
fungi in multispecies biofilm settings. Fungal-bacterial biofilms differ from bacterial or fungal
biofilms as fungi can already present a surface to which bacteria can adhere to (Seneviratne et al.,
2007). Studies on fungal-bacterial biofilms have also revealed better growth and colonization abilities
to influence each other compared to their monospecies counterparts (Boer et al., 2005; Elvers et al.,
1998). In any case, the general characteristics within multispecies settings point to the existence of
various microbial interactions. Different multispecies biofilm model systems combined with a variety
of analytical techniques are being developed (Rendueles & Ghigo, 2012). This has led to an increased
understanding of their complex relationships, functions, dynamics and interactions within these mixed
communities.
3.2 Microbial Co-aggregation
Co-aggregation mechanisms facilitated by cross-species protection, cross-feeding and co-metabolism
have been observed in multispecies biofilms. These in addition confer spatial and functional
heterogeneity in multispecies biofilms (Pande et al., 2016). Co-aggregation is a process by which
genetically distinct bacteria become attached to one another through specific cell to cell recognition
systems. This phenomenon is well studied in the case of oral biofilms (Kolenbrander et al., 1985;
Marsh, 2006). Co-aggregating species may act to serve as a bridge between different bacterial species,
e.g. Fusobacterium nucleatum bridging the gap between Treponema denticola and Porphyromonas
gingivalis in the presence of major outer sheath protein (MSP) during cooggregation process was
demonstrated (Socransky et al., 1998; Rosen et al., 2008). Other examples include Streptococci
colonizing the tooth surface together with Gram-positive rods such as Actinomyces naeslundii and
[INTRODUCTION]
12
Porphyromonas gingivalis co-aggregated with the primary colonizer Streptococcus gordonii (Lamont
et al., 2002) in dental plaque biofilms where streptococcal surface proteins (Ssp) were involved
during biofilm development. Tight co-aggregation between Candida albicans and streptococci has
been observed, owing to specific adhesins similar to those found in S. gordonii (Metwalli et al., 2013;
Silverman et al., 2010). Co-aggregating mixtures also resulted in spatial arrangement of biofilm
structure for e.g., in dental plaque biofilms (Mark Welch et al., 2016; Valm et al., 2011) and in flow
cells where Escherichia coli colonies occurred along the outer edge of the flow cell with the partner
P. aeruginosa accumulating in the centre (Klayman et al., 2009). Co-aggregation between lactobacilli
and E. coli strains isolated from various environments mediated by the production of a 32 kDa
proteinaceous aggregation‐promoting factor by Lactobacillus reuteri was previously reported (Kmet
et al., 1995; Reid et al., 1988). Hence, co-aggregation is essential for the orchestrated development of
multi-species biofilms (Katharios-Lanwermeyer et al., 2014).
Co-aggregation between microbial members enhances cross-species protection. Co-aggregation in
biofilms was observed to protect anaerobes from oxygen and susceptible species from antimicrobials
(Bradshaw et al., 1998; Gilbert et al., 2002). This was also demonstrated in bacterial-fungal biofilms
where Staphylococcus epidermis RP62A and Candida albicans appeared to protect each other against
antimicrobials thorough EPS production (Adam et al., 2002). Microbial aggregation favors co-
metabolism where one species utilizes the metabolite produced by a neighboring species. A classical
example includes a mixed community composed of Methanobacilus omelianskii and
Methanobacterium strain MOH, in which M. omelianskii oxidizes ethanol to acetate with the
liberation of hydrogen, while MOH can reduce CO2 to methane using the hydrogen released by M.
omelianskii under anaerobic conditions. However, M. omelianskii is not able to grow on ethanol when
alone while the cooperative interaction between these two microbes allowed lavish growth of the both
(Bryant et al., 1967). Similarly, metabolic cooperation occurred within Burkholderia- Pseudomonas
biofilms where Burkholderia xenovorans LB400 grown on medium with chlorophenyl, they excreted
chlorobenzoate that was subsequently metabolized by Pseudomonas sp. B13 (Nielsen et al., 2000).
Co-metabolic degradation of polycyclic aromatic hydrocarbons was enhanced in the presence of
fungal-bacterial co-cultures and accelerated remediation of chemical contaminated water systems was
suggested due to an increased co-metabolism when multiple species were present (Boonchan et al.,
2000; Farabegoli et al., 2008). Mixed-species biofilms composed of Variovarax sp., Comamonas
testosteroni and Hyphomicrobium sulfonivorans degraded linuron and its metabolic intermediate more
efficiently compared to single-species biofilms (Breugelmans et al., 2008).
The biofilm mode of life allows microbes to interact with each other and function as a group for
coordinated activities (Nadell et al., 2009). Metabolic cross-feeding interactions are ubiquitous in
multispecies biofilms and have been reported to play a crucial role for the entire microbial community
(Embree et al., 2015; Møller et al., 1998; Zelezniak et al., 2015). Metabolic exchanges form a
[INTRODUCTION]
13
strategy for group success (Morris et al., 2013; Ponomarova & Patil, 2015; Ren et al., 2015a), where
metabolic interactions facilitate, through division of labor, emergent properties at the community
level, such as biodegradation (Fowler et al., 2014; Lykidis et al., 2011), increased virulence (Alteri et
al., 2015; McNally et al., 2014) or enhanced biofilm biomass (Palmer et al., 2001).
3.3 Spatial organization in multispecies biofilms
Bacteria in biofilms generate a complex three dimensional structure and interactions within result in
physicochemical gradients and numerous microenvironments (Mark Welch et al., 2016). Two
scenarios of spatial structuring in biofilms were suggested (Nadell et al., 2016). First, cells at low
densities are predominantly solitary in their early phase of biofilm growth. In the second and later
stages of biofilm development, cells at higher densities contain segregated lineages or mixed lineages
(Fig. 5) (Nadell et al., 2016). The spatial arrangement of different strains and species within biofilms
strongly influences the relative fitness benefits of cooperative and competitive phenotypes, thus
influencing their overall community function (Inglis et al., 2009; Rumbaugh et al., 2009). Hence to
understand microbial biofilm communities, we need to understand the balance of cooperation and
competition within biofilm.
Figure 5: Cells appear as solitary cells. Later with biofilm development, segregated lineages indicate
a scenario where cooperative public goods are favored, and cells often present as clonal clusters. In
biofilms with mixed lineages, the interactions were expected to be predominantly antagonistic,
although inter-strain commensalism or mutualism could also be favored (Adapted from Nadell et al.,
2016).
3.4 Interactions in multispecies biofilm
In multispecies biofilms, microorganisms compete or cooperate with each other in interaction with the
external environment. This critically influences the structure, function and development of the biofilm
(Burmølle et al., 2014; Yang et al., 2011). Microbial biofilm communities interact with each other
[INTRODUCTION]
14
and with neighboring cells altering the community productivity. Interactions are classified
‘antagonistic’ (competition, parasitism, predation) if they negatively affect the community or ‘co-
operative’ (cooperation, synergism, altruism, mutualism) if they positively affect the community
(Ahmad et al., 2017).
Competitive antagonistic interactions between microbes are due to active struggles for nutrient
sources, oxygen and available space (Hibbing et al., 2010). The microbial composition of the biofilm,
the availability of nutrients along with competitive interactions and other growth parameters serve as
important driving forces for determining the structure and development of biofilms (Giaouris et al.,
2015). Investigations have shown how different microorganisms can effectively outcompete others as
a result of better utilization of a given energy source (Wuertz et al., 2004) and through the production
of bacteriocins, organic acids, biosurfactants and enzymes that may inactivate or inhibit the growth
(Tait & Sutherland, 2002), or prevent attachment or provoke the detachment of species from the
biofilm structures (Rendueles & Ghigo, 2012).
Several cooperative interactions occur, where all species benefit from the presence of others, leading
to an enhanced overall fitness (Stewart & Franklin, 2008). In multispecies biofilms, these fitness
benefits include; increase in biofilm formation capacity of the whole community, protection of
community members from antimicrobial action or modification of the local microenvironment
supporting the growth of other organisms (de Beer et al., 1994; D’Urzo et al., 2014; Ramsing et al.,
1993). However, it should be noted that though the overall fitness of the biofilm is enhanced when
multiple species are present, the underlying behavior between different species in some cases could be
competitive (Burmølle et al., 2014). In addition, the general classification of cooperative and
antagonistic interactions are not clearly defined, as they are often based on the social behavior of two
populations (actor and recipient) (West et al., 2006). Natural communities are far more complex
where more than two species are involved. Therefore, it is important to address interactions and their
effect on the whole community as well as on the individual members present within the community.
3.5 Synergism and exploring synergistic interactions
Synergism in multispecies biofilms is when the combined functions of different microbes produce a
collective effect that would be greater than their individual effect (Burmølle et al., 2014). Synergy in
multispecies biofilms is when an overall increase in biofilm biomass, increase in cell numbers of
interacting microbes or enhanced community function is observed. These synergistic interactions have
been suggested to operate in concert and have been demonstrated to strengthen the protective effects
of biofilms when multiple species are present compared to planktonic and monospecies biofilm
communities. This has been verified in multispecies biofilm communities where the whole
community gained protection against disinfectants compared to monospecies biofilm (Burmølle et al.,
2006; Schwering et al., 2013). Another example includes a multispecies biofilm consortium
[INTRODUCTION]
15
composed of P. aeruginosa, Pseudomonas protegens, and Klebsiella pneumonia which was resistant
to tobramycin and sodium dodecyl sulfate (SDS). It was shown that multispecies consortia showed
increased community-level resistance due to cross protection offered by P. protegens to all members
in the community (Lee et al., 2014) and other example include multispecies biofilms formed by
drinking water isolates that were less susceptible to sodium hypochlorite disinfection compared to
monospecies biofilms (Simões et al., 2010) supporting the common belief that multispecies biofilms
are less susceptible than monospecies biofilms. The lowered susceptibility of multispecies biofilm
relative to single species biofilms was reported to depend on higher cell densities, the number of
species incorporated, the role played by each of these species or tolerance displayed by a key species
(Simões et al., 2010, Shakeri et al., 2007).
The prevalence of biofilm induction and synergistic effects in biofilm formation were examined in
various studies by co-culturing different species combinations (Madsen et al., 2016; Ren et al., 2014,
2015a). The effects were classified synergistic in the multispecies biofilms by relating the biomass of
multispecies biofilm to that of the best single species biofilm producer. It is based on the assumptions
that a) the cell densities and biofilm forming capacities under similar nutrient availabilities of mono
and multispecies biofilm are equal unless interactions causing synergistic or antagonistic effects occur
and b) the best biofilm former dominates the multispecies biofilm (Ren et al., 2015a).
Figure 6: Example of a classification scheme to assign ‘synergy’ or ‘no synergy’ to biofilms levels
formed using mixed bacterial co-cultures.
[INTRODUCTION]
16
The above classification scheme illustrates the synergistic regime based on the biofilm forming
capability of each member (A and B in monoculture) and of the co-cultures (black bar) based on two
different genotypes. When the biofilm formed by co-cultures is greater than the best single strain
biofilm producer (Genotype A in Fig. 6), there is a biofilm induction and when their ratio i.e., fold
change, Fd (biofilm levels of co-cultures/ biofilm levels of single strain) is ≥ 1, this denotes
synergistic biofilm induction. The biofilm formed by the co-cultures when less than the best single
strain producer, this denotes ‘no synergy’. Antagonisc reduction is when a lesser biofilm is formed
compared to the poorest biofilm producer (Genotype B). This regime is applicable to co-cultures with
any number of unique genotypes. Also, synergistic interactions inducing multispecies biofilms were
also observed when poor biofilm formers were co-cultured verifying that the biofilm forming ability
of individual species do not necessarily reflect their potential when present in multispecies conditions
(Bharathi et al., 2011; Burmølle et al., 2006). As discussed above, the varying levels of biofilm
formation among multispecies bacterial community can be the result of metabolic interactions (Møller
et al., 1998), enhanced co-aggregation (Rickard et al., 2003), organized spatial distribution (Skillman
et al., 1998) and/or facilitated initial surface attachment i.e. a bridging bacteria facilitates the
attachment of other species that do not co-aggregate directly with each other (Klayman et al., 2009).
Hence, bacterial species that do not form biofilms as single strains could benefit from being
associated to biofilm formers, with expanded niche and protection from external stress, by engaging
in multispecies communities (Ren et al., 2015a). These synergistic interactions can be further
explored to understand its effects on the population dynamics, monitoring gene and protein expression
within multispecies communities using techniques like quantitative polymerase chain reaction (PCR),
fluorescent based in-situ hybridization (FISH) and transcriptomics and proteomic based approaches
(Hansen et al., 2017; Herschend et al., 2017; Liu et al., 2017).
4. Current understanding on FLP and biofilm interactions
4.1 Protozoa and biofilms
The role of protozoa in pelagic and benthic food webs has been receiving increasing attention from
scientists in the last decades (Sherr & Sherr, 2002). Protozoa perform several important functions
within in the food web and being the main bacterial consumers, bacterial communities in soil and
marine ecosystems have been shown to be affected by predatory protozoa (Hahn & Hofle, 2001;
Jürgens & Güde, 1994; Rønn et al., 2002a). Parallel to the investigations of the pelagic microbial food
web, researchers also focus on the importance of microbial interactions and processes at different
interfaces (Arndt et al., 2003). Protozoa-bacteria interactions at solid-air, solid-liquid and liquid-liquid
interfaces occur and free-living protozoa fall into different categories (Table 1) based on their
interaction with interfaces and feeding preferences, according to Parry, 2004.
[INTRODUCTION]
17
Table 1: Different protozoan group characterized by their feeding preferences
Differences in habitat impact protozoa grazing. Though the majority of the interactions between
protozoan grazers and their prey are similar in aquatic and soil habitats, differences exist. The
physical nature of soil constrains the active movement of organisms compared to aquatic systems. In
addition, water-filled pores in soil may also protect bacteria from grazing (Postma & van Veen, 1990;
Wright et al., 1993). In addition, protozoa living at the water boundary layer around soil particles can
survive even when the pore water dried out (Fenchel, 1987b). It was also observed that in flowing
waters, protozoa live in the water boundary layer where the stream velocity is close to zero (Silvester
& Sleigh, 2006). Environmental bacteria frequently confronted with predatory protozoa cause major
bacterial mortality and impose a pronounced effect on prey fitness. These predator-prey interactions
could result in the development of diverse strategies against grazing pressure (Arndt et al., 2003; Matz
& Kjelleberg, 2005).
4.2 Protozoan communities in biofilms
Protozoa feed efficiently on attached bacteria and the grazing efficiency differs strongly between the
different protozoan groups. Protozoan colonization of bacterial biofilms occurs at successive stages;
early surface colonizers like heterotrophic flagellates colonize the surface due to their high mobility
and abundance. This is later followed by ciliates and amoebae (Arndt et al., 2003). Biofilms can be
colonized by various protozoan groups including amoebae, flagellates and ciliates (Parry, 2004).
Although many different protozoan species were found associated with biofilms, the level of their
association with and grazing impact on biofilm-prey differs (Parry, 2004). For e.g., in early stage
biofilms, the predators were generalists that fed on suspended and attached prey, whereas in later
stages/mature biofilms they were composed of specialists that could attach and feed on surface-
associated bacteria (Arndt et al., 2003). Another study showed that different assemblages of soil
protozoa produced varying effects on bacterial community structure that were dependent on the type
of protozoa present (Rønn et al., 2002b).
4.3 Biofilms as response against predation
The gel-like state of the biofilm matrix was conceptualized to limit the access of antibacterial agents,
such as antibodies and phagocytic eukaryotic cells, and in line with this, it was proposed that biofilm
bacteria are substantially protected from amoebae or immune cells, similar to the resistance against
Protozoan group Characteristics
Transient Predominantly free-swimming, feed on suspended prey
Sessile Attached to a surface, feed on suspended prey
Browser Free swimming, feed on suspended and attached prey. They can browse
over surfaces for prey
Amoebae Browse over surfaces and feed on attached prey only
[INTRODUCTION]
18
antibiotics (Costerton et al., 1987). Defense strategies against protozoan predation, initially observed
with respect to change in bacterial morphologies (Hahn et al., 1999; Jürgens & Güde, 1994) and later
as bacterial aggregate formation (Jürgens et al., 2000), paved the way for suggesting that biofilm
formation to serve as a protective niche against grazing (Matz & Kjelleberg, 2005).
According to Matz, 2007, the characteristic features of the biofilm mode of life (surface adherence,
encapsulation by extracellular matrix and subsequent high cellular densities) are effective in
compromising predation. Studies have shown bacterial hydrophobicity and bacterial surface charge to
alter the feeding rates of nanoflagellates. The EPS matrix forms a physical barrier against the
attacker; evidence for such physical defense mechanisms comes from P. aeruginosa and Vibrio
cholerae biofilms co-cultivated with flagellate grazers (Matz et al., 2004, 2005). Grazing induced
large micro-colonies of alginate producing Pseudomonas and enriched the biofilm-forming strains of
V. cholerae, which consequently reduced the grazing efficiency. The indirect effect of the EPS matrix
on anti-predator fitness of biofilm cells was reported in the formation of high cell density consortia.
The close proximity of cells allows bacterial populations to communicate and cooperate via quorum
sensing (QS). QS favoring anti-predatory mechanisms, as was reported in P. aeruginosa biofilms
where QS signals induced the formation of micro-colonies and production of rhamnolipids to resist
protozoan grazing (Matz et al., 2004). Similarly, QS controlled differentiation of Serratia marcescens
cells into filaments and cell chains in biofilms was shown to protect from grazing (Queck et al.,
2006). While many of the above observations were tested under laboratory conditions, similar
protective effects of biofilm formation were also observed under semi-natural conditions. Grazing by
flagellates stimulated the abundance of bacterial micro-colonies within river biofilms (Wey et al.,
2008) and in other study, using activated sludge, showed that protozoa grazing initially reduced the
biofilm development but later stimulated the biofilm growth (Rychert & Neu R, 2010).
4.4 Evolutionary aspects of predation
Predation presents a selective force where adaptations against grazing increase the bacterial fitness
that can be evolutionarily favored over time. Formation of inedible microcolonies and QS mediated
secretion of virulence factors together with the fitness advantage of biofilm growth could in turn lead
to the evolution of multicellular traits and cooperative behavior (Matz, 2007) including
multicellularity and pathogenesis (Matz & Kjelleberg, 2005). The notion also exists that the survival
and successful replication of bacteria within the protozoa niche could act as a driving force in the
evolution of some bacteria as pathogens such as Listeria, Mycobacterium and Legionella (Brown &
Barker, 1999). However, our understanding of predator-prey interactions remains unclear due to its
complexity. Multispecies interactions in biofilms further add to the complexity and studies of biofilm
adaptations to predation may provide insights on how microorganisms persist and diversify in the
environment.
[AIMS]
19
Aims
Biofilms presents as a complex networks of inter-connected organisms and the complexities varies
from single species biofilm populations to species rich biofilms. Biofilm include bacteria, algae,
fungi, protozoa, nematodes and their study has revealed them to be complex and diverse. Research on
microbial interactions within biofilm communities have begun to shed light on different metabolic
functions and how different organisms cooperate.
The general aims of this doctoral research were to assess interactions between multiple species of
bacteria during biofilm development. Synergism among multiple species has been shown to
strengthen the protective effects of biofilms when multiple species were involved during biofilm
development. In this thesis, several bacterial isolates were tested for their ability to form synergistic
multispecies biofilms. Further, we investigated to what degree synergistic interactions can lead to
enhanced overall biomass productivity or affect prey fitness with respect to resistance to grazing. To
this end, mono and multispecies biofilms of four bacterial cultures in the presence and absence of a
pelagic predator were cultivated and analyzed.
The specific goals addressed during this study were:
1. Expanding the current understanding of protozoan ecology. In this study, toothbrush samples were
evaluated for the detection and presence of FLP. This aspect underlies the importance to incorporate
protozoan diversity in microbiological surveys.
2. Deciphering the microbial community composition of biofilms formed on the rubber seals of
household dishwashers. Dishwashers pose a range of growth constraining factors where microbes tend
to establish as biofilms under such conditions. In this study, both fungal and bacterial community
composition of biofilms were elucidated using next generation sequencing. Further, it was shown how
different conditions of the dishwashers affected the microbial biofilm composition.
3. Multispecies biofilm development. The isolates obtained from the dishwashers were tested for in-
vitro biofilm formation both in mono and four-species conditions. Synergistic interactions
contributing to an overall increase in multispecies biofilm formation were identified. Further,
multispecies bacterial communities and their ability to harbor the opportunistic fungal pathogen,
Exophiala dermatitidis within the biofilm were examined. E. dermatitidis were most commonly
detected in indoor environments and isolated at highest frequencies from dishwashers. These black
yeasts showed higher affinity for rubber and hence, their establishments as mixed bacterial- fungal
biofilms on different dishwasher surfaces commonly used in the industry were investigated.
4. Assessing the protective effect of multispecies biofilm under grazing pressure. Free-living
protozoa found in natural environments like soil are considered as important predators of bacteria.
[AIMS]
20
Ciliates exhibit high ingestion rates and at such extreme grazing pressure, biofilms are likely to
present as an alternate survival strategy for bacteria to overcome predation. Towards this, a bacterial
model consortium composed of four different species isolated from soil, namely Xanthomonas
retroflexus, Stenotrophomonas rhizophila, Microbacterium oxydans and Paenibacillus amylolyticus
was subjected to grazing by the ciliate Tetrahymena pyriformis under different mono and multispecies
conditions. Impact of grazing on inter- and intra-species interactions and population dynamics of this
bacterial consortium were further investigated.
[SUMMARY IN ENGLISH]
21
Summary of Research
Interspecies interactions are vital for the development of any complex communities including
multispecies biofilms, which are receiving increasing attention due to their ubiquitous presence in
most natural but also man-made habitats. Several studies have shown that species residing within
complex bacterial communities interact both inter and intra-specifically, and that these interactions are
instrumental in shaping the community structure and distribution of bacterial species within
multispecies biofilms. These complex interactions often lead to emergent properties in biofilms, such
as enhanced tolerance against antibiotics, host immune responses, and other stresses, which have been
shown to provide benefits to the biofilm members. Co-culturing studies using in-vitro multispecies
settings have shown these to enhance the overall biomass produced and many studies have revealed
the formation of microbial aggregates, microcolonies or biofilm formation as a response to grazing.
Predation by bacteriovorous protists can influence physiological status of the bacterial communities
and can result in bacterial responses at the community and species levels, which is in turn influenced
by the interplay of several complex interactions and parameters.
In the introductory sections, a comprehensive overview is provided of our current understanding of
multispecies biofilms, biofilm development and interactions between FLP and bacteria in these
biofilms. Special attention is given to predator-prey interactions.
This doctoral thesis aims to address various aspects of bacterial interactions inducing multispecies
biofilm formation and the role of biofilms to serve as a protective growth environment under grazing
pressure. More specifically, we examined microbial composition and diversity of multispecies
biofilms associated with toothbrushes and dishwashers, including FLP occurrence on toothbrushes,
and characterized a range of bacterial communities in dishwasher systems with respect to the species
ability to form biofilms individually and in co-cultures. Further, the influence of bacterial interactions
and its impact on population dynamics in a four-species bacterial model system under grazing
pressure was investigated.
This PhD thesis has resulted in 3 published manuscripts in peer-reviewed journals and one draft
manuscript. The manuscripts follow the order of my work on describing the inter-bacterial
interactions with a focus on biofilm formation and its protective effects observed for different
bacterial species under grazing pressure.
In Manuscript 1, special attention was paid to the occurrence of FLP on toothbrushes. In total, 6 out
of 28 toothbrushes were FLP positive. We show that FLP and bacteria, including some opportunistic
pathogens, were detected and identified from toothbrushes. Amoebae were the dominant FLP
morphotype recovered from toothbrush samples, which may be due to the fact that amoebae have a
higher attachment capacity compared to other FLP morphotypes. Bacterial isolates identified in this
[SUMMARY IN ENGLISH]
22
study which are classified as opportunistic pathogens include Acinetobacter johnsonii, Enterobacter
faecalis, Enterobacter cloacae, Klebsiella oxytoca, Staphylococcus aureus and Streptococcus
salivarius. Toothbrush head design had a significant influence on bacterial diversity and composition
where designed heads fitted with additional projections had a reduced bacterial load on their surfaces
compared to conventional toothbrushes. The result from this study corroborates previous findings that
closely arranged bristles on toothbrushes increase microbial retention.
Biofilm associated microbial communities can thrive in extreme or hostile environments, where
growing as individuals members could be challenging. This aspect was investigated in manuscripts 2
and 3. In Manuscript 2, microbial composition of a man-made system that is household dishwashers
that offer challenging conditions for microbial survival was determined using next generation
sequencing. Growth limiting factors like high temperatures in the rnage between 30 – 80 °C, varying
pH levels ranging 7-12, high salt concentrations, use of detergents and mechanical shear generated
from water ejectors during washing cycles constrain microbial survival in this extreme system. 24
different household dishwashers were investigated for both fungal and bacterial diversity within the
biofilm formed on the rubber seals. In most samples, bacterial genera such as Pseudomonas,
Escherichia and Acinetobacter, known to include opportunistic pathogens, were represented. The
most frequently encountered fungal genera in these samples belonged to Candida, Cryptococcus and
Rhodotorula and representatives of Candida spp. were found at highest prevalence in all sampled
dishwashers. It was also observed that conditions of dishwashers including its age, usage frequency,
and the hardness of the influent water supply to these dishwashers had a significant impact on
bacterial and fungal composition. Pairwise correlations revealed certain bacterial groups to co-occur
and so did the fungal groups. Early adhesion, contact and interactions were speculated to be vital in
the process of mixed fungal- bacterial biofilm, where complexes of the two, bacteria and fungi, could
provide a preliminary biogenic structure for biofilm establishment. In Manuscript 3, four
dishwashers were selected and screened the composition of bacteria and fungi, isolated from a defined
area of one square centimeter of rubber from 4 domestic dishwashers. A total of 80 isolates (64
bacterial and 16 fungal) were obtained and tested for their ability to form multispecies biofilms in-
vitro. 32 out of 140 tested (23%) four-species bacterial combinations displayed consistent synergism
leading to an overall increase in biomass. Bacterial isolates from two of the four dishwashers
generated a high fraction of synergistically interacting four-species consortia. Furthermore, two
synergistic four-species bacterial consortia were tested for their ability to incorporate an opportunistic
fungal pathogen, Exophiala dermatitidis, and establish as biofilms on sterile ethylene propylene diene
monomer M-class (EPDM) rubber and polypropylene (PP) surfaces. When the bacterial consortia
included E. dermatitidis, the overall cell numbers of both bacteria and fungi increased and a
substantial increase in biofilm biomass was observed. This study indicates a novel phenomenon of
cross-kingdom synergy in biofilm formation and further studies are needed to determine their
[SUMMARY IN ENGLISH]
23
potential implications for human health. Our research shows that persisting poly-extremotolerant
groups of microorganisms in household appliances are well established under these unfavorable
conditions, supported by the biofilm mode of growth.
Bacteria protect itself from a predator through various mechanisms and one such mechanism is
biofilm formation that has been shown to confer protection against grazing. While in nature, as most
biofilms were known to harbor different bacterial species, less is known on the effect of grazing with
respect to multispecies biofilm settings. Towards this aspect, in Manuscript 4, the effects of grazing
on the interactions and dynamics of a multispecies bacterial consortium by a pelagic protozoan
predator were investigated. Mono- and multispecies biofilms of four bacterial soil isolates, namely
Xanthomonas retroflexus, Stenotrophomonas rhizophila, Microbacterium oxydans and Paenibacillus
amylolyticus, were cultivated and subjected to grazing by the ciliate Tetrahymena pyriformis. Grazing
strongly reduced the planktonic cell numbers of P. amylolyticus, S. rhizophila and X. retroflexus in
monocultures while the cell numbers in the underlying biofilms of S. rhizophila and X. retroflexus
increased, but not in P. amylolyticus. This may be due to the fact that while grazing enhanced biofilm
formation in the former two species, no biofilm was formed by P. amylolyticus in monoculture, either
with or without grazing. However, in four-species biofilms, biofilm formation was observed to be
higher than in the best monoculture and a strong biodiversity effect was observed in the presence of
grazing. While cell numbers of X. retroflexus, S. rhizophila, and P. amylolyticus in the planktonic
fraction were greatly reduced in the presence of grazers, cell numbers of all three species strongly
increased in the multispecies biofilm. Our results demonstrate that synergistic interactions between
the four-species induced biofilm formation and further suggest that the best biofilm producer X.
retroflexus when exposed to the grazer not only protect itself but also extend supported to other
members that were sensitive to grazing, indicating a scenario of “shared grazing protection” within
the four-species biofilm model.
In conclusion, this PhD thesis demonstrates that studies using multispecies conditions, though low in
complexity comapred to natural bacterial communities, still enable us to get closer to natural settings
where biofilm communities are often present as multispecies microbial communities. The resulting
emergent functions, like increased biomass production and fitness benefits (e.g. grazer protection)
associated within biofilm architecture, underline the prevalence of synergistic interactions in
multispecies biofilms and their impact on individual species during biofilm development.
[SUMMARY IN DUTCH]
24
Samenvatting (Summary in Dutch)
Interspecies interacties zijn van vitaal belang voor de ontwikkeling van complexe gemeenschappen,
waaronder multispecies biofilms, die steeds meer aandacht krijgen door hun alomtegenwoordige
aanwezigheid in de meeste natuurlijke maar ook door de mens gemaakte habitats. Verschillende
studies hebben aangetoond dat soorten die verblijven in complexe bacteriële gemeenschappen inter-
en intraspecifiek interageren, en dat deze interacties behulpzaam zijn bij het vorm geven van de
gemeenschapsstructuur en verspreiding van bacteriesoorten binnen multispecies biofilms. Deze
complexe interacties leiden vaak tot bijzondere eigenschappen in biofilms, zoals verbeterde tolerantie
tegen antibiotica, immuunreacties van de gastheer en andere stressfactoren waarvan is aangetoond dat
ze de biofilm gemeenschap voordelen bieden. Uit in vitro multispecies cultuurstudies is gebleken dat
deze de totale geproduceerde biomassa verbeteren, en veel studies hebben de vorming van microbiële
aggregaten, microkolonies of biofilmvorming als reactie op begrazing aangetoond. Predatie door
bacteriovore protisten kan de fysiologische status van de bacteriegemeenschappen beïnvloeden en kan
resulteren in bacteriële responsen op gemeenschaps- en soortniveau, die op hun beurt worden
beïnvloed door het samenspel van verschillende complexe interacties en parameters.
In de inleidende paragrafen wordt een overzicht gegeven van de huidige kennis van multispecies
biofilms, biofilm ontwikkeling, en interacties tussen vrijlevende protozoa (VLP) en bacteriën. Het
onderzoek in het proefschrift richt zich op verschillende aspecten van bacteriële interacties die in de
biofilmvorming mogelijks optreden, en de rol van biofilms als een beschermend milieu tegen
protozoaire begrazing. Meer in het bijzonder onderzochten we de microbiële samenstelling en
diversiteit van multispecies biofilms geassocieerd met tandenborstels en vaatwassers, inclusief het
voorkomen van VLP op tandenborstels, en karakteriseerden een reeks bacteriële gemeenschappen in
vaatwasser types met betrekking tot het vermogen om biofilms individueel en in co-culturen te
vormen. Verder werd de invloed van bacteriële interacties en de impact ervan op populatiedynamiek
in een bacterieel systeem met vier soorten onder begrazingsdruk onderzocht.
Dit proefschrift heeft geresulteerd in 3 reeds gepubliceerde manuscripten in peer-reviewed
tijdschriften en één manuscript werd ingestuurd.
In Manuscript 1 werd het voorkomen van VLP op tandenborstels onderzocht. In totaal bleken 6 van
de 28 onderzochte tandenborstels VLP-positief. Amoeben waren het dominante VLP-morfotype dat
werd teruggewonnen uit tandenborstelmonsters, wat mogelijk te wijten is aan het feit dat amoeben een
hogere aanhechtingscapaciteit hebben in vergelijking met andere VLP-morfotypes. Bijkomend
werden in deze studie bacteriële isolaten geïdentificeerd die zijn geclassificeerd als opportunistische
pathogenen waar onder Acinetobacter johnsonii, Enterobacter faecalis, Enterobacter cloacae,
Klebsiella oxytoca, Staphylococcus aureus en Streptococcus salivarius. Het ontwerp van de
tandenborstel had een aanzienlijke invloed op de diversiteit en samenstelling van bacteriën, waarbij
[SUMMARY IN DUTCH]
25
koppen met extra uitsteeksels een verminderde bacteriële belasting op hun oppervlakken hadden in
vergelijking met conventionele tandenborstels. Het resultaat van deze studie bevestigt eerdere
bevindingen dat dicht bij elkaar geplaatste haren op tandenborstels de microbiële retentie verhogen.
Met biofilm geassocieerde microbiële gemeenschappen kunnen gedijen in extreme of vijandige
omgevingen, waar groeien als individueel organisme een uitdaging zou kunnen zijn. Dit aspect werd
onderzocht in manuscripten 2 en 3. In Manuscript 2 is de huishoudelijke afwasmachine de
uitdagende omstandigheid voor microbiële overleving. Groei beperkende factoren zoals hoge
temperaturen (30 - 80° C), variërende pH-waarden van 7-12, hoge zoutconcentraties, gebruik van
detergentia en mechanische acties die optreden tijdens wascycli, beperken de microbiële overleving in
dit extreme systeem. In de studie werden 24 verschillende huishoudelijke afwasmachines onderzocht
op zowel schimmel- als bacteriële diversiteit binnen de biofilm gevormd op de rubberen afdichtingen.
In de meeste monsters werden bacteriële genera zoals Pseudomonas, Escherichia en Acinetobacter,
waarvan bekend is dat ze opportunistische pathogenen omvatten, aangetoond. De meest voorkomende
schimmelsoorten in deze monsters waren Candida, Cryptococcus en Rhodotorula, waarbij Candida
spp. bijna in alle afwasmachines werden aangetroffen. Ook werd waargenomen dat de
omstandigheden van vaatwassers, waaronder de ouderdom, gebruiksfrequentie en de hardheid van het
water een significante invloed hadden op de samenstelling van bacteriën en schimmel
gemeenschappen. Analyse bracht aan het licht dat bepaalde bacteriegroepen en schimmelgroepen
gelijktijdig voorkwamen. Vroege adhesie, contact en interacties werden als hypothese naar voren
geschoven als essentieel in het proces van gemengde schimmel-bacteriële biofilms, waarbij
complexen van de beiden, bacteriën en schimmels, een voorlopige biogene structuur konden bieden
voor de vestiging vaneen biofilm. In Manuscript 3 werden vier vaatwassers geselecteerd en
gescreend op de samenstelling van bacteriën en schimmels, geïsoleerd uit een afgebakend gebied van
een vierkante centimeter rubber. Een totaal van 80 isolaten (64 bacteriële en 16 schimmel) werden
verkregen en getest op hun vermogen om in vitro multispecies biofilms te vormen. Tweeëndertig van
de 140 (23%) geteste bacteriesoorten met vier soorten vertoonden consistente synergie, wat leidde tot
een algehele toename van biomassa. Bacteriële isolaten van twee van de vier vaatwassers
produceerden een hoge fractie van synergistisch wisselwerkende consortia van vier soorten. Verder
werden twee synergetische bacteriële consortia met vier soorten getest op hun vermogen om een
opportunistische schimmelpathogeen, Exophiala dermatitidis, op te nemen en zich te vestigen als
biofilm op steriele EPDM rubber en polypropyleen (PP) oppervlakken. Toen de bacteriële consortia
E. dermatitidis omvatten, namen de cel aantallen van zowel bacteriën als schimmels toe en werd een
aanzienlijke toename in biofilm-biomassa waargenomen. Deze studie wijst op een nieuw fenomeen
van synergie tussen de verschillende organismen in biofilm vorming. Verdere studies zijn nodig om
de mogelijke implicaties voor de volksgezondheid te bepalen. Ons onderzoek toont alvast aan dat
[SUMMARY IN DUTCH]
26
persistente poly-extremotolerante groepen van micro-organismen in huishoudelijke apparaten onder
deze ongunstige omstandigheden goed zijn aangepast, ondersteund door de biofilm-modus van groei.
Bacteriën beschermen zichzelf tegen een predator door verschillende mechanismen, en een dergelijk
mechanisme is biofilm vorming waarvan is aangetoond dat het bescherming biedt tegen protozoair
grazen. Van de meeste natuurlijk voorkomende biofilms is bekend dat ze verschillende
bacteriesoorten herbergen, maar is er minder bekend over het effect van begrazing met betrekking tot
multispecies biofilm samenstellingen. Daarom werd in Manuscript 4 de effecten onderzocht van
begrazing op de interacties en dynamica van een multispecies bacterieel consortium door een
pelagisch protozoönroofdier. Mono- en multispecies biofilms van vier bacteriële bodem isolaten,
namelijk Xanthomonas retroflexus, Stenotrophomonas rhizophila, Microbacterium oxydans en
Paenibacillus amylolyticus, werden gekweekt en onderworpen aan begrazing door de ciliaat
Tetrahymena pyriformis. Begrazing verminderde sterk de planktonische cel aantallen van P.
amylolyticus, S. rhizophila en X. retroflexus in monoculturen, terwijl de cel aantallen in de
onderliggende biofilms van S. rhizophila en X. retroflexus toenamen, maar niet in P. amylolyticus. Dit
kan te wijten zijn aan het feit dat tijdens begrazing de biofilm vorming bij de eerste twee soorten werd
verbeterd, maar geen biofilm werd gevormd door P. amylolyticus in monocultuur, met of zonder
begrazing. In biofilms met vier soorten werd echter waargenomen dat de biofilm vorming hoger was
dan in de beste monocultuur en een sterk biodiversiteitseffect werd waargenomen in de aanwezigheid
van begrazing. Terwijl de cel aantallen van X. retroflexus, S. rhizophila en P. amylolyticus in de
planktonfractie sterk waren verminderd in aanwezigheid van grazers, namen de cel aantallen van alle
drie soorten sterk toe in de multispecies biofilm. Onze resultaten tonen synergetische interacties aan
tussen de door vier soorten geïnduceerde biofilmvorming, en suggereert verder dat de beste biofilm
producent, X. retroflexus, bij blootstelling aan de grazer zichzelf niet alleen beschermt maar ook
ondersteuning aan andere leden biedt die gevoelig zijn voor begrazing. Dit duidt op een scenario van
"gedeelde begrazingsbescherming".
Concluderend toont dit proefschrift aan dat studies met multispecies-condities, hoewel ze laag in
complexiteit zijn vergeleken met natuurlijke bacteriële gemeenschappen, ons toch in staat stellen om
dichter bij natuurlijke omgevingen te komen waar biofilmgemeenschappen vaak aanwezig zijn als
multispecies microbiële gemeenschappen. Reacties zoals verhoogde biomassaproductie en
fitnessvoordelen (bijv. grazer-bescherming) geassocieerd met biofilmarchitectuur, benadrukken het
voorkomen synergistische interacties in multispecies biofilms en hun invloed op individuele soorten
tijdens biofilmontwikkeling.
[SUMMARY IN DANISH]
27
Resumé (Summary in Danish)
Interaktioner mellem bakterier tilhørende forskellige arter er vitale for udviklingen af komplekse
mikrobielle samfund, inklusiv multispecies biofilm, og da multispecies biofilm er udbredte i de fleste
naturlige- og menneskeskabte miljøer, er de genstand for stadig øget opmærksomhed. Flere studier
har vist, at bakteriearter der lever i komplekse bakterielle samfund interagerer både intra- og
interpecifikt, og at disse interaktioner er instrumentelle for, hvordan samfundsstrukturen etableres og
distributionen af bakteriearterne i multispecies biofilm forekommer. Disse komplekse interaktioner
resulterer ofte i, at bakterierne udvikler egenskaber, som de ikke ville have, hvis de voksede alene, og
derfor ikke var en del af et komplekst samfund. Disse ”emergent properties” er blandt andet øget
tolerance over for antibiotika, værtsimmunresponser og andre stressfaktorer, hvilket har vist sig at
give medlemmerne af biofilm særlige fordele frem for de artsfæller, der lever uden for en
samfundsstruktur. Studier af co-kultivering i in-vitro multispecies opsætninger viser, at bakterier i
blandende samfund generelt producerer øget biomasse, og mange studier kan dokumentere dannelse
af mikrobielle aggregater, mikrokolonier eller biofilmformation som respons på tilstedeværelsen
protozoer, der lever af ”frie” bakterier (grazing). Truslen fra bakteriespisende protozoer kan influere
bakteriesamfundets fysiologiske status og kan resultere i responser fra bakterierne på både arts- og
samfundsniveau, hvilket igen er influeret af sammenspillet mellem forskellige interaktioner og
parametre.
Formålet med denne ph.d.-afhandling er at adressere forskellige aspekter ved bakterielle interaktioner,
der alle understøtter dannelse af multispecies biofilm, og at undersøge biofilmenes rolle som
beskyttende vækstmiljøer under forhold, hvor grazing er udbredt. Mere specifikt har jeg undersøgt
den mikrobielle diversitet af multispecies biofilm og udvalgte eukaryote organismer (hhv. protozoer
og mikrosvampe) associeret med tandbørster (manuskript 1) og opvaskemaskiner (manuskript 2 og
3). Jeg karakteriserede flere bakterielle samfund i opvaskemaskiner i henhold til deres evne til at
producere biofilm; både individuelt og i co-kulturer. Jeg undersøgte ydermere indflydelsen af
bakterielle interaktioner på populationsdynamikker i en modelkultur med 4 forskellige
bakteriestammer, der udsættes for grazing (manuskript 4).
Resultaterne præsenteret i denne afhandling viser, at studier foretaget under multispecies
omstændigheder, også selvom de er mindre komplekse end naturligt forekommende bakteriesamfund,
gør det muligt for os karakterisere biofilm, som de findes i deres naturlige miljøer, hvor de oftest
eksisterer som multispecies mikrobielle samfund. De deraf resulterende emergent properties så som
øget biomasseproduktion og fitness fordele (beskyttelse mod grazing), som associeres med
biofilmstrukturen, underbygger forekomsten af synergistiske interaktioner i multispecies biofilm og
fremhæver deres indflydelse på individuelle arter under biofilm dannelse.
[DISCUSSION]
28
General Discussion
Microbiota in many ecosystems occur as adherent biomasses or biofilms often display complex
ecological and evolutionary relationships between the different microbial members like fungi,
bacteria, virus, protozoa and nematodes (Brown & Barker, 1999). Biofilm formation represets a
simplistic survival strategy where microorganisms exist under diverse environmental conditions and
may serve as a protective niche for pathogens in the natural environment, when not associated with a
host (Lindsay & von Holy, 2006). For e.g. Vibrio cholerae biofilms on crustacean shells, aquatic
insects and plants could release higher amounts of cells from these biofilms to cause human infections
(Hall-Stoodley & Stoodley, 2005). Variuos virulence factors like flagella, pili, alginate as in the case
of P. aeruginosa, found expressed during biofilm development (Kipnis et al., 2006) and
heterogeneous microenvironments that occur within biofilms support the emergence of altered
phenotypic and genotypic variants capable of surviving changing environmental conditions and might
also facilitate infection (Hall-Stoodley & Stoodley, 2005). Also, other microrganisms like fungi
capable for biofilm formation represent emerging novel opportunistic agents responsible for many
nosocomial infections. Examples include Candida sp., Aspergillus sp, Trichoderma sp., and Fusarium
sp. (Jabra-Rizk et al., 2004; Patterson, 2005). Further, biofilm formation by foodborne spoilage and
pathogenic bacteria cause economic loss as they often lead to product contamination during food
processing, and medical devices and prostheses related infections (Donlan and Costerton, 2002;
Lindsay et al., 2006). However, not all biofilms are detrimental as mixed species biofilms with
coordinated functions have been exploited for beneficial roles for e.g. in the bioremediation processes
of human and manufacturing wastes (Skariyachan et al., 2016; Norton et al., 2000; Hiraishi et al,
1998). Protective roles of bacterial biofilms have been described for e.g. in human gut, where mixed
consortia of commensal bacteria attached to gut epithelial cells provide a barrier against foodborne
pathogens; tooth decay and dental plaque as a result of proliferation of disease-producing strains are
usually kept in check by the natural commensal bacteria present in these biofilms on tooth surfaces
(Whittaker et al., 2001; Lee et al., 2000). Hence, the ability of bacteria to communicate and behave as
multi-cellular organism shaped by social interactions during biofilm formation has provided
significant benefits to bacteria in host colonization, defense against competitors/predators, and
adaptation to changing environments.
Understanding the microbial ecology of and the underlying interactions among microbial species in
biofilm communities form the core of this PhD research. The focus to perceive biofilm formation as a
consequence of microbial interactions has paved the way to better understand their dynamics (Yang et
al., 2011). Social interactions in multispecies biofilms determine the function and/or composition of
microbial community processes either through synergistic or antagonistic interactions or both between
its members (Flemming et al., 2016). Social interactions can be particularly more pronounced in
multispecies biofilm settings (Burmølle et al., 2014) and have been shown to enhance biomass
[DISCUSSION]
29
production compared to single species biofilms in soil (Manuscript 4; Ren et al., 2015), sea water
(Burmølle et al., 2006), industrial settings (Røder et al., 2015) and household appliances, e.g.
dishwashers (Manuscript 3). It has also been reported that biofilm growth and structural integrity
promote cooperation (Nadell et al., 2009), and extensive studies on such cooperative effects in
multispecies biofilms have, in addition to increases in overall biomass production, also reported
enhanced co-aggregation, resistance of the community against external stresses and an expanded
niche availability benefitting the community (Rickard et al., 2003; Burmølle et al., 2006, Nadell et al.,
2009).
The resilience of biofilm mode of growth to hostile environments, and investigating the microbial
diversity of biofilms in both man-made systems and extreme natural environments have gained
popularity in recent years. Examples include studies on microbial biofilm communities formed on
cave walls (Borsodi et al., 2012), in acidic geothermal areas (Urbieta et al., 2015), hot springs
(Boomer et al., 2009) and lakes with high salinity, and in the presence of extreme pH values and
heavy metal contamination (Rascovan et al., 2016). Likewise, man-made systems like coffee
machines, dishwashers and washing machines have been observed to support the growth of microbial
communities (Gümral et al., 2016; Mattick et al., 2003; Vilanova et al., 2015). Toothbrushes present
another system where different microbes could attach and establish as biofilms, resulting in potential
health effects. In Manuscript 1, the microbiota of biofilms on toothbrushes with specific head/bristle
designs were examined. In Manuscript 2, the microbial community composition of both bacterial and
fungal origin, from different dishwasher associated biofilms was elucidated using next generation
sequencing. Furthermore, in manuscript 3, fungal and bacterial isolates obtained from these systems
by traditional culture methods were tested for their ability to engage in in vitro multispecies biofilm
development. Our observations showed that some four-species bacterial combinations led to an
overall increase in biomass, suggesting synergistic interactions. The bacterial consortia also displayed
the ability to incorporate an opportunistic fungal pathogen, Exophiala dermatitidis and facilitate its
establishment in biofilms. Growth limiting factors, such as varying temperatures, high and low pH,
action of detergents and shear force generated by water ejectors determine the survival of microbes in
this extreme system. Manuscript 2 and Manuscript 3 support the general notion that different
microbial groups become well established and persist under unfavorable conditions by growing as a
biofilm. Microbial growth in these systems, in turn, has been speculated to select and drive their
evolution towards acquiring polyextremotolerant traits (Gostincar et al., 2010).
Our observations show that bacterial fitness is enhanced in multispecies biofilms. If the fitness
advantage applies to the whole community, then the underlying interactions are characterized as
cooperative (West et al., 2007). In Manuscript 3, though we did not determine if all individual
members gained benefit within the four-species assembly, these mixed co-cultures increased their
[DISCUSSION]
30
biofilm biomass compared to their monospecies counterparts. Also, the poor biofilm producers
benefitted by joining the multispecies settings. This was observed in the case of E. dermatitidis, where
the fungal cells do not attach themselves when grown alone, but becomes well established in biofilms
when together with bacterial consortia. These results corresponds to previous findings that the biofilm
forming ability of an individual microbe is not necessarily an indicator for its potential in multispecies
biofilm formation (Bharathi et al., 2011; Burmølle et al., 2014).
Studies on inter-species interactions within two-species models using bacterial isolates from tree-hole
rainwater pools showed that the majority of bacterial interactions tend to follow a competitive trend
affecting the individual species and overall productivity negatively (Foster & Bell, 2012). However,
follow up studies showed that species interactions between five co-occurring bacterial species
stimulated divergence in resource use, leading to enhanced productivity of the entire community
(Lawrence et al., 2012), and that the multispecies biofilm over time evolves to optimize resource
partitioning between the bacterial members present in tree-hole rainwater pools (Fiegna et al., 2014).
In line with this, it was also showed that an increase in multispecies biofilm formation correlated with
long-term coexistence of microbial communities in different environments (Madsen et al., 2016). This
underlines the importance of multiple interactions where all organisms co-occur with many other
species in diverse systems and influence how component species adapt with respect to environment,
which in turn, could have consequences for ecosystem functioning (Lawrence et al., 2012).
As explained above, the key to whether bacterial species compete or co-operate lies in their possibility
of long-term co-adaptation and degree of niche overlap (Rivett et al., 2016). In our biofilm
cultivation experiments in Manuscript 3, the bacterial isolates were obtained from different
dishwasher systems and tested for their ability to form multispecies biofilms. We observed that the
numbers of synergistic bacterial consortia were highest for a dishwasher used for eight years. Also,
biofilm formation was induced when these isolates were given the commonly used ethylene propylene
diene monomer M-class (EPDM) rubber and polypropylene (PP) substrates in dishwasher systems for
biofilm attachment. This further supports the experimental evidence that long-term coexistence
facilitated biofilm formation among bacteria that have coexisted in their original environment
(Madsen et al., 2016). We also found that synergistic interactions indicative of cooperative effects
were dominant in two of the dishwashers (dishwasher sample 1 and 4) where most four-species
assembly contained phylogenetically diverse members, whereas in dishwasher 2 and 3, the isolate
composition was less phylogenetically diverse, having lower numbers of synergistic combinations.
This trend was also reported in a pairwise interaction study where that the probability of antagonism
increased among closely related bacteria with similar carbon metabolism (Russel et al., 2017). In
addition, divergence in resource utilization owing to multiple species consuming different resources
may contribute to increased community productivity. This was evidenced in a study by Rivett et al.,
2016 where bacterial communities displaying stronger antagonistic interactions during the early stages
[DISCUSSION]
31
of colonization and acclimatization, to novel biotic and abiotic factors, declined over time to produce
a stable community.
Our observations in Manuscript 3 focus on the overall community level as it is often difficult to
address the specific interspecies interactions directly. This leaves the question of species interactions
open when exploring the population dynamics. To address the above issue, there is a need for a well-
studied and reproducible microbial model community to study individual population dynamics during
biofilm development, as in the case of the four-species model applied in Manuscript 4. Briefly, the
four-species biofilm model comprising X. retroflexus, M. oxydans, S. rhizophila and P. amylolyticus,
showed a significant increase in biofilm biomass and in their individual cell numbers when the four
species were co-cultured together as compared to their monospecies biofilms (Ren et al., 2014,
2015a). Furthermore, meta-transcriptome analysis revealed distinct gene expression patterns in the
four-species biofilm model compared to single and two-species species combinations; with a set of
genes found to be expressed only in the four-species combination. Also, a number of genes were
down-regulated, underlying the lifestyle benefits in multispecies biofilm that allowed X. retroflexus to
mute certain costly functions which otherwise were vital for its existence in monoculture (Hansen et
al., 2017). A meta-proteomic approach on this four-species model also showed that community
development depended on cooperative interactions facilitated by cross-feeding of specific amino acids
between its community members (Herschend et al., 2017).
Predation by protozoa limits bacterial population sizes and can influence bacterial community
composition and structure (Murase et al., 2006). Data on free living protozoa (FLP) in biofilms
however are limited to a few studies (e.g. Gourabathini et al., 2008; Vaerewijck et al., 2011) and
microbiological surveys from natural environments often do not include a characterization and
identification of protozoa. They are also hampered by the current isolation techniques (Chavatte et al.,
2016). In addition, finding the optimal conditions to maintain protozoan cultures is largely a process
of trial and error and few standard protocols exist. Moreover, their detection and identification is time-
consuming and relies mostly on microscopic identification. Recently, NGS methods are being
developed for protozoan identification (Hu et al., 2015; Jung et al., 2015; Moreno et al., 2018).
However, sequencing based approaches requires efficient DNA extraction and applying directly to
detect protozoa from environmental samples where their cell numbers are scarce, pose a challenge.
Our labs have previously detected FLP on vegetable sprouts, poultry and have repeatedly worked with
environmental samples (Baré et al., 2009; Chavatte et al., 2014, 2016). Building on this expertise and
knowledge, we identified and detected protozoa from toothbrushes in Manuscript 1, where we report
that most protozoa belonged to the amoeba morphotype. This could be due to the choice of isolation
method, as it has been reported that amoebae were the most abundant morphogroup following
cultivation on stomachered homogenate (Chavatte et al., 2016). Other reasons include the general
higher attachment capacity of amoebae and also the possible formation of amoebal cysts that enables
[DISCUSSION]
32
these organisms to become dormant in changing environments and return to trophozoite mode of
growth when conditions become favorable.
Apart from controlling bacterial populations, grazing by FLP has been implicated in the persistence of
food-borne pathogens. While most of the efforts to characterize FLP-bacteria interactions focus on
single bacterial species, more studies to elucidate the role of FLP in biofilm-related environments are
needed. With one proposed view that biofilms offer grazing resistance in bacterial communities, the
protective nature of multispecies biofilm and the corresponding effects on population dynamics were
examined under grazing pressure in the four-species biofilm model community described above in
Manuscript 4. Our results showed a synergistic induction of four-species biofilm formation under
grazing by T. pyriformis. Moreover, P. amylolyticus and S. rhizophila were most susceptible to
grazing under monospecies conditions and the poor biofilm producer P. amylolyticus gained
protection in the four-species co-culture. It was also shown that grazing promoted a surface-associated
life style in the consortia, i.e. increased biofilm biomass based on crystal violet staining and cell
numbers based on colony forming units, while planktonic cells numbers decreased due to grazing.
This indicates the necessity of structured environments for synergistic interactions to occur that could
be favored by co-metabolism (Ren et al., 2015) and hence offer a protective environment (Burmølle et
al., 2014). Similar to our results, protozoa grazing has been reported to also induce microbial
aggregate formation in aquatic ecosystems where interspecies interactions significantly increased
productivity in terms of overall bacterial numbers and carbon transfer efficiency (Corno et al., 2015).
The view that multispecies biofilms offer a protective habitat was challenged by Kart et al., 2014,
where it was observed that the extent of protective effects were mainly characteristic of individual
members that constitute the biofilm. Also, the protective effects of multispecies biofilm formation
under grazing pressure were dispelled in other studies (Huws et al., 2005; Weitere et al., 2005). These
conflicting observations could be due to the choice of bacterial isolates, biofilm cultivation methods,
different definitions of synergism/cooperation and the type of protozoa used in grazing experiments.
Studies have shown protozoa feeding traits to influence grazing resistance in bacterial biofilms (Seiler
et al., 2017) and surface associated bacteria can be even more consumed when exposed to a
specialized grazer (Rogerson & Laybourn-Parry, 1992). Therefore, more studies with different grazers
are needed for a comprehensive understanding of the effect of grazing by protozoa on bacterial
biofilms. Moreover, in our studies, each multispecies consortium tested (Manuscripts 3 and 4)
consisted of bacteria originating from same place and isolated from smaller areas with a likelihood of
having encountered each other in their original environment. All co-cultured bacteria also have
similar resource patterns (use of same culture conditions) during their growth. This allows to perform
experiments that closely resemble the natural environment, as mentioned in a review investigating
bacterial biofilms (Røder et al., 2016).
[DISCUSSION]
33
In conclusion, based on our results, there is strong evidence that synergistic interactions are important
for co-existence and protection of bacteria. The overall increase in biofilm formation was indicative of
strong positive interactions between different microbial species adapting to each other in their relevant
ecological settings and under stressful conditions such as grazing. It is recognized that multispecies
biofilms are dynamic communities within a vast network of interactions between different species.
The analyses of such communities are consequently very complex and the emergent properties within
these systems have a wide range of consequences for the survivability of the bacteria. This illustrates
the need to further broaden our view on multispecies systems and to unravel the complexity of
interspecies interactions as their emergent functions cannot be solely determined using mono-species
experiments.
[FUTURE PERSPECTIVES]
34
Future Perspectives
Studies on multispecies biofilms in recent years have revealed that microbial community physiology
and functions depend on complex interactions between the community members, together with
several abiotic factors. These interactions need to be taken into account when assessing the potential
of communities for both beneficial uses in e.g. degradation of waste and when addressing pathogenic
scenarios, e.g. in clinical settings. In this thesis, the potential of different bacteria to form stable
multispecies consortia were assessed. This aspect could be applied to formulate novel beneficial
microbial consortia and thereby devise an eco-friendly approach for enhanced degradation of wastes.
It was also identified that different species contribute as key biofilm players and developing strategies
and limiting their role could broaden our knowledge on eliminating biofilms.
It would be of interest also to investigate the effects of a particular stress or combinations of different
stresses (e.g. the dishwsasher conditions) on biofilm development, as multispecies biofilms confer
benefits such as increased tolerance to antibacterial compounds and enhanced protection from
desiccation. Also, interactions at multispecies levels and their subsequent biofilm formation result in
the production of new types of polysaccharides having different composition, and this research aspect
could be applied to further understand the role and composition of EPS in multispecies biofilms.
Correspondingly, the use of a specialized surface-associated grazer could also benefit our studies in
explaining if biofilms always offer a protective niche during predation or whether this depend on the
type of grazer present. Recent technological advancements including meta-genomics, meta-
transcriptomics and CLASI - FISH probing can be employed to further our insights into the structural
and functional dynamics within complex communities. This will open a new door leading to further
in-depth studies enabling to predict the outcome of specific interactions. However, it must also be
emphasized that multispecies interaction studies benefit and have benefitted from studies using mono-
species biofilms and addressing single cell functions, where researchers have identified different
genes and regulators important for biofilm formation, that could be used to help and understand
complex communities.
Thus, combinations of simple and complex approaches in microbiology are necessary to appropriately
understand how these complex biofilms are shaped and how they function. Obtaining this will lead to
a new understanding of multispecies biofilm and the possibilities for efficient utilization of
communities.
[BIBLIOGRAPHY]
35
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Identification of Free Living Protozoa (FLP) and bacterial composition on toothbrushes
Prem K Raghupathi a, b
, Charles Dumolinc, Mette Burmølle
a, Søren J. Sørensen
a, Koen Sabbe
d, Kurt
Houf b, c
a Molecular Microbial Ecology Group, Section of Microbiology, Department of Biology, University of
Copenhagen, Copenhagen, Denmark; b
Department of Veterinary Public Health and Food Safety, Faculty
of Veterinary Medicine, Ghent University, Merelbeke, Belgium; c Laboratory of Microbiology,
Department of Biochemistry and Microbiology, Ghent University, Ghent, Belgium and d Department of
Biology, Faculty of Sciences, Ghent University, Ghent, Belgium
Interactions between free-living protozoa (FLP) and bacteria have been implicated in the
persistence of pathogenic bacteria in various environments. In this study, we show that FLP and
bacteria, including some opportunistic pathogens, were detected and identified from toothbrushes.
Amoebae were the dominant morphotype recovered from toothbrush samples. The toothbrush
head design had a significant influence on bacterial diversity and composition where designed
heads fitted with additional projections had reduced bacterial load on its surfaces compared to
conventional toothbrushes. The results from this study support previous findings that closely
arranged bristles on toothbrushes increase microbial retention.
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Introduction
The oral cavity represents an ecosystem where different microorganisms can prosper and oral biofilm
harbors the most diverse microbes within the human body (Chandki et al., 2011; Edlund et al., 2015; Hall
et al., 2017). The microbiome of the oral cavity have been reported to host over 700 different bacterial
species (Beneduce et al., 2010; Dewhirst et al., 2010; Hall et al., 2017; Kilian et al., 2016) and provides
the perfect portal of entry for microbes to access new hosts (Edlund et al., 2015). Toothbrushes can act as
a reservoir and transmission tool for microorganisms, including pathogenic bacteria (Caudry et al., 1995).
At this moment, studies focused on revealing the microbial communities on toothbrushes have gained
momentum (Eichenauer et al., 2014; Karibasappa et al., 2011; Rodrigues et al., 2012). In addition to the
oral cavity itself; human contact, storage conditions of toothbrush and aerosols all serve as possible
contamination routes of toothbrushes (Beneduce et al., 2010; Frazelle & Munro, 2012). Storage in
bathrooms as well contribute as a site for contamination due to their humid and wet environment (Glass &
Jensen, 1988; Scott et al., 1982).
Commercially available toothbrushes are produced with different configurations in the bristles type,
bristle cluster, type of plastic moldings and/ or rubber fittings. Very few studies have assessed the impact
of such designs on bacterial accumulation. It was reported that bacteria become more trapped within the
insides of the bristles (Bunetel et al., 2000) and closely arranged bristles showed increased bacterial
retention (Goldschmidt et al., 2004). Bacterial survival also increased with the retention of moisture and
oral debris within the bristles (Mehta et al., 2007). In addition to bacteria, fungi and yeasts were reported
to contribute to the microbial load of toothbrushes (Malmberg et al., 1994; Mobin et al., 2011). Bacteria
attach to, accumulate and survive on toothbrushes, and it has been shown that a toothbrush in regular use
becomes heavily colonized over time (Verran & Leahy-Gilmartin, 1996).
The main use and purpose of a toothbrush is in the removal of the dental biofilm formed by different
microorganisms on the teeth (Collins, 2014); which, in turn, results in the colonization by microorganisms
onto toothbrush surfaces. Studies have shown that toothbrushes support the growth of biofilm forming
microbial strains (Abubakar et al., 2013; Devine et al., 2007; Sammons et al., 2004). Microbial growth as
biofilms could be due to structural or protective nature, i.e. the physical structure that the toothbrush
bristle filaments offer or the regular contact with microbial disinfectants. Microbial biofilms have been
shown to confer resistance against disinfectants and enhance persistence of certain microorganisms
(Bridier et al., 2011).
Free-living protozoa (FLP) have been isolated from various habitats like soil, marine and fresh waters,
geothermal springs, dental units and hospital water networks (Aguilera et al., 2010; Arias Fernandez et
al., 1989; Armand et al., 2016; Bass & Bischoff, 2001; Hikal et al., 2015; Trabelsi et al., 2016). Bacteria
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and protozoa were found to coexist in biofilm environments (Matz et al., 2008) and several studies have
assessed the relationship between FLP and pathogenic bacteria (Lambrecht et al., 2015; Matz et al.,
2008). FLP are widely recognized as important bacterial consumers controlling bacterial biomass, and
form an important trophic link in aquatic and terrestrial food webs (Barker & Brown, 1994; Pernthaler,
2005; Sherr & Sherr, 2002). FLP were reported to be present in refrigerators (Vaerewijck et al., 2010) and
be part of the ‘in-house microbiota’ of food related environments (Vaerewijck et al., 2014).To date, most
studies on microbial colonization of toothbrushes have addressed bacterial and fungal diversity; at present
no information on the occurrence of FLP and bacterial pathogens on toothbrushes is available.
Toothbrushes could present a habitat where FLP and opportunistic pathogenic bacteria coexist. The aims
of the present study were to investigate bacterial load and diversity and the presence and diversity of
FLPs on toothbrushes and to assess the impact of toothbrush head design on microbial colonization.
Materials and Methods
Study design and sample preparation
Two commercially available toothbrush head designs (‘conventional’ and ‘designed’, Fig 1) were used in
assessing the FLP diversity, and to enumerate the total bacteriological load. Fourteen conventional and
fourteen designed toothbrushes were provided to 28 unrelated individuals and were used for 6 weeks.
Five toothbrushes from each design were also provided to 10 individuals and these were left open in their
respective bathroom environment but were not used to brush with, in order to evaluate environmental
cross contamination. Toothbrushes were collected in sterile bags; air dried and processed the next day.
New, unopened toothbrushes were included as blank controls. Each toothbrush sample was processed by
removing the head from the handle and the bristles were cut using a sterile scalpel. The head and bristles
of each sample were then transferred into a sterile stomacher bag containing 20ml 1X PAS solution and
homogenized with a peristaltic homogenizer for 5 minutes.
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Figure 1: (A) Conventional and (B) designed head of toothbrushes selected in this study.
Isolation and detection of FLP
A previously established protocol to isolate and identify FLP from sample surfaces (Chavatte et al., 2014)
was used in the present study. 19 ml of the homogenate were transferred to 25 cm2 tissue culture flask and
Page’s amoeba saline (PAS) was added to a final volume of 20 ml. Sterile uncooked rice grains were
included as a carbon source to stimulate microbiological growth. The culture flasks were stored in dark at
room temperature and the cultures were examined after two days and for up to 1 week for the presence of
FLP. Repeated examinations were essential as there could be rapid turnover of FLP species in cultures.
FLP in sample homogenate were detected by inverted light microscopy (magnification ×400) and were
identified based on their morphology and mobility. Organisms were assigned to morphotypes (ciliates,
flagellates or amoeba) as specified by Smirnov & Goodkov, 1999 and Smirnov & Brown, 2004.
Bacteriological Enumeration and Identification
One ml of the homogenate was used for bacteriological enumeration and isolation. 10-fold serial dilutions
were performed and 100 µl of the suspensions were plated on plate count agar (PCA, Oxoid, Basingstoke,
UK) to enumerate the total aerobic bacteria; 5% blood agar plates (BA, Oxoid England) and mannitol salt
agar plates (MSA, Oxoid England). All plates were supplemented with 50µg/ml cycloheximide to inhibit
the growth of fungi. The PCA plates were incubated aerobically at 28 °C for 48 hours, BA plates at 37 °C
for 24 hours and MSA plates at 30 °C for 48 hours. Colonies formed after incubation were isolated as
pure cultures. Bacterial smears from these pure cultures were prepared according to manufacturer’s
instruction (Standard operating procedure (SOP); Direct Transfer Method) and examined using Bruker
LT/SH microflex MALDI-TOF MS (matrix-assisted laser desorption/ionization time-of-flight mass
spectrometer) (Bruker Daltonics, Bremen, Germany). Each series of measurements was preceded by a
calibration step with a bacterial test standard (BTS 155 255343; Bruker Daltonics) to validate the run. The
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spectra were generated using MALDI Biotyper automation control, using Flex Analysis software 3.4
(Bruker Daltonics) following manufactures operating methods in linear positive ion detection mode.
Identifications were obtained by comparing the mass spectra to the Bruker MSP database (version
DB5989) using the Bruker Compass 4.1.31 software (Bruker Daltonics) at default. Identification scores ≥
2.0 were considered reliable for identifying bacteria classified to the species level and scores ≤ 1.7 were
not included in the analysis. Scores between 1.7 and 2.0 indicated ambiguous species identification and
were re-examined using bacterial cell extracts prepared according to manufacturer’s instruction (SOP;
Formic Acid Extraction Method).
Data Analysis
Results of bacterial enumeration were expressed as colony forming units (CFU) /toothbrush head. A
sample was considered FLP positive if at least one of the morphogroup was observed; these were then
recorded as binary variables (presence/absence) as performed previously (Chavatte et al., 2016). The
bacteria isolated and identified from a sample were compared against the total isolates obtained from each
sample. Hence, the relative abundances of bacterial isolates were calculated and log10 transformed.
Redundancy Analysis (RDA) based variation partitioning was used to determine the degree to which the
toothbrush designs and the presence of protozoa related to the variation in bacterial community
composition. Significance of the model, of the RDA axes and each of the factors were tested using an
ANOVA- like permutation test (999 permutations). Differences in the bacterial community composition
between the two toothbrush designs were evaluated using the software package STAMP 2.1.3 (Parks et
al., 2014) and significance checked using ANOVA (Tukey-Kramer post-hoc test). Benjamini-Hochberg
FDR corrections were applied to account for multiple testing. Dissimilarity indices were computed based
on ‘Euclidean distances’ and scaled heat maps were hierarchically clustered (hclust) using ‘coniss’
method. Heat maps of significant bacterial species were generated using available packages: gplots,
vegan, rioja and Rcolorbrewer for Rstudio3.2.0.
Results
Bacterial load and community composition in toothbrushes
Total aerobic bacterial counts ranged from 2 – 5.6 log10 CFU/head in designed toothbrush heads and 3.9–
7.5 log10 CFU/head in conventional heads. Among the toothbrush groups used for checking cross-
contamination, the total aerobic counts ranged between 1.5 – 2.3 log10 CFU/ head and 0.6 – 2.6 log10 CFU/
head for designed and conventional toothbrushes respectively (Table S1, supplementary information). No
colonies were obtained from unopened and unused toothbrushes. A total of 1187 of which 1035 bacterial
isolates (526 isolates from conventional and 509 isolates from designed toothbrushes) with MALDI-TOF
MS based identification score ≥ 2.0 were obtained. The remaining 132 isolates rendered no possbile
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identification using Bruker system. Further, a total of 186 (100 from conventional heads and 86 from
designed heads) isolates were observed on toothbrushes that were left open in the environment and not
used (cross-contamination control). Both toothbrush groups had isolates represented by the genera
Enterobacter, Kocuria, Nocardioides, Pseudomonas, Micrococcus and Staphylococcus. Bacterial isolates
belonging to the genera Staphylococcus, Pseudomonas and Micrococcus were present in more than 80%
(n>12) of each sample group (Fig 2A). Isolates belonging to the genera Pantoea, Brevibacterium,
Stenotrophomonas, Streptococcus and Raoultella were recovered in conventional toothbrushes, and
Microbacterium was present only in designed toothbrushes (Fig 2A). Among the cross-contamination
control groups, 10 bacterial genera were identified. Micrococcus was present in all samples of both head
designs (n=5/5). Other genera included Kocuria, Staphylococcus and Pseudomonas identified at varying
levels across samples. Enterobacter was isolated from 4 out of the 5 conventional toothbrushes whereas
Nocardioides was identified in 3 out of the 5 designed toothbrush samples (Fig 2 B and C).
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Figure 2: (A) Bacterial genera that were represented in conventional (CB) and designed (DB) toothbrush
head samples (n ≥ 5, N=14). Bacterial genera that were present as cross contaminants in the toothbrush
groups; (B) conventional toothbrush controls and (C) designed toothbrush controls (n ≥ 3, N= 5)
Identification by MALDI-TOF MS revealed varying species diversity, i.e. the percentage of samples
containing the species. Micrococcus luteus, Nocardioides sp, Staphylococcus hominis and Staphylococcus
epidermis were the most prevalent species isolated from both sample groups (Fig 3). Enterobacter
faecalis was highly represented in designed toothbrush heads compared to conventional toothbrush heads.
Also, a higher diversity of different Pseudomonas spp. were identified and isolated from conventional
toothbrush heads (Fig 3).
B C
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Figure 3: Distribution of bacterial species across the conventional (CB) and designed (DB) toothbrush
samples.
Isolates identified in this study, were classified as opportunistic pathogens and few examples include
Acinetobacter johnsonii, Enterobacter faecalis, Enterobacter cloacae, Klebsiella oxytoca, Staphylococcus
aureus and Streptococcus salivarius (Hoffmann et al., 2010; Kang et al., 2004; Stuart et al., 2006; Tong
et al., 2015; Wilson et al., 2012; Wisplinghoff et al., 2004). The complete isolation and species
composition across all sample toothbrush heads are listed in Table S2 (supplementary information)
Occurrence and detection of FLP in toothbrushes
0% 20% 40% 60% 80% 100%
Brevibacterium casei
Pseudomonas extremorientalis
Pseudomonas plecoglossicida
Pseudomonas rhodesiae
Staphylococcus saprophyticus
Staphylococcus capitis
Enterobacter cloacae
Kocuria rhizophila
Pseudomonas libanensis
Staphylococcus hominis
Stenotrophomonas maltophilia
Streptococcus salivarius
Nocardioides sp
Pseudomonas fluorescens
Pseudomonas monteilii
Pseudomonas synxantha
Raoultella ornithinolytica
Staphylococcus epidermidis
Micrococcus luteus
Enterobacter fecalis
Staphylococcus epidermidis
Kocuria palustris
Staphylococcus hominis
Staphylococcus warneri
Nocardioides sp
Micrococcus luteus
Distribution across samples
DB
CB
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FLP were present in 4 out of 14 conventional toothbrush head samples. In the case of designed toothbrush
heads, 2 out of 14 samples were FLP positive. Observations indicated that only one morphogroup i.e.
amoeba was detected among the toothbrush groups examined. Other morphotypes (flagellates and
ciliates) were not detected in suggesting an overall lower FLP diversity on toothbrushes. Fig 4 A-D shows
micrographs of free-living amoeba that were detected in four conventional toothbrush heads (Sample 4,
6, 7 and 14) and Fig 4E & F depicts free-living amoeba detected in two designed toothbrush heads
(Sample 3 and 14).
Figure 4: Micrographs showing amoeboid morphotypes that occurred in different toothbrush samples,
obtained using light microscopy. A, B, C and D depicts the detection of amoeba from conventional
toothbrushes. E and F depict the detection from designed toothbrushes.
Toothbrush design affects bacterial diversity and abundance
Redundancy analysis with toothbrush head design and protozoa occurrence as included factors showed
that the toothbrush head design significantly contributed to explaining variation in bacterial composition
(Table S3 in supplementary information and Fig 5A). No significant relationship between protozoan
occurrence and bacterial community composition was observed.
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60
Figure 5: Redundancy analysis on normalized isolate abundance data showing toothbrush design as
significant constraining variables (arrows). The significance of the model, axes and the factors were
determined by ANOVA (999 permutations, *0.05 ≤ p-values < 0.01; ***0.001 ≤ p-values). Samples from
conventional toothbrush heads are shown in red symbols and from designed heads in blue symbols.
The total aerobic bacterial counts obtained after plating were significantly higher in conventional
toothbrush heads compared to the designed toothbrush heads (two sample t-test, p<0.05; Figure S1,
supplementary information). Samples grouped according to the toothbrush design revealed a significant
impact on bacterial species distribution (ANOVA, p< 0.05). Micrococcus luteus and Kocuria palustris
were the most abundant species in designed toothbrushes. Raoultella ornithiolytica, Enterobacter
cloacae, Pantoae agglomerans, Dermacoccus nishinomiyaensis and various Pseudomonas species were
represented at significantly higher abundance in conventional toothbrushes (Fig 6).
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Figure 6: Heatmap showing the relative percentage abundance of isolates of significant bacterial species
across the conventional (CB) and designed (DB) toothbrushes.
Discussion
The present study revealed the presence of diverse bacterial communities and amoebae on toothbrush
heads. Currently, toothbrushes commonly do not only have bristles but also with projections between
bristle filaments that help in improving oral hygiene (Casemiro et al., 2008). Bacterial load was
significantly higher on conventional toothbrush heads than on toothbrushes with designed projections.
This observation corroborates previous findings that microbes become entrapped within the bristle
filaments and that closely arranged bristles increase bacterial retention (Fig 1) (Bunetel et al., 2000;
Goldschmidt et al., 2004). The genera Pantoea, Brevibacterium, Stenotrophomonas, Streptococcus and
Raoultella were detected on conventional toothbrushes at higher isolation abundance. Species like
Pantoae agglomerans, Streptococcus salivarius, Raoultella ornithiolytica, Brevibacterium casei and
Stenotrophomonas maltophilia have earlier been reported to be associated with the human oral cavity
(Anesti et al., 2005; Derafshi et al., 2017; Dewhirst et al., 2010; Leão-Vasconcelos et al., 2015; Roger et
al., 2011). Micrococcus luteus, Nocardioides sp HKS-04, Staphylococcus hominis and Staphylococcus
epidermis were isolated from both toothbrush designs, showing the ability of these organisms to colonise
any toothbrush surface. Micrococcus and Staphylococcus were also identified on toothbrushes used as
cross-contamination controls, probably due to the fact that both genera are abundant members of the
natural flora found on human skin (Grice & Segre, 2011). As such they can be released into the indoor air
supply (Kooken et al., 2012) and aerosol particles in moist environments (Ankola et al., 2009).
Opportunistic pathogens like Micrococcus luteus has been found associated with oral bacteraemia
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62
especially after brushing (Lucas et al., 2008). In this present study, many identified bacterial isolates
belonged to Nocardioides sp.; their presence could attribute to their ability to utilize polymers (Mattes et
al., 2005) present in toothbrush plastics, as energy source.
Interestingly, protozoan communities are completely dominated by amoeboid morphotype, although only
a few of the toothbrush samples were positive for amoebae. Ciliates and flagellates were not detected.
This may be due to the fact that amoebae have a higher attachment capacity compared to these other
morphotypes (Chavatte et al., 2016). Other reasons include a) the choice of method used to identify FLP.
While the stomacher protocol is a widely used and standardized method (Wu et al., 2003), Chavatte et al.,
2014 reported differences in the recovery of ciliates and amoebae, with stomacher protocol showing
higher recovery of amoebae; b) the culturing conditions could as well present a selective niche for FLP
species as some species do not grow in enrichment medium (Fenchel et al., 1997; Smirnov, 2003); and c)
effective attachment to toothbrush bristle surfaces by free-living amoeba through influent water or from
the oral cavity could contribute to its direct detection (Bergquist, 2009; Dupuy et al., 2014; Thomas &
Ashbolt, 2011). In addition, the toothbrush samples were also completely dried after collection which
could also impact FLP. Free-living amoebae were reported to shift into dormancy under unfavorable
conditions and grow at a constant rate under favorable conditions (Khan et al., 2015). This aspect could
very well contribute to their establishment on and detection from toothbrush heads. Further, identification
of FLP in liquid cultures, in combination with the traditional light microscopy approach has been reported
to underestimate FLP numbers (Caron, 2009; Chavatte et al., 2014) and this might account for their
observed lower abundances and diversity in this study.
Chemical disinfection of toothbrush and sanitation procedures have been assessed by different studies
(Basman et al., 2016; Karibasappa et al., 2011; Mobin et al., 2011; Spolidorio et al., 2011), yet the
microbial contamination of toothbrushes is apparent. Microorganisms grown as biofilms attached to
surfaces display resistance to common disinfectants like chlorine, peracetic acid, hydrogen peroxide,
chlorhexidine and sodium hypochlorite (Bridier et al., 2011). Biofilms are often found to be associated
with grazing protozoa (Arndt et al., 2003; Lawrence & Snyder, 1998) and multispecies biofilms have
shown to provide shelter for bacteria against protozoa grazing (Raghupathi et al., 2018). The presence and
detection of bacteria and amoebae in this study suggest that toothbrushes can provide a habitat for surface
associated microbial biofilms, where FLP and bacteria can interact. Similar results were reported where
FLP positive status of refrigerators was linked to higher bacterial loads (Vaerewijck et al., 2010).
Persistence of pathogenic bacteria in food related habitats were also found to be in association with FLP
(Brown & Barker, 1999). Opportunistic pathogens identified in this study and pre- and post-ingestion
adaptations developed by these bacteria could lead to the rise of grazing resistant strains and as a result,
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are able to survive and grow inside FLP cells that aid in their transmission to new habitats and hosts
(Matz & Kjelleberg, 2005). Moreover, FLP (and their cysts forms) have shown to protect and shelter
pathogenic bacteria against harsh environmental conditions (Barker & Brown, 1994; King et al., 1988;
Lambrecht et al., 2015; Snelling et al., 2006) enhancing their transmission.
In most cases, FLP are not routinely incorporated in microbiological surveys and as a result information
on FLP is scarce (Chavatte et al., 2016). The data obtained in the present study contribute to the growing
knowledge on the occurrence of FLP in human associated environments. It is shown that toothbrushes
also harbor diverse bacterial communities, and their interactions with FLP could constitute to the survival
and transmission of pathogenic bacteria, in general. Further research on the impact of this finding on
bacterial ecology and epidemiology is needed.
Declarations
Consent for publication
All authors read and approved the final manuscript.
Competing interest
The authors declare that they have no conflicts of interest.
Acknowledgements
Our acknowledgements go to all the volunteers who kindly participated and provided the toothbrush
samples. We thank Emma Heyman and Jolien Bonte for their assistance with enumeration, isolation and
identification of bacteria and protozoa.
Funding
This research was funded by BOF Special Research Fund Belgium, 01SF1614 and the Danish Council for
Independent Research grant, DFF-1335-00071. The funding bodies had no influence on the design of the
study and collection, analysis, and interpretation of data and in writing the manuscript.
[MANUSCRIPT 1]
64
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Supplementary Information
Supplementary Table S1: Bacterial load enumerated from the two toothbrush head designs.
Sample Group
Conventional Head Designed head
CFU Log10 CFU/head CFU Log10 CFU/head
1 19000 4.3 25460 4.4
2 940340 6.0 3120 3.5
3 600000 5.8 10420 4.0
4 14650000 7.2 15000 4.2
5 31000000 7.5 100 2.0
6 12200000 7.1 50000 4.7
7 9000 4.0 2630 3.4
8 101000 5.0 110 2.0
9 710000 5.9 432000 5.6
10 20400 4.3 1070 3.0
11 68000 4.8 820 2.9
12 80000 4.9 150 2.2
13 90000 5.0 100 2.0
14 31000000 7.5 600 2.8
Control Group Conventional Head Designed head
CFU Log10 CFU/head CFU Log10 CFU/head
C1 194 2.3 39 1.6
C2 100 2 65 1.8
C3 202 2.3 4 0.6
C4 29 1.5 367 2.6
C5 74 1.87 96 1.98
Supplementary Figure S1: Total aerobic counts from the two toothbrush design groups. The data shows
the mean log10CFU counts ± Std.error of means; *p-value < 0.05.
1.00E+00
1.00E+01
1.00E+02
1.00E+03
1.00E+04
1.00E+05
1.00E+06
1.00E+07
CB DB
Cell
num
bers
/to
oth
bru
sh
*
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Supplementary Table S2: Number of isolates of different bacterial species obtained from conventional and designed toothbrush heads. The species
identified here were based on MALDI-TOF MS with identification scores ≥ 2.0 that is considered reliable for classifying bacteria to the species level.
Sample Conventional Toothbrush Heads (CB) Designed Toothbrush Heads (DB)
Species 1 2 3 4 5 6 7 8 9 10
1
1
1
2
1
3
1
4 1 2 3 4 5 6 7 8 9
1
1
1
2
1
3
1
4
1
5
Acinetobacter johnsonii
2
1 1
2
Aerococcus viridans
1
5 1
7
6
2
6 2
Alcaligenes faecalis
1
Bacillus altitudinis
1
Brevibacterium casei
1 1 1
1 1
2
Brevibacterium celere
2 1
Brevibacterium sanguinis
1
Brevundimonas diminuta
1
1
1
Chryseobacterium
scophthalmum
4
Chryseobacterium sp
1
1
Citrobacter freundii
1
1
Corynebacterium afermentans
2 1
Delftia acidovorans
1
1
1
1
Dermacoccus nishinomiyaensis
1 1
1
2
Enterobacter cloacae
3 1
1
1 9 3
2 4
1
Enterobacter faecalis
2
2
4
24
2
1 1
Enterobacter xiangfangensis
1
Escherichia vulneris
1
Klebsiella oxytoca
4
1
Kocuria carniphila 1
1
1
1
Kocuria kristinae
2
2
Kocuria palustris
7
1
2
13
2 1
4 3
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Kocuria rhizophila
1
5
5 3 4 1 2
1
5 3 9
2
Massilia timonae
1 6
Microbacterium liquefaciens
2
1
3
Microbacterium maritypicum
1
1
Microbacterium oxydans
7
Microbacterium paraoxydans
1
Micrococcus luteus 1 1 2 1 1 3 2
1 1
1 1 2 6 5 13 4 1
1
1
3
1 7
1
2 6 4 6 2
Micrococcus terreus
1
2
Nocardioides sp 2
2
8
1 3 3
1 1 1 2
4
10
1 1 3 3 6
1
Pantoea agglomerans
1
4 1 2
1
Pseudomonas azotoformans
1
2 2 1
Pseudomonas extremorientalis
1 1 2 1
1
Pseudomonas fluorescens 2
1 2 4 1 3
1
1
4
1
Pseudomonas mendocina
1
3
Pseudomonas putida
1
3
1
6
Rothia amarae
1
6
Rothia dentocariosa
1
Serratia liquefaciens
3
4
2
0 2
Pseudomonas graminis
1
1
Pseudomonas libanensis 1 1
9
2
2
1
1
Pseudomonas monteilii 2 1
1 1 2 2
1
7
2
1
Pseudomonas plecoglossicida
1
1
1
1 1
2
Pseudomonas rhodesiae 1
3 1
1
2
Pseudomonas synxantha 1 2
3 3 1 5 1 2
4
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71
Pseudomonas tolaasii
1
1
Raoultella ornithinolytica
1 1 4 2 3
1
5 3 1
6
Raoultella planticola
1
Roseomonas mucosa
1
3
1
Sphingomonas paucimobilis
2
Staphylococcus aureus
3
1
9
Staphylococcus capitis
4 1 2
1
2
1
3
2
1
1
Staphylococcus condimenti
2
4
Staphylococcus cohnii
1
1
1 1
Staphylococcus epidermidis
2
0 1
1 3 4 2 1
2
7 1
1
2
1
3
2
7
Staphylococcus haemolyticus
1
2
1
2 5
Staphylococcus hominis 7
1 1
1
1
0
1 1 3 4 2 1
2
7
Staphylococcus pasteuri
1
2
Staphylococcus pettenkoferi
1
1
Staphylococcus saprophyticus
1
1 1 1
2
1
2
5
Staphylococcus warneri
1
1 1
1
2
16
5 3
1 1
Stenotrophomonas maltophilia
2 5 1
4 4 5 9
1
2
Stenotrophomonas rhizophila
1
Streptococcus gordonii
2
Streptococcus oralis
1
1
Streptococcus parasanguinis
3 2
1
1
5
1
Streptococcus salivarius 2 2
3 2 2 1 5
4 1
Streptococcus vestibularis
5
6
Total No. of Isolates
4
0
7
1
2
0
2
7
5
9
6
5
6
0
4
0
4
2 5
4
9 6
1
6
2
6
8
4
3
3
11
0 6 5
1
4
4
6
1
4
8
2
1
6
2
2
3
3
3
2
1
2
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Supplementary Table S3: Redundancy based analysis on toothbrush design and protozoa occurrence
and its impact on bacterial community. The composition of the bacterial community was significantly
affected by toothbrush head design. df: degrees of freedom; F: ratio of the between groups variance
and within groups variance; p: p-values, *p<0.05.
Factors df
Variance
%
F p
Toothbrush design (CB, DB) 1 5.54 3.13 0.002*
Protozoa (presence/absence) 1 1.90 1.07 0.379
Residuals 25 44.26
73
Raghupathi PK, Zupančič J, Brejnrod AD, Jacquiod S, Houf K, Burmølle M, Gunde-Cimerman N,
Sørensen SJ. Microbiomes in Dishwashers: Analysis of the microbial diversity and putative
opportunistic pathogens in dishwasher biofilm communities. Appl Environ Microbiol. 2018 Jan 12.
pii:AEM.02755-17.Doi:10.1128/AEM.02755-17
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Microbiomes in Dishwashers: Analysis of the microbial diversity and putative opportunistic
pathogens in dishwasher biofilm communities
Prem Krishnan Raghupathi a, cϮ
, Jerneja Zupančič bϮ
, Asker Daniel Brejnrod a, Samuel Jacquiod
a, Kurt
Houf c, Mette Burmølle
a, Nina Gunde-Cimerman
b, Søren J. Sørensen
a
Molecular Microbial Ecology Group, Section of Microbiology, Department of Biology, University of
Copenhagen, Universitetsparken 15, bldg. 1, DK2100 Copenhagen, Denmark a; Department of
Biology, Biotechnical Faculty, University of Ljubljana, Jamnikarjeva 101, 1000 Ljubljana, Sloveniab;
Department of Veterinary Public Health and Food Safety, Faculty of Veterinary Medicine, Ghent
University, Salisburylaan 133, 9820 Merelbeke, Belgium c
Extreme habitats are not only limited to natural environments, but also apply to man-
made systems, for instance household appliances such as dishwashers. Limiting factors, such as
high temperatures, high and low pH, high NaCl concentrations, presence of detergents and
shear force from water during washing cycles define the microbial survival in this extreme
system. Fungal and bacterial diversity in biofilms isolated from rubber seals of 24 different
household dishwashers were investigated using next generation sequencing. Bacterial genera
such as Pseudomonas, Escherichia and Acinetobacter, known to include opportunistic pathogens,
were represented in most samples. The most frequently encountered fungal genera in these
samples belonged to Candida, Cryptococcus and Rhodotorula, also known to include
opportunistic pathogenic representatives. This study showed how specific conditions of the
dishwashers impact the abundance of microbial groups, and investigated on the inter- and intra-
kingdom interactions that shape these biofilms. The age, the usage frequency and hardness of
incoming tap water of dishwashers had significant impact on bacterial and fungal composition.
Representatives of Candida spp. were found at highest prevalence (100%) in all dishwashers and
are assumingly one of the first colonizers in recent dishwashers. Pairwise correlations in tested
microbiome showed that certain bacterial groups co-occur and so did the fungal groups. In
mixed bacterial-fungal biofilms, early adhesion, contact and interactions were vital in the
process of biofilm formation, where mixed complexes of the two, bacteria and fungi, could
provide a preliminary biogenic structure for the establishment of these biofilms.
Ϯ Shared First Author
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Introduction
Extreme natural environments have for decades attracted the interest of microbiologists. However,
only fairly recently have microbial communities in extreme environments in households and common
household appliances been studied. The selection pressure within some of these is reflected in reduced
microbial diversity allowing only the most fitted species to withstand the stressful conditions. In recent
years, in addition to understanding the biodiversity of extreme natural habitats (1–5), ecological
investigations into various man-made ecosystems like kitchens (6), bathrooms (7), trash bins (8), tap
water pipes (9–11), automated teller machines (12), coffee-machines (13), washing machines (14, 15)
and dishwashers (16–19) have gained momentum.
Amongst different household ecosystems studied so far, kitchens are colonized with the broadest
diversity of extremotolerant microorganisms (20–22). And only recently, it was discovered that some
microbes can survive and grow even under extreme conditions in certain domestic appliances (13, 16)
as for instance dishwashers (DWs). DWs are extreme habitats with constantly fluctuating conditions,
where only microbial communities with poly-extremotolerant properties can survive. The individual
community members, as well as the whole community itself must possess key phenotypic traits that
enable them to resist alternating wet and dry periods, frequent changes of temperatures during the
washing cycles (from 20°C and up to 74°C), oxidative detergents elevating the pH from 6.5 to 12, high
organic loads, high NaCl concentrations and shearing generated by water sprinklers. The metal, plastic
and rubber parts of DWs may enable the establishment and growth of mixed bacterial/fungal
communities that are protected by copious amounts of extracellular polymeric substances (EPS) and
thus, confer on the biofilm communities, the extremotolerant properties that go beyond the
extremotolerance of each individual species (17, 18, 23).
The obvious choice for microbes when exposed with extreme conditions in DWs is the biofilm mode
of growth, providing shelter against external stresses (24) and where intimate cross-species boundaries
may occur (25). Such surface communities (26, 27) may also provide a link to emerging disease
pathogenesis (28), since biofilms formed in DWs (and other appliances) could contribute to the
dispersion and persistence of bacterial/fungal groups outside the common spectrum of saprobes (16).
Fungi, able to cause opportunistic infections in humans, have been documented inside DWs (17–19)
and the incidence of domestically-sourced fungal infections has been increasing steadily over the last
decades (17, 29, 30).
However, to date, the diversity of the whole microbiota in DWs has not been investigated. Therefore,
we have studied both the bacterial and fungal communities (with a special focus on mixed biofilms)
associated with household DWs, using high-throughput sequencing. Furthermore, we studied how
specific conditions of the DWs impact the abundance of certain microbial groups and how well inter-
and intra-kingdom interactions shape the structure of microbial communities within DWs.
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Materials and methods
Dishwasher Sample Information
Microbial biofilm grown on the rubber seals of 24 different DWs in private dwellings across different
Slovenian cities was sampled in this study (Table 1). The water supply at source of these DWs was
characterized into soft or hard water based on ion analysis method as previously performed (10). Final
concentrations were determined following the method from ISO Standard SIST EN ISO 11885:2009.
Biofilm sampling and genomic DNA extraction
Biofilm formed on up to 1 cm2 of the rubber seal surface (Fig 1) was scraped off using sterile scalpel
and the collected biomass was placed into a sterile tube and stored at -20°C till use. Genomic DNA
extraction from 50 – 100 mg of biofilm biomass was performed using MoBio Power Biofilm DNA
isolation Kit (Carlsbad, CA, USA) according to the manufacturer’s instructions. Extraction controls
were included during DNA extraction. Negative control contained ultrapure water (Milli-Q) in the
same quantity as dishwasher biofilm biomass. The processing of the sample and the negative control
were performed simultaneously and in the same way, according to manufacturer’s instructions. DNA
concentrations were quantified for all samples using Qubit® dsDNA HS Assay and measuring the
fluorescence on Qubit® fluorometer (InvitrogenTM, UK).
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Table 1: Different dishwasher machines and its associated characteristics from which biofilm samples were obtained. ‘Ticks’ represent the samples that were
sequenced based on 16S rRNA genes /ITS rRNA region. S1-S24: 24 sampled dishwashers; 16S rRNA: gene for 16S rRNA, partial sequence obtained within
sample; ITS rRNA: ITS rRNA gene partial sequence obtained within sample; Age: years used; Freq. of use: Frequency of use i.e. the no. of times the DW
was used per week; Washing cycle: Temperature of washing cycle; Water hardness characteristics: H – hard (above 2.0 mmol/L CaCO3); MH –
moderately hard (1.5- 2 mmol/L CaCO3); SH – slightly hard (1.0 -1.5 mmol/L CaCO3); MS- moderately soft (0.5 -1 mmol/L CaCO3); S- soft (below 0.5
mmol/L CaCO3).
Sample S1 S2 S3 S4 S5 S6 S7 S8 S9 S10 S1
1
S1
2
S1
3
S1
4
S1
5
S16 S17 S18 S1
9
S2
0
S2
1
S2
2
S2
3
S2
4
16S
rRNA
ITS
rRNA
Years
used
2.
5
2.
5
7 3 2 8 5 8 5 1 7 0.5 1 2 8 1 1 4 4 3 8 5 1 8
Freq. of
use
7 7 3 14 14 7 3 7 3 2 7 3 3 2 7 7 14 3 3 1 7 7 1 7
Washing
cycle
(°C)
60 60 60 60 60 60 60 60 60 65 60 65 50 65 65 65 65 65 65 60 50 60 65 60
Water
Hardnes
s
S
H
S
H
M
H
M
H
M
H
M
H
M
H
M
H
M
H
M
H
SH SH SH SH MS M
H
M
H
M
H
H SH MS H SH MS
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16S rRNA gene and nuclear ribosomal internal transcribed spacer (ITS) rRNA amplicon based
sequencing
To determine the microbial diversity in biofilms associated with the selected DW rubber seals,
amplicon sequencing based on 16S rRNA gene and nuclear ribosomal internal transcribed spacer
(ITS) region for bacterial and fungal diversity, respectively, was applied. The concentration of
extracted DNA was quantified and adjusted to 5 ng /µl for all samples using ultrapure water (Milli-Q).
For the PCR reaction, 1 µl of above prepared DNA was used. The variable regions V3 and V4 were
used for bacterial identification by primers targeting the flanking conserved regions and amplified
using the primers PRK341F (5’-CCT AYG GGR BGC ASCAG-3’) and MPRK806R (5’-
GGATCTACNNGGTATSTAAT-3’) (70). The general eukaryote primers ITS7 (5’-
GTGAATCATCGAATCTTTG-3’) (71) and ITS4 (5’-CAGACTTRTAYATGGTCCAG-3’) (72) were
used to amplify the ITS2 region for sequencing. Blank was included as a negative control. PCR
amplifications were done in two steps.
The first PCR amplification was done using PuRe Taq ready-To-Go PCR Beads (GE Healthcare,
United Kingdom) containing 1µl of each primer. Bacterial PCR-I mix was amplified according to
following conditions: 94°C for 2 min, 35 cycles of 94°C for 20 s, 56°C for 20s and 68°C for 30s, and
final extension at 68°C for 5 min. Eukaryotic PCR-I amplifications were 94°C for 2 min, 35 cycles of
94°C for 30 s, 56°C for 30 s, 72°C for 30 s, followed by 72°C for 5 min. The final products were then
cooled on ice to minimize hybridization between specific PCR products and short nonspecific
amplicons. Products were checked by running 5 μl on a 1.5% agarose gel. Sequencing primers and
adaptors were added to the amplicon products in the second PCR step: 2.0 µl 10x AccuPrime™ PCR
Buffer containing 15 mM MgCl2, Invitrogen), 0.15 µl (MSM2) AccuPrime™ Taq DNA Polymerase (2
units/µl, Life Technologies), 1.0 µl of each fusion primers, 2 µl of 10X diluted PCR product from first
PCR and water to a total of 20 µl reaction volume. The PCR-II conditions were: 94°C for 2 min,
followed by 15 cycles of 94°C for 30 s, 56°C for 30s and 68°C for 30s, and final extension at 68°C for
5 min. Amplicons were size separated on a 1 % agarose gel and purified using Montage Gel Extraction
Kit (Millipore, Billerica, MA, USA). Amplicon concentrations were quantified for all samples using
Qubit® dsDNA HS Assay and measuring the fluorescence on Qubit® fluorometer (InvitrogenTM,
UK). Samples were sequenced using an Illumina MiSeq sequencer, employing paired-end reads, as
described previously (73). De-multiplexing was performed by the Miseq Controller Software. Raw
fastq files for both 16S and ITS data were processed with qiime_pipe, a wrapper around the QIIME
(1.7) pipeline available at https://github.com/maasha/qiime_pipe. The preprocess_illumina.rb script
handles quality control, and for both datasets this was done using the same parameters. Merging of
paired-end reads was done with a maximum of 20% mismatches and minimum length overlaps of
15bp. Primers were identified with 2 maximum mismatches and the amplicon was trimmed to only
contain the sequence between the primers. Sequences were discarded if the average quality was less
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79
than 30. Chimera checks were performed with the QIIME script identify_chimeric_seqs.py using the
usearch61 method. For 16S, it was checked against GreenGenes (4 Feb. 2011) and for ITS, against
UNITE (09 Feb 2014 dynamic version) databases. OTU picking were done at 97% for both datasets
with QIIMEs pick_otu.py, and representatives of these were picked with pick_rep_set.py, both at
default settings. For 16S, trees were generated by aligning to the GreenGenes set using PyNast and
FastTree through the QIIME wrappers. Representative sequences were classified through the RDP
classifier at the default 0.8 confidence threshold. The same databases were used for classification as
well as for chimera checking.
Microbial Interaction Network
Positive correlations signifying co-occurrence and negative correlations signifying mutual exclusions
were characterized by generating the Spearman co-occurrence network. The network based on
normalized abundance was generated using the CoNet 1.0b6 plugin for Cytoscape 3.2.1 on the basis of
the in-built nonparametric Spearman correlation coefficient with a minimal cut-off threshold of r ≥
|0.65| (P≪0.01, Bonferroni corrected) (74, 75). In this study, we present the correlation data for
bacterial and fungal members that were coexisting in the same dishwashers (N=18). OTUs with ≥50%
sample representation were selected for the microbial network (n≥9, N=18).
Data Analysis
Alpha diversity analyses were performed in the PAST software ver.2.17 (76). Alpha diversity indices
were calculated on OTU counts rarefied to 4000 counts per sample for bacterial sequences and
samples below 4000 bacterial counts were not included in this analysis. Fungal counts were rarefied to
400 counts per sample for this particular analysis. Fungal sequences rarefied to the lowest sequencing
depth allowed DW samples with replicate conditions to be maintained. The following indices were
used to assess the diversity: the sample richness, the Shannon (H), the Chao-1. The effect of DW
conditions on alpha diversity indices were statistically assessed using t-test (Wilcoxon-Mann-Whitney,
p< 0.05).
Multivariate beta-diversity analyses were done using non-rarefied counts (77). As the contingency
tables featured 1000-folds variation in abundance, a log10 transformation was applied to have all the
information possible and to satisfy distributional assumptions. The transformed compositional dataset
was subject to a Redundancy analysis (RDA) using DW conditions as factors. The significance of the
model, of the RDA axes and factors were estimated by ANOVA on Euclidean distances and 999
permutations, p<0.05 significance. In addition, PERMANOVA using 999 permutations and Bray-
Curtis dissimilarity index was performed to assess the significance of DW conditions. Selections of
taxa (phyla and genus level) with significant changes in prevalence grouped by DW conditions were
performed in STAMP 2.1.3 (78) using multiple and group comparisons and significance checked
using in-built ANOVA (Tukey-Kramer post-hoc tests) and Welch’s t- test (Bonferroni corrected). The
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selected significant taxa were plotted in heatmap using dissimilarity indices computed based on
euclidean distances, average clustering and scaled counts. The heat maps were clustered based on
‘coniss’. RDA, PERMANOVA, principal component (PC) plots and heatmaps were generated using
various R packages: gplots, vegan, rioja and Rcolorbrewer available for Rgui 3.2.0 (79).
Results
Characterization of bacterial and fungal communities in DW-associated biofilms
Twenty-four biofilms grown on rubber seals of DWs (Fig 1) were sampled to assess both bacterial and
fungal communities. DWs characteristics varied in terms of years, frequency of use, temperature of
washing cycles and influent water hardness (WH) as shown in Table 1. In the first PCR reactions, 21
samples generated DNA fragments that were processed by a second PCR and further sequenced. A
total of 221, 032 partial 16S rRNA gene and 313, 420 ITS rRNA gene transcript sequences were
obtained from 21 DW samples, respectively. Raw sequences of all the DWs are available from the
NCBI Sequence Read Archive (SRA) under the Bioproject IDs: PRJNA315977 for bacterial and
PRJNA317625 for fungal reads, respectively. Overall, sequence reads were assigned to 309 bacterial
OTUs and 194 fungal unique OTU classifications. The predicted number of bacterial genera ranged
from 29 to 150 across all samples and the fungal genera ranged from 15 to 104 (Supplementary
information, Table S1).
Figure 1: Biofilm formed on the rubber seal in residential DWs. Microbial biofilm formation on DW
rubber seal, the square (in red) represents the 1cm2 sampling site. Biofilm samples from 1 cm2 were
collected by scrapping the surface of rubber seal with sterile scalpel for DNA extraction and further
analysis. Sampling was done in-situ, with the seal at its original place.
Alpha diversity were calculated based on rarefied sequences and are presented (Supplementary
information, Table S2). Richness and evenness of the bacterial community were not affected by the
DW conditions (Supplementary information, Table S3). However, the alpha diversity indices of fungal
community were found to be significantly influenced by DW conditions. The years of usage and DW
with influent hard water had significant impact on species richness, species evenness and abundance
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81
indicating that these specific environmental conditions enrich fungal community profiles (Wilcoxon-
Mann-Whitney tests, p < 0.05) (Supplementary information, Table S4).
Microbial phyla in DW associated biofilms
DWs biofilms were composed of diverse fungal and bacterial phyla. Based on non-rarefied 16S rRNA
gene reads, the sequences were assigned to 16 different bacterial phyla, of which, Proteobacteria were
dominating across all the samples followed by Actinobacteria and Firmicutes (Supplementary
information, Fig S1). All DW samples also contained sequences representing Bacteroidetes. The
remaining phyla, (Chloroflexi, Cyanobacteria, Deinococcus-Thermus, Spirochaetes, TM7,
Verrucomicrobia, Synergistetes and Acidobacteria) contributed up to 9% of the total prokaryotic
sequences. Among the subclasses of Proteobacteria, the bacterial community was dominated by α-
Proteobacteria (46 ± 7 %) and γ-Proteobacteria (45 ± 7 %) in all the DW biofilm samples. Bacilli were
the most abundant subclass of Firmicutes dominating in all biofilm samples. The most abundant taxa
in all DNA samples belonged to the genera Gordonia, Wautersiella, Rhodobacter, Nesterenkonia,
Stenotrophomonas, Exiguobacterium, Acinetobacter and Pseudomonas. Bacterial OTU represented by
the genus Meiothermus belonging to the phylum Deinococcus-Thermus and the phyla TM7 were
present in 19/21 DW samples. The genera Escherichia/Shigella and Pseudomonas were identified in
62% and 67% of DWs, respectively (Fig 2).
OTUs based on non-rarefied ITS rRNA gene reads were assigned across four fungal phyla
(Supplementary information, Fig S1). Ascomycota dominated in the samples followed by
Basidiomycota. Among the subclasses of Ascomycota, the fungal biofilm community was dominated
by Saccharomycetes characterized by genera Candida, Debaryomyces, and Saccharomyces (Fig 2)
present in all DWs. Filamentous fungal genera like Cladosporium, Fusarium and Aspergillus were
present in more than 16 DW samples, and the black yeast genera, Aureobasidium and Exophiala, were
present in 33% of DWs. Basidiomycota represented by genera Rhodotorula and Cryptococcus were
present in 90% and 86% of DWs, respectively. The genera Wallemia and Trichosporon were present
in more than half of DWs. Microbiota based on absolute sample count i.e. the bacterial and fungal taxa
classified at the genus level that occurred in N > 5 samples, where 21 ≥ N ≥ 5 is shown in Fig 2.
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Figure 2: The different bacterial and fungal genera that were represented across 21 DW samples. The numbers represent sample count i.e. the number of DW
samples that contained the representative genera.
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83
Abiotic conditions of DW affect microbial composition
Changes in the structure of microbial communities were investigated by multivariate beta-diversity
analysis. Redundancy analyses (RDA) were performed to test the relationship between different DW
conditions and their impact on the microbial community composition. RDA with two included factors
(years of use and frequency of use) showed that the DWs years of use was the most significant driving
force (ANOVA, p<0.05) affecting bacterial communities followed by frequency of use (Freq). RDA
explained 84% total variance, of which 43.8 % is described by RDA1 and RDA2 (respectively,
27.32% and 16.44%, Fig 3A & B). The most explanatory variables were frequency of use (27%) and
the number of years in use (23%) (Supplementary information, Table S5). RDA with three included
factors (year, water hardness and frequency of use) showed that years of use, frequency of use and
WH significantly affected fungal community (ANOVA, p<0.05). Out of 38% total variance, 23.5%
was explained by the first two components (respectively, 15.43% and 8.08%, Fig 4A, B&C). In this
case, WH was found to be the most explanatory variable (18.9%) (Table S5, Supplementary
information). PERMANOVA on Bray-Curtis dissimilarity index confirmed the observed trends where
DWs frequency of use (14%) and WH (17%) were the most significant factors (p<0.05) impacting the
bacterial and fungal community profiles, respectively (Supplementary information, Table S6).
Figure 3: Principal component plots of redundancy analysis (RDA) performed to log10-transformed
16S rRNA amplicon sequencing data using (A) year and (B) frequency of use as explanatory factors.
Significance of the model, axes and factors was determined by ANOVA (999 permutations; p < 0.05).
Stars stand for the level of significance according to the code: (*) 0.05 ≤ P-values < 0.01; (**) 0.01 ≤
P-values< 0.001. Factors ‘y’ represent years of use (0-4, 5-7 and 8 years) and ‘Freq’ represents
frequency of use (1-3, 7 and 14 times/week).
RD scatter-plot shows DW with bacterial composition grouped into recent DW used between 0-4; old
DWs at 5-7/8 years (Fig 3A). Based on frequency of use, DWs were grouped under three categories:
low frequency (1-3 times / week), intermediate frequency (7 times /week) and high frequency (14
-6 -4 -2 0 2 4 6
-4-2
02
RDA1 ** - 27.32%
RD
A2
*-
16
.44
%
S24
S17
S22
S19
S6
S15
S21
S13
S1S2
S10
S4
S14
S7
S3
S23
S5
S12
S11
Year 0-4
Year 5-7Year 8
Model P= 0.04*
Variance = 84%
8 5-7
0-4
A
S21
S16
-6 -4 -2 0 2 4 6
-4-2
02
RDA1 ** - 27.32%
RD
A2
*-
16
.44
%
Freq ≥14
Freq 1-3Freq 7
S24
S17
S22
S19
S6
S15
S21
S13
S1S2
S10
S4
S14
S7
S3
S23
S5
S12
S11
Model P= 0.04*
Variance = 84%
≥ 14 7
1-3
B
S21
S16
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84
times /week) (Fig 3B). Samples with fungal composition were also grouped based on DW conditions
(year, frequency of use and influent WH) as shown in Fig 4A, B & C, respectively.
Figure 4: Principal component plots of redundancy analysis (RDA) performed to log10-transformed
ITS rRNA amplicon sequencing data using (A) year and (B) frequency of use and (C) water hardness
as explanatory factors. Significance of the model, axes and factors was determined by ANOVA (999
permutations; p < 0.05). Stars stand for the level of significance according to the code: (*) 0.05 ≤ P-
values < 0.01; (**) 0.01 ≤ P-values< 0.001. Factors ‘y’ represent years of use (0-4, 5-7 and 8 years);
‘Freq’ represents frequency of use (1-3, 7 and 14 times/week) and ‘H’ represents hard water, ‘MH’
represents moderately hard water, ‘SH’ represents slightly hard water and ‘MS’ represents moderately
soft water.
The influence of DW’s age and frequency of use on bacterial communities
-6 -4 -2 0 2 4
-3-2
-10
12
3
RDA1** - 15.43%
RD
A2
**
-8
.08
%
Year 0-4
Year 5-7
Year 8
S19
S22
S3
S18 S12
S16
S24
S11
S13
S7
S21
S4
S9
S6
S1
S23S2S8
S17S15
S20
Model P= 0.004**
Variance = 38%
-6
8 5-7
0-4
A
-6 -4 -2 0 2 4
-3-2
-10
12
3
RDA1** - 15.43%
RD
A2
**
-8
.08
%
Freq 1-6 Freq 7-14
S19
S22
S3
S18 S12
S16
S24
S11
S13
S7
S21
S4
S9
S6
S1
S23S2S8
S17S15
S20
Model P= 0.004**
Variance = 38%
-6
7-14 1- 6
B
-6 -4 -2 0 2 4
-3-2
-10
12
3
RDA1** - 15.43%
RD
A2
**
-8
.08
%
H
S19
S22
S3
S18 S12
S16
S24
S11
S13
S7
S21
S4
S9
S6
S1
S23S2S8
S17S15
S20
Model P= 0.004**
Variance = 38%
-6
H MH
MS SH
SH
MS
MH
C
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Based on the above results, further analyses of DW groups on taxonomic profiles were performed in
the software package STAMP. Actinobacteria, Firmicutes and Proteobacteria were the three major
phyla significantly affected by DW’s age (ANOVA, p< 0.05) and Firmicutes further influenced by
frequency of use (ANOVA, p< 0.05). Recent DWs (0-4 years) contained Proteobacteria (48%),
Actinobacteria (29%) and Firmicutes (13%) indicating that these phyla members were the early
settlers in biofilms of recent DWs. DWs between 5-7 years of age seem later to be increasingly
populated by Actinobacteria (36%) and Firmicutes (34%). DWs at 8 years of age showed increased
levels of Actinobacteria (49%) with further reduction in Proteobacteria and Firmicutes (Fig 5A). DWs
that were used more often (frequency) had reductional shift in Firmicutes diversity (Fig 5B). Thus,
bacterial members belonging to Actinobacteria and Proteobacteria groups could dominate in DWs and
Firmicutes being susceptible to operational conditions of DWs.
Figure 5: Impact of DWs age and frequency of use on the relative abundance of bacterial taxa present
in the samples. (A & B) Mean ± S.E.M of percentage abundance in samples grouped by (A) years and
(B) frequency of use at the phyla level. (C & D) Heat map of significant bacterial genera in DW
samples grouped by (C) years of use (0-4, 5-7 and 8 years) and grouped by (D) frequency of use (1-3,
7 and 14times/week), respectively.
A B
% R
ela
tive
Ab
un
da
nce
% R
ela
tive
Ab
un
da
nce ▪
●
Proteobacteria
Firmicutes
Actinobacteria
0
10
20
30
40
50
60
0-4 years 5-7 years 8 years0
10
20
30
40
50
60
1-3 7 ≥14
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Investigation on abundance shift patterns at genus level between different DW age groups is shown in
Fig 5C. A total of 11 bacterial taxa showed significant prevalence levels according to the three DW
age groups (ANOVA, p< 0.05). Most young DWs (0-4 years) had varying levels of bacterial
abundance represented by genera Rhizobium, Escherichia/Shigella, Azospira, Exiguobacterium,
Rhizobium
Escherichia/Shigella
Azospira
Exiguobacterium
Chryseobacterium
Staphylococcus
Arthrobacter
Treponema
Aerococcus
Brevibacterium
Starkeya
Aeromicrobium
Kaistella
Ancylobacter
Salana
-3 -1 1 3
Relative abundance0- 4
5- 78
C
S1
S1
0
S1
2
S1
3
S1
4
S1
7
S1
9
S2
S2
3
S4
S5
S1
1
S2
2
S3
S7
S1
5
S2
1
S2
4
S6
S2
0
S1
6
-3 -1 1 3
Relative abundance
S2
3
S1
0
S1
2
S1
3
S1
4
S2
0
S1
9
S3
S1
S1
1
S1
5
S1
6
S2
S2
1
S2
2
S2
4
S6
S1
7
S5
S7
S4
1- 3
7
14
D
Cellulosimicrobium
Helicobacter
Brevibacterium
Nesterenkonia
Xanthobacter
Roseomonas
Aeromicrobium
Micrococcus
Leptothrix
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Chryseobacterium and Staphylococcus. Taxa belonging to genera Exiguobacterium, Arthrobacter,
Staphylococcus, Aerococcus, Treponema and Lactobacillus were represented in DWs 5-7 years.
Further, genera like Aeromicrobium, Salana, Ancylobacter, Starkeya, Kaistella, Brevibacterium and
Ancylobacter dominated in DWs used for 8 years. Genera Azospira, Escherichia/Shigella and
Rhizobium dominated in younger DWs (Welch’s t-test, p< 0.05) whereas, genera Aeromicrobium and
Brevibacterium dominated in older DWs used for 5-7 and 8 years (Welch’s t-test, p< 0.05,
respectively). The frequency of use showed 9 bacterial taxa to be significantly influenced (ANOVA,
p<0.05) between the three levels (Fig 5D). The genera Brevibacterium, Aeromicrobium, Roseomonas,
Xanthobacter, Helicobacter and Cellulosimicrobium were prevalent in DW used more frequently
compared to DWs used at low frequencies (1-3times /week) (Welch’s t- test, p < 0.05).
Fungal community of DWs rubber seal is influenced by age, frequency of use and hardness of
incoming tap water
The conditions of DWs and their impact on fungal taxonomic profiles revealed a significant difference
between the three fungal phyla, Ascomycota and Basidiomycota (ANOVA, p<0.05). Young
dishwashers (0-4 years) were abundant in Ascomycota (92%) with Basidiomycota contributing only
5%. Over time, the two phyla became more equally represented at 55% and 43%, respectively (Fig
6A). A higher shift in abundance between the phyla, Ascomycota and Basidiomycota was observed
under two frequency levels (Welch’s t-test, p< 0.05, respectively) (Fig 6B). This could indicate
Basidiomycota susceptible to decline in DWs used more frequently while ascomycetous fungi remain
well established. The impacts of WH were shown to affect the phyla Ascomycota, Basidiomycota and
Zygomycota between the four levels of water harness (ANOVA, p< 0.05). Hard and moderately hard
water influenced fungal genera belonging to Basidiomycota and Zygomycota compared to soft and
slightly hard water (Welch’s t-tests, p< 0.05) (Fig 6C).
Figure 6: Impact of DWs age and frequency of use on the relative abundance of fungal taxa present in
the samples. (A & B) Mean ± S.E.M of percentage abundance in samples grouped by (A) years, (B)
0
20
40
60
80
100
0-4 years 5-7 years 8 years
%R
ela
tive
ab
un
da
nce
0
20
40
60
80
100
1-6 7-14
%R
ela
tive
ab
un
da
nce
0
20
40
60
80
100
H MH MS SH
%R
ela
tive
ab
un
da
nce
A B C
▪●
Ascomycota
Basidiomycota
Zygomycota
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frequency of use and (C) water hardness at the phyla level. (D, E & F) Heat map of significant fungal
genera in DW samples grouped by (D) years of use (0-4, 5-7 and 8 years); (E) grouped by frequency
of use and (F) grouped by influent water hardness, respectively. ‘H’ represents hard water, ‘MH’
represents moderately hard water, ‘SH’ represents slightly hard water and ‘MS’ represents moderately
soft water.
At genus level, 6 taxa were influenced on abundance levels by years of use (Fig 6D). Most DWs used
between 5-7 years were shown to be colonized by the genera Wallemia, Rhodotorula, Candida,
Aureobasidium and Cryptococcus. Though recent DWs had varying fungal representations between
these genera; Candida was significantly abundant in recent DWs (0-4 years) and Rhodotorula was
significantly higher in abundance in DWs used for 5-7 years compared to DWs at 8 years (Welch’s t-
test, p<0.05), respectively. DWs frequency of use had significant impact on the fungal genera Candida
and Rhodotorula where, most frequently used DWs enriched Candida and less frequently used DWs
were settled with Rhodotorula (Fig 6E) (Welch’s t- test, p<0.05). The DW samples with incoming WH
also influenced the fungal biota as shown in Fig 6F. Genera Phoma, Thelebolus, Stagonaspora,
Neobulgaria, Perisporiopsis Cladosporium were represented significantly at higher abundance in
samples with hard water compared to samples with other WH characteristics (Welch’s t- test, p< 0.05).
Cryptococcus
Hyphodontia
Aureobasidium
Candida
Rhodotorula
Wallemia
-2 -1 0 1 2
Relative abundance
0- 4
5- 7
8
S1
S1
2
S1
3
S1
6
S1
7
S1
8
S1
9 S2
S2
0
S3
S7
S4
S9
S1
5
S2
1
S2
4
S6
S8
S2
2 S3
S1
1
D
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Positive microbial interactions may shape the DWs biofilm communities
Microbes sharing the same environmental niche may co-exist or exclude each other while competing
for the same resources (40–42). Correlation analyses were performed to explore relationships among
the microbial flora associated with DW’s biofilm communities, as positive pairwise correlations may
indicate microbial collaboration or dependencies. The survey revealed 140 significant interactions at
the genus level in biofilm samples including both fungal and bacterial genera (Spearman rank
correlation cut-off, r > |0.65|, permutation test, p<0.05) with 90% (125/140) of the predicted
interactions positively correlated (Table 2).
S1
2
S1
3
S1
8
S1
9
S2
0
S2
3
S2
4
S3
S7
S9
S1
S1
1
S1
5
S1
6
S1
7
S2
S2
1
S2
2
S4
S6
S8
Rhodotorula
Candida
1- 6
7- 14
-2 -1 0 1 2
Relative abundance
E
Debaryomyces
Cladosporium
Rhodotorula
Sterigmatomyces
Stagonospora
Thelebolus
Neobulgaria
Perisporiopsis
Mortierella
Cryptococcus
Phoma
Alternaria
Trichosporon
Pilidium
Articulospora
Ascochyta
Dioszegia
Clitopilus
Rhizophagus
Gibberella
Hannaella
Candida
-4 -2 0 2 4
Relative abundance
H
MH
MS
SH
S1
9
S2
2
S1
6
S1
7
S1
8
S3
S4
S6
S7
S8
S9
S1
5
S2
1
S1
S2
4
S1
2
S2
S2
0
S2
3
S1
3
S1
1
F
[MANUSCRIPT 2]
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Table 2: Pair-wise correlations between the bacterial and fungal OTUs observed in 18 DW biofilms
samples. Interactions shown here are based on Spearman Rank correlations (r > cutoff=|0.65|,
permutation test, p<0.05).
Surprisingly, 95% (119/125) of the predicted positive correlations were intra-kingdom bacterial
interactions dominated by Proteobacteria, Actinobacteria and Firmicutes. Representatives of genera
detected in these biofilms were shown to positively correlate with other taxa. For example, γ-
Proteobacteria genus Escherichia/Shigella spp. was positively correlated with Ochrobactrum spp.;
Staphylococcus spp. was positively correlated with γ-Proteobacteria genus Pseudomonas.
Enterococcus spp. was positively correlated with the γ-Proteobacteria genus Stenotrophomonas.
Positive inter-kingdom correlations i.e. interaction between bacterial and fungal members were
observed only in 2.5% of total positive interactions (N=125). Example includes Saccharomycetes
(Candida spp.) and α-Proteobacteria in one case and between β–Proteobacteria and Dothideomycetes
in two cases. Fungal members tend to mutually co-occur in DWs community (2.5% (3/125)) as no
negative correlations within the fungal taxa were observed. In fungi, positive co-occurrence was
observed between Rhodotorula spp. and Cladosporium spp.; Cryptococcus spp. and Cladosporium
spp.; Debaryomyces spp. and yeasts classified as Saccharomycetes. However, mutual exclusions
among inter-kingdom interactions accounted for 46% (7/15) of the total negative correlations (N=15).
Cross-domain negative interactions i.e. mutual exclusions between bacteria and fungi were observed
between the fungal phylum Ascomycota and the bacterial phyla Actinobacteria and Bacteroidetes.
Basidiomycota negatively correlated with γ–Proteobacteria; α-Proteobacteria and Sphingobacteria
(Supplementary information, Fig S2).
Discussion
Indoor and household environments, including household appliances, offer diverse habitats for
microorganisms to adapt and flourish. Most current knowledge on DWs microbiology is focussed on
opportunistic pathogenic black yeasts and other members of DWs mycobiota based on classical
cultivation techniques (16, 17, 19). To date, the molecular approach to identify and characterize the
microbial diversity of DWs was documented in a single study that was limited to the presence of fungi
Total interactions 140
Total Positive Interactions 125
Bacterial -Bacterial positive interactions 119
Fungal-Fungal positive Interactions 3
Bacterial-Fungal positive Interactions 3
Total Negative Interactions 15
Bacterial -Bacterial negative interactions 8
Bacterial-Fungal negative Interactions 7
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in new and old biofilms (17). This study, using high throughput sequencing, assessed both bacterial
and fungal community coexistence in biofilms, which were developed on the rubber seals of DWs.
These mixed bacterial/fungal biofilms are very likely to enable protection against harsh environmental
conditions contributing to persistence. The formation of biofilms on DWs rubber seals also reflects
invasion, settlement and growth of microorganisms under extreme conditions in these appliances.
Combination of different stress factors within DWs allows for the survival of only well adapted and
complementary microbial species that enable the formation of biofilms. We found that on rubber seals
in DWs, most abundant OTUs were dominated by Gram-positive members like Gordonia spp.,
Micrococcus spp., Exiguobacterium spp and a Gram- negative Chryseobacterium spp. The presence of
these genera are usually associated with natural environments in which biotic conditions are extreme
(31–36). Few species within these genera were earlier reported as halotolerant and tolerate a broad
range of pH (5-11), high levels of UV radiation and heavy metal stress (including arsenic) (31–33).
The most abundant bacterial genus Gordonia was found in all DWs sampled in this study. These
representatives were reported to degrade different polymers and xenobiotics (35) that presumably
facilitate their presence on DW rubber seals. Bacterial taxa represented by putative thermophilic
genus, Meiothermus and phylum TM7 were present in most DW samples. Both of these thermophilic
genera could be expected in biofilms formed on DW rubber seals, since the bacterial members of
these genera can tolerate short periods of up to 70 °C and grow optimally from 50 to 65°C and at
alkaline conditions (pH ~8.0) (36). This study reports the detection of putative Meiothermus spp.
within a domestic system.
Microbial diversity within indoor environments is influenced by human effects. The abundance of
bacterial composition of indoor environment was shown to closely mirror the microbial profiles of its
human residents (37). Sampled DWs included a subset of bacterial genera known to have
representatives associated with humans, for example Staphylococcus, Streptococcus, Lactobacillus,
Corynebacterium and Enterococcus. These bacterial genera, common on human skin and in the gut,
have been detected in other studies investigating the domestic microbiome (30, 38, 39), yet their
presence within the extreme conditions has not been reported. Furthermore, our study revealed the
presence of sequences affiliated to genera known to harbor some of the most common and potential
human opportunistic pathogens, namely Escherichia/Shigella and Pseudomonas as integral members
of DWs microbial biofilms. In fact, more than 60% of samples contained genera like Acinetobacter,
Escherichia/Shigella and Pseudomonas, indicating that DWs could shelter these bacterial groups in
private homes. However, it should be noted that the presence of potential bacterial pathogens in
household DW can be tempered by the fact that pathogenicity can be strain specific (the methods used
here do not provide such resolution). In addition, the approach applied in our study cannot distinguish
between cells that were alive/dead/spores. Multispecies biofilm formation may help these well-adapted
(and other) opportunistic bacteria to survive in harsh environment of DWs, being protected with large
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amounts of bacterial and fungal EPS protecting them in immediate close vicinity (40, 41). EPS
immobilizes individual cells in biofilms and keep them in close proximity, allowing interactions
including cell-cell communication and the formation of synergistic micro-consortia (40) and thereby,
determine the development and structure of multi-species biofilms (42). The primary colonizers of any
surface are predominantly bacteria (43), which modify surface characteristics, enabling subsequent
colonization by secondary microorganisms (44, 45). Recent studies of microbial biofilm communities
on natural and artificial substrates (ceramic, glass, plastic, aluminium, and coral skeleton) showed that
early-stage biofilms were established by successive colonization of Proteobacteria, Firmicutes and
Actinobacteria (46–48). Similarly, in this study, it was observed that in recent dishwashers, the main
biofilm community members belonged to Proteobacteria, Actinobacteria and Firmicutes. α-
Proteobacteria that prevail in aquatic-related biofilms (49) were well represented besides γ-
Proteobacteria in DWs. γ-Proteobacteria were found to be the major contributors to biofouling (49, 50)
formed on polymers (51) and their presence in DWs could constitute to different polymers used in DW
components.
Taxonomic identification of the fungal DW community by gene marker-based amplicon sequencing
showed similar results to previous cultivation based approaches (16, 17, 19). The prevalence of
Ascomycota in relation to Basidiomycota was confirmed (17). However, in this study, we report
differences in distribution at the genus level. Sequencing results from fungi in DWs biofilms from a
previous study showed an higher abundance of Cryptococcus (17), while in this study the dominance
of Candida is reported. Also, the prevalence of fungi belonging to genera Rhodotorula, Cryptococcus
was higher; and genera Exophiala and Aureobasidium, the two black yeast-like fungi classified as
opportunistic pathogens were also detected. The presence of black yeast-like fungi was lower than in
previous cultivation based studies (16–19). This reduced incidence rate of these colonizers in DWs
could be due to the consequence of less efficient DNA extraction from black yeast cells due to the
presence of melanin in cell walls, large polysaccharide production or meristematic growth forms (52–
54). In addition, lower or higher incidence of microbial composition can also result from the chosen
methodology as sequencing analyses will also detect non-viable cells (55), resulting in an increased
microbial richness.
The density of microbial settlement on 1 cm2
rubber seals in DWs is only known for fungal
community (17) and not for bacterial community. The main colonizer black yeast E. dermatitidis was
detected at up to 106 CFU/cm
2; E. phaeomuriformis, R. mucilaginosa and C. parapsilosis were
detected in the range between 102 and 10
5 CFU/cm
2 (17). Ascomycetous fungi, namely Candida
dominated the recent DWs. These early fungal colonisers were followed by other co-occurring
Ascomycota and Basidiomycota members. It cannot be excluded that early colonisers may benefit
from the so-called “priority effect” (56), giving them the advantage to occupy the surface first, and
subsequently filter/chose the new comers, resulting in differential community assemblies. The
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hardness of tap water also significantly affected the fungal community in DWs where, more diverse
fungal species were present in DWs with hard and moderately hard water. Hard and moderately hard
water have increasing contents of Ca2+
and Mg2+
ions. Though water disinfection procedures have also
shown to influence fungal diversity (58, 59), it is noteworthy that the existence and morphology of
certain fungi depends on the presence of some ions, in particularly cations, such as Ca2+
(57). Other
ions, such as Cl−, generated by water chlorination, generally do not affect fungi (60, 61). Our results
indicate that establishment and diversity of fungi within DW biofilms will be greater with hard water,
while in soft water fungal biomass tends to be dominated by the fungal phyla Ascomycota. Further,
fungal composition could also be influenced by their adaption to different disinfectants used during
wash cycles in DWs.
In most natural environments, individual organisms do not live in isolation, but rather form a complex
community of different species that shape the structure, community itself and the evolution of the
individual species (62). In mixed bacterial-fungal biofilms, the early contact and adhesion are likely to
be important in the process of biofilm formation where mixed complexes of the two, bacteria or fungi
might provide biotic support for the establishment of biofilms (63–65). Microbes that live in the same
ecological habitat may co-occur or exclude each other. Studies have shown that coexistence can
facilitate interspecies interactions in biofilms (41, 66). This aspect was investigated as the fungal and
bacterial communities from 18 samples in this study that shared the same habitat and that they
therefore could provide insights into their possible interactions. Positive pairwise correlations indicate
mutual co-occurrence which also may point to symbiosis, mutualism or commensalism, whereas
negative pairwise correlations indicate mutual exclusion which may reflect competition, mutual
exclusion or parasitism (67). Our results indicate that bacterial groups co-occur with each other and so
do the fungal groups with other fungal members. Interestingly, cross-kingdom pairwise correlations
between fungi and bacteria were dominated by negative correlations which may reflect that they
occupy different locations on the biofilms.
However, in-vitro studies have shown close interactions between yeast cells forming the biofilm core,
and bacteria in the biofilm periphery create a protective coating for yeasts cells and pseudo-hyphae
(68, 69). Stressful conditions in DWs and the presence of bacteria could stimulate growth of fungi like
Candida spp. as pseudo-hyphae, thus enabling formation of the multispecies biofilm core on which
further bacteria could associate. This could be attributed to positive cross-kingdom correlation seen
between Saccharomycetes (mostly represented by the genus Candida) and Proteobacteria (α-
Proteobacteria). This suggests that the early members of the DW biofilm community and their
associations have possibly developed overtime in these environments. These could possibly support
the idea of strong priority effects, where first colonizers will strongly determine the chrono-succession
of events leading to establishment of successful biofilm structures in DWs.
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Conclusion
In this study, we investigated diverse bacterial and fungal communities in biofilms formed on different
DW rubber seals. Furthermore, abiotic conditions in DWs were shown to influence the microbial
community composition and several putative microbial pathogens that are important to food safety and
human health were presented. This study confirms that household appliances like dishwashers,
colonized by poly-extremotolerant bacteria and fungi, could present potential sources of domestically-
sourced infections. To understand and possibly prevent these phenomena, more studies should
investigate fungal interactions on bacterial physiology and vice-versa in biofilms formed on household
appliances.
Declarations
Acknowledgements
Our acknowledgements go to all the people who kindly provided samples from their dishwashers. We
also thank Karin Vestberg for her assistance with NGS and prof. Børge Diderichsen for careful and
critical reading of the manuscript.
Ethics approval and consent to participate
In this study, field sampling was performed, and to our knowledge, no endangered or protected species
were involved. All of the samples studied here were obtained from the discussed sampling areas, for
which permission was obtained from the owners.
Consent for publication
All authors read and approved the final manuscript.
Availability of data and materials
The sequence data sets generated during and analysed during the current study are available at the
NCBI Sequence Read Archive (SRA) under the Bioproject IDs: PRJNA315977 for bacterial reads and
PRJNA317625 for fungal reads.
Competing interest
The authors declare that they have no conflicts of interest.
Funding
This research was funded by the Ministry of Higher Education, Science and Technology of the
Republic of Slovenia, as a Young Researcher grant to JZ (grant no. 382228-1/2013). We also thank
the Slovenian Research Agency (Infrastructural Centre Mycosmo, MRIC UL) and the Danish Council
for Independent Research grant: 1323 00235 for providing financial support. The funding bodies had
no influence on the design of the study and collection, analysis, and interpretation of data and in
writing the manuscript.
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Authors’ contributions
SJS, NGC and MB designed the experimental setup. PKR and JZ performed the experiments. ADB, SJ
and PKR analysed the data. PKR, JZ, ADB, SJ, KH, NGC, MB, and SJS compiled the manuscript.
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Supplementary information
Supplementary Figure S1: Microbial composition in individual samples at the phylum level. (A)
Fungal and (B) bacterial abundance (%) across DW samples. The low abundant phyla Acidobacteria,
Chloroflexi, Cyanobacteria, Spirochaetes, Synergistetes, Verrucomicrobia and other
unclassified/unassigned reads are combined in the miniscule ‘others’. Gap in the dataset refer to
samples where sequence data could not be generated.
A
0% 20% 40% 60% 80% 100%
S1
S2
S3
S4
S5
S6
S7
S8
S9
S10
S11
S12
S13
S14
S15
S16
S17
S18
S19
S20
S21
S22
S23
S24
Abundance
Actinobacteria
Bacteroidetes
Firmicutes
Proteobacteria
TM7
Deinococcus-Thermus
Others
B
0% 20% 40% 60% 80% 100%
S1
S2
S3
S4
S5
S6
S7
S8
S9
S10
S11
S12
S13
S14
S15
S16
S17
S18
S19
S20
S21
S22
S23
S24
Abundance
Ascomycota
Basidiomycota
Zygomycota
Others
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Supplementary Figure S2: Significant co-occurrence and co-exclusion interactions among bacterial
(‘blue circles’) and fungal (‘grey circles’) OTUs in the DWs microbiome. Predicted pairwise
interaction network between the bacterial and fungal OTUs generated from the OTU matrix. The
displayed pairwise co-occurrences appeared in at least 50% of all samples and were dominated by
positive correlations indicated by ‘green’ connectors. ‘Red’ connectors indicate negative correlations.
[MANUSCRIPT 2]
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Supplementary Table S1: Summary table of 16S rRNA gene and ITS gene based amplicon profiles
obtained. ‘No reads’ indicates samples where sequences reads could not be generated. SRR represents
the SRA accession numbers obtained after deposition of raw reads to NCBI database.
Bacterial Richness Fungal Richness
Samples OTUs Counts SRR Samples OTUs Counts SRR
S1 97 12056 3335213 S1 26 4536 3354512
S2 104 14880 3279031 S2 30 47413 3354513
S3 35 13294 3335242 S3 64 52407 3354514
S4 105 11086 3343755 S4 32 53664 3354515
S5 56 4812
3343756
S5 No
reads
No
reads
No
reads
S6 104 8872 3343757 S6 43 9514 3354535
S7 112 5790 3343758 S7 26 28574 3354573
S8 No
reads
No
reads
No
reads
S8 22 37174
3354574
S9 No
reads
No
reads
No
reads
S9 60 24041
3354575
S10 115 17912
3343759
S10 No
reads
No
reads
No
reads
S11 55 1511 3343760 S11 33 14060 3354576
S12 60 961 3343761 S12 26 19847 3354577
S13 104 14995 3343763 S13 28 9602 3354578
S14 96 6661
3343796
S14 No
reads
No
reads
No
reads
S15 107 9683 3343797 S15 15 10340 3354579
S16 50 127 3343798 S16 20 3520 3354580
S17 107 3580 3343799 S17 18 3601 3354581
S18 No
reads
No
reads
No
reads
S18 30 2162
3354582
S19 95 24429 3343800 S19 104 4372 3354583
S20 29 100 3343802 S20 25 660 3354584
S21 71 8529 3343803 S21 45 891 3354585
S22 110 26115 3343805 S22 27 472 3354587
S23 92 15960 3343810 S23 30 436 3354588
S24 150 13036 3335236 S24 40 49511 3354600
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Supplementary Table S2: Dishwasher alpha diversity summary post rarefaction of sequence counts.
‘NA’ indicates samples in which sequences reads were not generated.
Bacterial Diversity Fungal Diversity
Samples Richness
Taxa_S
Shannon_H Chao-
1
Samples Richness
Taxa_S
Shannon_H Chao-
1
S1 69 2.7 78 S1 10 0.7633 11
S2 75 2.9 82 S2 5 0.32 5
S3 18 1.2 19 S3 16 1.658 16
S4 75 2.2 103 S4 10 0.9061 12
S5 48 1.7 54 S5 NA NA NA
S6 72 2.0 83 S6 13 1.441 16
S7 87 2.7 108 S7 3 0.03494 4
S8 NA NA NA S8 19 1.601 24
S9 NA NA NA S9 NA NA NA
S10 78 2.4 92 S10 15 1.655 17
S11 - - - S11 7 0.3897 9
S12 - - - S12 8 0.8721 8
S13 68 2.5 79 S13 NA NA NA
S14 73 2.8 103 S14 4 0.697 4
S15 84 3.0 105 S15 10 1.039 25
S16 - - - S16 7 0.261 10
S17 - - - S17 12 1.632 14
S18 NA NA NA S18 42 2.714 46
S19 51 2.2 62 S19 14 0.8477 16
S20 - - - S20 26 1.481 32
S21 50 2.0 55 S21 21 2.483 23
S22 59 2.2 74 S22 26 1.276 30
S23 57 1.3 83 S23 22 1.669 31
S24 110 3.6 133 S24 10 0.7633 11
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Supplementary Table S3: Summary of p-values from pairwise comparisons on bacterial alpha
diversity indices using Wilcoxon-Mann-Whitney.
Supplementary Table S4: Summary of p-values from pairwise comparisons on fungal alpha diversity
indices using Wilcoxon-Mann-Whitney (* p ≤ 0.05)
Chao-1 Year 0-4 5-7 Freq 0-6 WH MS SH MH
0-4 - 0.72 7-14 0.20 MS - 0.19 0.27
5-7 0.72 - SH 0.19 - 0.71
8 0.91 0.82 MH 0.27 0.71 -
H 0.19 0.01* 0.02*
Shannon_H Year 0-4 5-7 Freq 0-6 WH MS SH MH
0-4 - 0.04* 7-14 0.31 MS - 0.28 0.64
5-7 0.04* - SH 0.28 - 0.38
8 0.84 MH 0.64 0.38 -
H 0.02* 0.0009* 0.003*
Richness Year 0-4 5-7 Freq 0-6 WH MS SH MH
0-4 - 0.56 7-14 0.4 MS - 0.33 0.25
5-7 0.56 - SH 0.33 - 0.85
8 0.98 0.60 MH 0.25 0.85 -
H 0.05* 0.01* 0.01*
Chao-1 Year 0-4 5-7 Freq 1-3 7 WH MS SH MH
0-4 - 0.43 1-3 - 0.56 MS - 0.55 0.31
5-7 0.43 - 7 0.56 - SH 0.55 - 0.62
8 0.47 0.21 14 0.99 0.70 MH 0.31 0.62 -
H 0.27 0.49 0.72
Shannon_H Year 0-4 5-7 Freq 1-3 7 WH MS SH MH
0-4 - 0.52 1-3 - 0.16 MS - 0.35 0.07
5-7 0.52 - 7 0.16 - SH 0.35 - 0.28
8 0.37 0.22 14 0.63 0.16 MH 0.07 0.28 -
H 0.28 0.71 0.66
Richness Year 0-4 5-7 Freq 1-3 7 WH MS SH MH
0-4 - 0.41 1-3 - 0.28 MS - 0.42 0.24
5-7 0.41 - 7 0.28 - SH 0.42 - 0.67
8 0.30 0.13 14 0.99 0.46 MH 0.24 0.67 -
H 0.20 0.46 0.65
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105
Supplementary Table S5: Summary from redundancy based ANOVA analysis done using each of
the factors (999 permutations). The composition of the bacterial community was significantly affected
by the factors ‘years of use’ and ‘frequency’ of DWs. The composition of fungal community was
significantly affected by years of use, frequency of use and tap water hardness. df: degrees of freedom;
F: ratio of the between groups variance and within groups variance; p: p-values. Star indicates the
level of significance of each factor: ‘*’ p < 0.05.
Source Factors df Variance (%) F p
Bacteria
Diversit
y
Years (0-4y, 5-7y and 8y) 2 23.44 1.34 0.047*
Temperature (≥65°C, ≤60°C) 1 9.63 1.11 0.299
Frequency of use ( 1-3, 7 and 14) 2 27.21 1.55 0.022*
Water Hardness ( H,MH,SH and MS) 3 24.00 0.91 0.660
Fungi
Diversit
y
Years (0-4y, 5-7y and 8y) 2 9.47 1.32 0.043 *
Temperature (≥65°C, ≤60°C) 1 4.04 1.12 0.280
Frequency of use ( 0-7 and 14) 1 5.78 1.61 0.034 *
Water Hardness ( H,MH,SH and MS) 3 18.87 1.76 0.017 *
Supplementary Table S6: PERMANOVA based on Bray-Curtis dissimilarity index and dishwasher
conditions using 999 permutations. The analyses were executed using the vegan package for R
software. df: degrees of freedom; R2: coefficient of determination; p: p-values. Star indicates the level
of significance of each factor: ‘*’ p < 0.05.
Source Factors df R2 p
Bacteria
Diversity
Years of use 2 0.11 0.186
Temperature 1 0.05 0.386
Water hardness 3 0.13 0.519
Frequency of use 2 0.14 0.045*
Residuals 12 0.56
Fungi
Diversity
Years of use 2 0.10 0.200
Temperature 1 0.06 0.135
Water Hardness 3 0.17 0.018*
Frequency of use 1 0.09 0.127
Residuals 13 0.58
106
Zupančič J, Raghupathi PK, Houf K, Burmølle M, Sørensen SJ, Gunde-Cimerman N. Synergistic
Interactions in Microbial Biofilms Facilitate the Establishment of Opportunistic Pathogenic Fungi in
Household Dishwashers. Front Microbiol. 2018 Jan 30; 9:21. Doi: 10.3389/fmicb.2018.00021.
MANUSCRIPT 3
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Synergistic interactions in microbial biofilms facilitate the establishment of opportunistic
pathogenic fungi in household dishwashers
Jerneja Zupančič1Ϯ
, Prem Krishnan Raghupathi2, 3Ϯ
, Kurt Houf3, Mette Burmølle
2, Søren J. Sørensen
2,
Nina Gunde–Cimerman1
1 Department of Biology, Biotechnical Faculty, University of Ljubljana, Ljubljana, Slovenia;
2
Molecular Microbial Ecology Group, Section of Microbiology, Department of Biology, University of
Copenhagen, Copenhagen, Denmark; 3 Faculty of Veterinary Medicine Department of Veterinary
Public Health and Food Safety, Laboratory of Hygiene and Technology, Merelbeke, Belgium
Biofilms formed on rubber seals in dishwashers harbour diverse microbiota. In this
study, we focussed on the microbial composition of bacteria and fungi, isolated from a defined
area of one square centimetre of rubber from four domestic dishwashers and assessed their
abilities to in-vitro multispecies biofilm formation. A total of 80 isolates (64 bacterial and 16
fungal) were analysed. Multiple combinations of bacterial isolates from each dishwasher were
screened for synergistic interactions. 32 out of 140 tested (23%) four-species bacterial
combinations displayed consistent synergism leading to an overall increase in biomass, in all
experimental trails. Bacterial isolates from two of the four dishwashers generated a high number
of synergistically interacting four-species consortia. Network based correlation analyses also
showed higher co-occurrence patterns observed between bacterial members in the same two
dishwasher samples, indicating cooperative effects. Furthermore, two synergistic 4-species
bacterial consortia were tested for their abilities to incorporate an opportunistic fungal
pathogen, Exophiala dermatitidis and their establishment as biofilms on sterile ethylene
propylene diene monomer M-class (EPDM) rubber and polypropylene (PP) surfaces. When the
bacterial consortia included E. dermatitidis, the overall cell numbers of both bacteria and fungi
increased and a substantial increase in biofilm biomass was observed. These results indicate a
novel phenomenon of cross kingdom synergy in biofilm formation and these observations could
have potential implications for human health.
Ϯ Shared First Author
[MANUSCRIPT 3]
108
Introduction
Biofilms are defined as highly structured communities of microorganisms that are attached to each
other, commonly surface associated and enclosed within a self-produced matrix of extracellular
polymeric substance (EPS) (Costerton et al., 1995). The advantages obtained by organisms from
producing biofilms include protection from harsh environments, enhanced tolerance to physical and
chemical stress, metabolic cooperation and community-coordinated adjustment of gene expression.
Microorganisms in biofilms adapt their physiology and stress responses and display collective and
coordinated behaviour (Donlan, 2002; Chmielewski and Frank, 2004; Van Houdt and Michiels, 2010).
Multispecies biofilms are common and often dominant in natural environments (Donlan, 2002; Hall-
Stoodley et al., 2004). Resident microorganisms interact with each other in both synergistic and
antagonistic manner affecting the biofilm biomass, functionality and tolerance compared to mono-
species biofilms (Sharma et al., 2005; Filoche et al., 2004, Ren et al., 2015; Burmølle et al., 2006;
Pathak et al., 2012; Wen et al., 2010; Schwering et al., 2013, Madsen et al., 2016; Lee et al., 2014;
Moons et al., 2009).
Biofilms are a source of food contamination and food safety related problems (Srey et al., 2013;
Carpentier and Chassaing, 2004, Røder et al., 2015). In food production facilities, pathogenic bacteria
may benefit from biofilm formation (Klayman et al., 2009) as biofilms can withstand higher
temperatures, standard cleaning procedures (Marouani-Gadri et al., 2010) and commonly used
disinfectants (Corcoran et al., 2014) thereby, leading to biofilm related outbreaks (Donlan, 2002; de
Souza et al., 2015). Most studies focus on the biology and persistence of monocultures of a particular
bacterial pathogen in biofilm (Lister and Horswill, 2014; Tolker-Nielsen, 2014), however there is a
growing need to understand the impact of interspecies interactions on the formation and architecture of
biofilms (Sheppard et al., 2016; Elias and Banin, 2012). Increasing evidence points to the role of fungi
in biofilms involved in human diseases (Ramage et al., 2009, Hoarau et al., 2016, Kalan et al., 2016).
In mixed bacterial and fungal biofilms, it was reported that bacterial cells gained protection within the
matrix and increased its tolerance to antimicrobials and stress (Kong et al., 2016; De Brucker et al.,
2015).
Recently, it was discovered that the extreme depauperate ecosystem of household appliances, such as
dishwashers, washing machines and coffee machines, harbour selected poly-extremotolerant bacteria
and fungi (Zalar et al., 2011; et al.Callewaert et al., 2015; Novak Babič et al., 2015; Vilanova et al.,
2015, Zupančič et al., 2016; Raghupathi et al., 2017). These microbes resist both high and low pH,
temporary increase in temperatures up to 74 °C, desiccation, high organic loads, high concentrations
of NaCl and mechanical stress from water ejectors (Zalar et al., 2011; Zupančič et al., 2016). They are
represented by diverse human opportunistic fungi (Zalar et al., 2011; Döğen et al., 2013; Gümral et
al., 2015; Zupančič et al., 2016) and bacteria (Raghupathi et al., 2017).
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We have focussed on mixed biofilms in dishwashers since there is a worldwide increase in demand for
household appliances (Freedonia, 2016) and opportunistic pathogens detected in these machines could
be an emerging threat to human health (Binder et al., 1999; Morens et al., 2004). Despite the ubiquity
of microbial communities and the presence of dishwashers in many private households, interspecies
interactions among different bacteria and fungi have not been investigated in these systems. The focus
of present research was to identify the species composition of bacteria and fungi from the rubber seals
of four different dishwashers. The viable bacterial and fungal isolates were identified using a
combination of classical and molecular methods. Multiple combinations of different bacterial isolates
from each these dishwashers were co-cultured in-vitro and their ability to form stable, four-species
biofilms was assessed. The synergistic bacterial consortia were tested for their ability to incorporate
Exophiala dermatitidis (the most common opportunistic fungal pathogen found in dishwashers) (Zalar
et al., 2011; Döğen et al., 2013; Gümral et al., 2015; Zupančič et al., 2016) and their establishment as
mixed bacterial-fungal biofilm on different surfaces commonly used in dishwashers were investigated.
Experimental Procedures
Cultivation and identification of the microbial community from 1 cm2 of dishwasher biofilms
Microbial biofilms formed on 1 cm2 area of rubber seal from 4 different dishwashers were sampled in
this study (Table 1).
Table 1: Dishwashers sampled for microbial composition in this study. The dishwashers varied in age,
frequency of use and influent water hardness characteristics. DW1=dishwasher 1; DW2= dishwasher
2; DW3=dishwasher 3; DW4 = dishwasher 4; SH = slightly hard (1-1.5 mmol/L CaCO3); MH =
moderately hard (1.5 – 2.0 mmol/L CaCO3); MH= moderately soft (0.5-1.0 mmol/L CaCO3). ‘SRR’
represents the sequence read archive assigned after deposition of 16s rRNA gene marker-based
amplicon reads to NCBI database.
Dishwasher Country; City;
GPRS coordinates
Age
(years in
use)
Frequency of
use/week
Influent
water
NCBI
SRR
DW1
SI; Žalec;
46°15′3.59″N
15°9′50.18″E
3 7 SH 3279031
DW2 SI; Ljubljana;
46°03′′N 14°30′′E 5 3 MH 3335242
DW3 SI; Brezovica;
45°58′11.68″N 7 3 MH 3343759
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14°26′9.95″E
DW4
SI; Novo Mesto;
45°47′54.88″N
15°10′26.08″E
8 7 MS 3335236
The dishwashers varied in age, i.e. years in operation; frequency of use, i.e. the no. of. Times the
dishwasher was used per week; and incoming tap water hardness. The water supply connected to these
DWs was characterized based on ion analysis method (Babič et al., 2013). Final concentrations were
determined following the method from ISO Standard SIST EN ISO 11885:2009. Biofilm samples
were collected with sterile swabs (Invasive sterile EUROTUBO® collection swab). Sampling of
microbiota was performed by rubbing a cotton swab moistened with physiological saline over 1 cm2
rubber seal surfaces, immediately after the termination of the washing cycle in these dishwashers.
Swab samples were stored in sterile collection tubes at 4 °C and were processed within a day.
Cultivable microbes living in close contact from each of these dishwashers were cultivated by plating
methods to obtain individual bacterial and fungal colonies. For each dishwasher sample, 3 ml of sterile
physiological saline was added into the collection tube containing swabs and vortexed intensely for 1
min at maximum speed. Subsequently, for bacterial screening, aliquots of 100 µl of the sample were
diluted 10-fold and plated on different bacteriological agar media i.e. nutrient agar (NA), Brain-Heart
Infusion agar (BHI), Reasoner´s 2A agar (R2A), and Minimal Media agar (M9) (Vogel & Bonner,
1956). All plates were supplemented with cycloheximide (CYC, 50 µg ml-1
, Sigma) to ensure only
bacterial growth. Plates were incubated aerobically at 37 °C for 2 days (NA and BHI) and up to 7 days
for M9. In case of R2A, plates were incubated for 7 days at 35 °C. Isolation of fungi was performed by
inoculating same aliquots of 100 µl of the above diluted suspension on Malt Extract Agar (MEA)
(Oxoid, Hampshire, UK) supplemented with 0.05 g/l chloramphenicol, and incubated at 30 °C and 37
°C for up to 7 days.
Microbial colonies of various morphotypes (both bacterial and fungal) were restreaked several times
on chosen media plates (Luria Bertani (LB) for bacteria and MEA for fungi until pure cultures were
obtained. The pure cultures were deposited and can be obtained from the Ex Culture Collection, part
of the Infrastructural Centre Mycosmo (MRICUL) at the Department of Biology, Biotechnical
Faculty, University of Ljubljana, Slovenia.
Identification of isolates using Sanger sequencing
DNA extraction and molecular identification of fungal isolates from dishwashers was performed as
previously described (Zupančič et al., 2016). Briefly, pure fungal cultures were transferred to fresh
MEA medium and after 3 -7 days of incubation and DNA extractions were performed with methods
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specific to the type of fungal isolates. For yeasts, DNA extraction was using PrepMan Ultra Sample
Preparation Reagent (Applied Biosystems) according to the manufacturer’s instructions. DNA
extractions of filamentous fungi and Exophiala strains were done according to Gerrits van den Ende
and de Hoog (1999), after mechanical lysis of the mycelium. Fusarium strains were identified using
nuclear translation elongation factor 1-alpha (tef) sequences, amplified with the EF1 and EF2 primers
(O'Donnell et al., 1998).
Bacterial identification was performed using the extracted genomic DNA from overnight grown pure
cultures (LB plates incubated at 37 °C) using PrepMan Ultra Sample Preparation Reagent (Applied
Biosystems) according to the manufacturer’s instructions. PCR amplifications based on 16S rRNA
gene with oligonucleotide primers 27F and 1492R targeting bacterial 16S ribosomal gene (Lane, 1991)
were applied for bacterial identification. The amplified fragments were Sanger sequenced (Microsynth
AG) and the 16S rRNA gene sequences were trimmed to approx. 800bp amplicons and identification
was done using Ribosomal Database Project-II (RDP) (http://rdp.cme.msu.edu) and National Center
for Biotechnology Information (NCBI) BLAST tool searching GenBank. RDP Seqmatch was used
against the 16S rRNA database with sequences from isolated bacteria in order to determine the closest
known relatives. The amplicon sequences were also compared against GenBank non-redundant
nucleotide database using NCBI BlastN (Megablast). The isolates were assigned at species level with
the Seqmatch score (S-ab) ≥ 0.99 (99% similarity) or at genus level with S-ab score of ≥ 0.95 (95%
similarity). Sequences were uploaded to the NCBI database and the accession numbers are provided
(Table 2).
Growth Media and Conditions
To determine the optimal growth conditions and evaluate the biofilm-forming capabilities of
microorganisms obtained in this study, we selected 7 bacterial isolates from each of the four
dishwashers providing a total of 28 bacterial isolates (Table 2). Selections of isolate were made
between different phylogenetically diverse bacterial species in each dishwasher. These isolates were
subcultured from frozen glycerol stocks onto LB (Luria-Bertani) agar plates and incubated for 24 h at
37 °C. A single colony of each bacterial isolate was inoculated into 5 ml LB media tubes, incubated
overnight at 37°C while shaken at 200 rpm.
In-vitro bacterial multispecies biofilm cultivation
The seven selected isolates from each dishwasher (Table 2) were screened for biofilm formation as
single species and in four-species combinations as described previously (Røder et al., 2015; Ren et al.,
2015) with few modifications. Serial 10-fold dilutions of bacterial cultures were performed from
overnight grown cultures (in LB media) where 1 ml of the dilutions were inoculated with 29 ml fresh
LB media, incubated overnight at 37°C and shaking at 200 rpm. Cell cultures in exponential phase
(OD600 between 0.3 – 0.7) were then selected, centrifuged at 8000 rpm (10 min, 21°C), washed with 1x
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phosphate buffer saline (PBS) and re-suspended in 10% w/v LB media (reduced). The optical density
OD600 of each bacterial culture was then adjusted to 0.15 in the reduced LB media. Biofilm cultivation
assay was performed using 96-well microtiter plates (NUNC, Roskilde, Denmark) and peg lids
(NUNC-TSP lid system, Roskilde, Denmark) placed on top of the plates, also referred to as the
Calgary method (Ceri et al., 1999). A total of 150 µl as mono-species or four mixed species (37.5 µl of
each species) cultures were added to each well. Each plate contained the representative mono-species
cultures. 150 µl 10% LB served as blank. Plates were incubated at 25°C for 24 hours.
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Table 2: List of selected bacterial isolates used in biofilm cultivation experiments. DW1 - dishwasher 1; DW2 - dishwasher 2; DW3 - dishwasher 3; DW4 -
dishwasher 4; * Phyla; P – Proteobacteria, F – Firmicutes, A – Actinobacteria. Strain ID represents the isolate identification after deposition (as ‘EXF’ for
fungal and ‘EXB-L’ for bacterial isolates) at the Microbial Culture Collection Ex (MRICUL EX).
Isolate
source
ID# Closest relative * Strain ID
EXF- / EXB L
Accession
no. of. the
closest
relative
NCBI
Accession
number
DW1
1 Pseudomonas aeruginosa P EXB L-1125 KR911837 MG59730
1
2 Ochrobactrum
pseudintermedium
P EXB L-1130 KF026284 MG59730
2
3 Klebsiella oxytoca P EXB L-1137 CP011636 MG59730
3
4 Stenotrophomonas maltophilia P EXB L-1167 KP185140 MG59730
4
5 Enterobacter hormaechei P EXB L-1135 KP303395 MG59730
5
6 Pseudomonas putida P EXB L-1149 KJ735915 MG59730
6
7 Bacillus cereus F EXB L-1175 KC969074 MG59730
7
DW2
8 Acinetobacter lwoffii P EXB L-1215 LN774665 MG59730
8
9 Bacillus cereus F EXB L-1223 KP988025 MG59730
9
10 Exiguobacterium aestuarii F EXB L-1196 FJ462716 MG59731
0
11 Exiguobacterium panipatensis F EXB L-1201 EF519705 MG59731
1
12 Kocuria rhizophila A EXB L-1199 AY030315 MG59731
2
13 Micrococcus luteus A EXB L-1190 KF993675 MG59731
3
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14 Pseudescherichia vulneris P EXB L-1211 JQ958880 MG59731
4
DW3
15 Bacillus circulans F EXB L-1279 KM349203 MG59731
5
16 Micrococcus luteus A EXB L-1261 KJ733861 MG59731
6
17 Microbacterium
hydrocarbonoxydans
A EXB L-1250 JQ954857 MG59731
7
18 Exiguobacterium aestuarii F EXB L-1244 FJ462716 MG59731
8
19 Exiguobacterium arabatum F EXB L-1278 JF775422 MG59731
9
20 Exiguobacterium panipatensis F EXB L-1260 EF519705 MG59732
0
21 Exiguobacterium profundum F EXB L-1270 KM873375 MG59732
1
DW4
22 Acinetobacter junii P EXB-L-1308 EU862296 MG59732
2
23 Haematomicrobium sanguinis A EXB-L-1326 EU086805 MG59732
3
24 Bacillus cereus F EXB-L-1176 GU568201 MG59732
4
25 Brevibacterium casei F EXB-L-1336 HM012705 MG59732
5
26 Exiguobacterium panipatensis F EXB-L-1316 EF519705 MG59732
6
27 Exiguobacterium aestuarii F EXB-L-1327 FJ462716 MG59732
7
28 Staphylococcus saprophyticus F EXB-L-1314 AB697718 MG59732
8
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Network analysis data
While competing for same resources, bacteria present in the same environment potentially co-occur or
exclude each other (Roggenbuck et al., 2014). This relationship was characterized by generating the
Spearman co-occurrence network (Barberán et al., 2012). The four selected dishwasher in this study,
sequenced using Illumina MiSeq platform and taxonomic classifications of the 16S rRNA gene
sequences based on RDP classifier, were described previously (Raghupathi et al., 2017). Sequence
raw reads (SRR) (Table 1) from these dishwashers were made available to NCBI Sequence Read
Archive (SRA) under the Bioproject ID: PRJNA315977. The network and predicted interactions were
generated on the basis of relative counts of different bacterial genera that had more than 50 sequence
observations and represented in 50% of the samples (n > 2, N=4). We present correlation data for log
transformed counts using CoNet 1.0b6 plugin in Cytoscape 3.2.1. The correlations were made on the
basis of in-built nonparametric Spearman correlation coefficient with a minimal cut-off threshold of r
≥ |0.85| (p ≪0.01, Bonferroni corrected).
In-vitro cultivation of Bacterial-Fungal biofilms
The bacterial isolates from DW4 were prepared as mentioned above. The fungal strain E. dermatitidis
genotype A (EXF-9777), also isolated from DW4 (Table 2), was subcultured from frozen glycerol
stocks onto MEA, supplemented with 0.05 g/l chloramphenicol and incubated 3-5 days at 37 °C. A
single colony of the black yeast was then inoculated into 5 ml 10% LB media tubes and incubated at
37°C while shaken at 200 rpm until an OD600 of approx. 0.7 was reached. Then, with the aim to work
with a uniform culture media which will provide a common niche for both bacteria and fungi, LB
media was replaced with 10% LB and OD600 adjusted to 0.15. A total of 150 µl as mono-species
(bacteria/fungi) cultures or 30 µl for each species in five mixed species (four bacteria and E.
dermatitidis) combinations were added to each well. Also, each plate contained the representative 75µl
of mono-species bacterial cultures together with 75µl fungal cultures. Plates were incubated at 25°C
for 24, 48 and 96 hours. 150 µl 10% LB served as blank.
Biofilm quantification and screening for synergistic interactions
Mixed species and monospecies biofilm cultivation in a 96-well Calgary Biofilm Device (CBD) and
its quantification using 1% w/v crystal violet were performed as described previously (Røder et al.,
2015; Ren et al., 2015). We classified synergy, as and when the measured absorbance from the CBD
assay of the multispecies biofilm (MSB) being greater than that of the best single strain (BSS) biofilm
producer present in the relevant combination when taking standard errors into account, i.e. (Abs590
MSB - Std. error) > (Abs590 BSS + Std.error) = Synergy, while (Abs590 MSB + Std. error) < (Abs590
BSS -Std. error) = No synergy (Ren et al.. 2015). In case of bacterial-fungal biofilms, synergy was
when the absorbance of multispecies bacterial-fungal biofilm was greater than that of the best single
strain biofilm producer present together with the fungi (BSS) in the relevant combination when taking
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standard errors into account. Fold change (Fd) is represented as ratio of the biofilm biomass of
multispecies consortia with/without fungi to its best biofilm producer with /without fungi within the
respective consortia i.e. Fold change = Abs590 MSB - Std. Err / Abs590 (BSS + Std. Err). Hence,
consortia with an Fd > 1 are designated as synergistic. The above cultivation and quantification of
biofilm was performed with three technical replicates and the assay was performed at three different
times.
In-vitro establishment of multispecies biofilm on dishwasher rubber and plastic material and its
quantification
Two four-species bacterial consortia from DW4 that showed an overall increase in biofilm formation
in all trials, were tested for the incorporation of E. dermatitidis using a 24 well plate; as this fungus
was found to be present on DW4 rubber seal. Enumeration of fungal and bacterial cells from the
biofilm formed on wells was done using fluorescent associated cell sorting system BD FACS Calibur
(BD Biosciences). The biofilm on the bottom of the plates were washed gently and the attached cells
were scrapped-off, homogenized in 500 µl 1X PBS and transferred into micro-centrifuge tubes. The
fungal cells were selectively stained using Calcofluor White Stain (Sigma-Aldrich) to differentiate
from bacterial cells.
Further, the biofilm formation on three different types of elastomer; EPDM (ethylene propylene diene
monomer (M-class)) referred to as 17, 18, 19 and three different types of polypropylene (PP) (C3H6)n
referred to as 1, 2, 3; used in dishwasher industry were tested. The elastomer and plastic material were
cut into slices of 1 cm2 size (with active surface 2 x 1 cm
2) and sterilized by autoclaving at 121 °C for
15 min. Bacterial and fungal cultures were prepared as described above. 24-well cell culture plates
(TPP® cat. no. 92024, Sigma-Aldrich, USA) were used to cultivate the biofilms on artificial materials
of EPDM and PP. A total of 1250 µl for monospecies bacterial or fungal cultures or four mixed
species (312, 5 µl of each bacterial culture), or five mixed species (250 µl of each bacteria and fungi
cultures) combinations were added to each well. The same volume of 10% LB medium was added as
blank. After inoculation, sterile elastomer or plastic parts were aseptically added into the plates. The
plates were incubated at 25 °C for 24h, 48h, and 120h. The biofilm assays were performed three times
on different days with three technical replicates each time.
The crystal violet method was applied to quantify biofilms formed on EPDM / PP (Røder et al., 2015;
Ren et al., 2015) as follows. Briefly, after incubation, in order to wash off loosely attached cells and
planktonic fractions, the EPDM / PP substrates were transferred using sterile forceps successively to
three 24-well microtiter plates containing 1200 µl of 1X PBS buffer per well, followed by staining of
the biofilms formed on the EPDM / PP with 1250 µl of an aqueous 1 % (w/v) CV solution. After 20
min, the EPDM / PP substrate was rinsed three times with 1X PBS and de-stained in 1250 µl 96 %
ethanol in each well of a new plate. After 20 min, the absorbance was measured as described above.
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Results
Variation in total cultivation community structure across four different dishwashers
Among the dishwashers that were screened for viable microbial population within 1 cm2 isolation area
from four dishwashers (DW), a total of 80 isolates (64 bacterial and 16 fungal) were obtained
(Supplementary Information, Table S1). Isolates from DW1 contained seven different fungal species
and 20 different bacterial species. The fungal isolates belonged to four different classes viz.
Saccharomycetes, Chaetothyriomycetes, Sordariomycetes and Urediniomycetes. Majority of the
isolated bacterial species belonged to Proteobacteria; and others belonged to four different bacterial
phyla. DW2 had 3 fungal species and 18 different bacterial species belonging to 3 different bacterial
phyla. 10 Gram-positive isolates belonging to two bacterial phyla, Firmicutes and Actinobacteria and
no fungal isolates were obtained from DW3. Majority of these bacterial isolates belonged to the genus
Exiguobacterium. Isolates from DW4 contained three different fungal species belonging to 3 fungal
classes and 16 different bacterial species. The 16 isolates belonged to 4 different bacterial phyla.
Bacterial isolates from DW3 and DW2 were represented by two or three families (DW3:
Microbacteriaceae and Bacillaceae; DW2:-Enterobacteriaceae Micrococcaceae and Moraxellaceae)
respectively. Bacterial isolates from DW1 and DW4 were represented by 5 families (DW1:
Pseudomonadaceae, Brucellaceae, Enterobacteriaceae, Xanthomonadaceae and Bacillaceae; DW4:
Moraxellaceae Bacillaceae Staphylococcaceae Brevibacteriaceae and Micrococcaceae). DW2 and
DW4 contained the black yeast E. dermatitidis, represented by two different genotypes, of which, the
clinically relevant genotype A was present in both DWs. Previous results showed the most abundant
microbial taxa in these four DW samples identified by 16S rRNA and ITS gene marker based
amplicon sequencing. Most abundant bacterial taxa belonged to genera like Exiguobacterium,
Gordonia, Nesterenkonia, Ochrobactrum, Chryseobacterium, Stenotrophomonas, Pseudomona and
Acinetobacter. Most abundant fungal taxa in these four DW samples were represented by genera
Candida, Cryptococcus, Rhodotorula and Exophiala (Raghupathi et al., 2017; Fig S1 Supplementary
information).
Bacteria classified as opportunistic pathogens like Pseudomonas aeruginosa, Ochrobactrum
pseudintermedium, Klebsiella oxytoca and Acinetobacter junii and opportunistic fungal pathogens like
E. dermatidis, Candida parapsilosis, Rhodotorula mucilaginosa and Fusarium oxysporum species
complex (FOSC) were isolated from these dishwashers. Bacterial and fungal isolates from DW1, 2 and
4 were represented by various opportunistic pathogens whereas; the isolates from DW3 were
represented by non-pathogenic “environmental” strains (Fig 1). This classification was made based on
known fungal and bacterial taxonomic literatures (de Hoog et al., 2014; Whitman WB, 11th ed.
Bergey's Manual of Systematics of Archaea and Bacteria (BMSAB, 2017)).
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Figure 1: Distribution of microbial population isolated from the rubber seals of 4 DWs. After isolation
and identification of both bacteria and fungi, isolates were classified as environmental or opportunistic
pathogenic strains based on the taxonomic literature (de Hoog et al., 2014; BMSAB, 2017) in each
dishwasher; DW1- dishwasher 1; DW2- dishwasher 2; DW3- dishwasher 3; DW4 – dishwasher 4.
Multi-species interactions enhance biofilm biomass
Screening for biofilm formation revealed that DW1 and DW4 had higher percentage of four-species
consortia with fd > 1, thus considered to be synergistic in biofilm formation, compared to DW2 and
DW3 (Fig 2). Overall 35 four-species combinations were tested per each DW, 140 combinations in
total per experiment. Results showed that DW1, DW2 and DW4 had 9, 2 and 21 stable four-species
combinations, respectively, (consistently synergistic (fold-change, fd >1) in all three trials. DW3 had
no four-species combinations interacting synergistically across all trials. The absorbance
measurements of single and 4-species combinations and their corresponding fold-change (fd)
calculated across the three biological trails are shown (Supplementary information, Table S2).
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Figure 2: Percentage of synergistic (fd value >1) 4-species bacterial combinations (total; N=35) from
DW1-4 based on CV quantification after 24h incubation at 25°C in 10% LB media. The number above
each column bar indicates the total number of 4 species consortia having fd value >1 in all replicates
and trials. The experiment was performed at three different times with three technical replicates each
time. The error bars denote the percentage mean ± standard error (S.E) from three biological trails.
The four-species consortia were analysed to identify the different species contributing as key biofilm
producers when present within the given consortia. Therefore, the isolates that contributed more
frequently to synergy in each 4-species combination were obtained. The analysis performed across
three trials gave a maximum count of 60 combinations per isolate (Fig 3). In DW1, 4-species
combinations containing Pseudomonas aeruginosa and Enterobacter hormaechei were more likely to
interact synergistically. In DW4, Acinetobacter junii was the most frequent isolate contributing to
synergistic interactions. In DW2 and DW3, the frequency of each isolate to engage in a synergistic 4-
species biofilm varied among different bacterial members. Escherichia vulneris and Exiguobacterium
aestuarii in different 4-species combinations were more likely to interact synergistically in DW2 and
DW3, respectively.
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Figure 3: Isolates from dishwashers that were assessed as key species contributing to biofilm synergy
when present in a 4-species combination. The number of times an isolate contributing to synergy
within a 4-species consortium were summarized from three trials. Each isolate participated in 20 four
species combinations/trial, thus N= 60 observations.
Potential interactions between different bacterial taxa using a network based approach
Bacterial diversity based on 16S rRNA gene sequencing of these four DW biofilm communities was
revealed in a previous study (Raghupathi et al., 2017). Significant pairwise interactions (p < 0.01)
between different bacterial genera from these four DW samples were analysed. The type of interaction
i.e. positive correlation hypothetically indicates symbiosis, mutualism or commensalism and negative
correlation hypothetically indicates mutual exclusions, competition or parasitism (Roggenbuck et al.,
2014). It was found that in DW1 and DW4, the numbers of positive correlations were higher than in
DW2 and DW3 (Fig 4). The interaction networks within different bacterial genera identified in this
study are presented (Fig S2, Supplementary information). The genera Pseudomonas and Acinetobacter
had highest numbers of positive correlations suggesting a potential to co-exist with other bacterial
genera.
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Figure 4: Network based analysis showing the number of co-occurrences and mutual exclusion
interactions among bacterial genera identified in the four dishwasher systems generated based on
Spearman correlation analysis.
Bacterial-Fungal biofilm development
E. dermatitidis is known for its dominant presence in household DWs (Zalar et al., 2011; Zupančič et
al., 2016). Therefore, its establishment within bacterial biofilms was investigated. Different four-
species bacterial consortia from DW4 were tested for their ability to incorporate E. dermatitidis (See
Table S3, Supplementary information). We found that two four-species bacterial consortia increased in
its overall biofilm production when E. dermatitidis was included. One bacterial consortium
(Consortium1) was composed of Acinetobacter junii (EXB-L-1308), Haematomicrobium sanguinis
(EXB-L-1326), Bacillus cereus (EXB-L-1176) and Exiguobacterium aestuarii (EXB-L-1327). The
other bacterial consortium (Consortium2) was composed of Acinetobacter junii (EXB-L-1308),
Bacillus cereus (EXB-L-1176), Brevibacterium casei (EXB-L-1336), and Exiguobacterium aestuarii
(EXB-L-1327). It should be noted that the bacterial consortium 1 when present alone increased in cell
numbers over time, however, the bacterial cell numbers reduced overtime in Consortium 1 in the
presence of fungal cells. Consortium 2 showed no change in bacterial numbers and fungal numbers
increased overtime. These results indicate a shift in population dynamics that could be observed due to
resource competition and complex interactions between different microbial species. Interestingly, E.
dermatitidis did not form biofilm when grown as fungal monocultures as it did not attach well to the
surface of the Calgary biofilm device (CBD) indicated by its low cell numbers. However, when E.
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dermatitidis was introduced to multispecies bacterial biofilm, the cell numbers increased leading to the
formation of the trans-kingdom biofilm (Fig 5).
Figure 5: Establishment of fungal cells into bacterial consortia. Log10 CFU counts obtained after 24
and 120 hours of incubation at 25°C harvested from biofilms formed on the wells using flow
cytometry. Consortium C1 were composed of A. junii, B. cereus, H. sanguinis & E. aestuarii species
and consortium C2 were composed of A. junii, B. cereus, B.casei & E. aestuarii species. C1 and C2
denote the total CFU counts from biofilm formed by the two 4-species bacterial consortia with no
fungal addition and not of individual isolates within the consortia. ‘C1+F’ and ‘C2+F’ denote the
counts of total bacterial and E. dermatitidis cells when these consortia were co-cultured with the fungi
E. dermatitidis. ‘F’ denotes the total cell counts of E. dermatitidis when present to form monospecies
fungal biofilm. The error bars denote the mean cell counts ± S.E from three biological trials
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123
Industrial implications
Synthetic surfaces in many machines and equipment, including household appliances and medical
utensils, may become established with microbial biofilm overtime. This could contribute to risks
associated with cross-contamination. As an applied aspect of this study, we wanted to assess the
establishment and colonization of bacterial-fungal biofilms on different elastomer (EPDM) and
polypropylene (PP) surfaces using bacterial Consortium 1 together with E. dermatitidis. This
multispecies bacterial–fungal biofilm was best formed on elastomer 18, which constitute as the actual
rubber material currently used in the industry as rubber seals. Biofilms were less successfully
established on elastomer type 17 and 19 (Fig 6A). E. dermatitidis grown as a mono-species fungal
biofilm also showed an increased attachment to elastomer 18 compared to elastomer 17 and 19 (Fig
6A). Thus, elastomer 18 represents a preferred surface for microbial biofilm formation. However,
based on the absorbance measurements from microbial biomass formed on different PP surfaces; our
observation point to PP surfaces providing an even better surface for microbial attachment (Fig 6B)
than elastomers. Similar results were observed on bacterial-fungal biofilms using bacterial isolates in
Consortia 2 (Supplementary information, Fig S3).
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124
Figure 6: Microbial biofilm formation on EPDM and PP materials. A) Biofilm establishment on three
EPDM rubber types after 24, 48 and 120 hours of incubation at 25 °C; B) Biofilm establishment on
three PP types after 24, 48 and 120 hours of incubation at 25 °C. The biofilm establishement were
absorbance (OD590) measurements quantified by 1% CV staining. The error bars denote the mean
biofilm formation ± S.E from three biological trails.
Discussion
Survival of microorganisms in extreme environments is often associated with formation of complex
biofilms attached on a suitable surface (Davey & O’Toole, 2000). In domestic environments, biofilms
were examined in tap water supply systems (Mullis & Falkinham, 2013; Lührig et al., 2015;
Burkowska-But et al., 2015; Ling et al., 2016; Iakhiaeva et al., 2016; Novak Babič et al., 2016) and in
wet niches such as shower heads (Abe et al., 2016). In this study, we focussed on the isolation of
microorganisms from biofilms formed on rubber seals of four dishwashers (DWs) and used these
isolates to determine their biofilm forming abilities in-vitro. Bacterial communities that colonised the
rubber seals of DWs comprised a wide variety of environmental bacterial species together with a
number of species represented as opportunistic pathogens.
The composition of the microbial communities differed considerably among the four DWs. Microbial
species obtained from these samples were well represented based on their abundance levels at their
genus level (Raghupathi et al., 2017). These results show that most abundant microbial representatives
identified by sequencing approach, remained viable in these extreme systems. The dominant bacterial
genus, Exiguobacterium was isolated in 3 out of 4 DWs. Different species of this genus were known
for their ability to proliferate in extreme natural environments like hot, alkaline and marine
environments (Vishnivetskaya et al., 2009). Another highly represented bacterial genus was Bacillus.
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125
Bacillus is ubiquitous in domestic environments (Gorny et al., 1999; Park et al., 2006) and is widely
used in industry as a microbial indicator for cleaning procedures during washing cycles (Ståhl
Wernersson et al., 2004; Lee et al., 2007; Nicolella et al., 2011). The diversity of fungi isolated from
DWs was in accordance with previous studies (Zalar et al., 2011; Döğen et al., 2013; Gümral et al.,
2015; Zupančič et al., 2016).Black yeasts, E. dermatitidis and E. phaeomuriformis were represented in
most dishwashers; followed by white yeasts, Candida parapsilosis and red yeasts, Rhodotorula
mucilaginosa. These four fungal species were classified as opportunistic human pathogens (Trofa et
al., 2008; Silva et al., 2012; Lunardi et al., 2006; De Almeida et al., 2008; Hiruma et al., 1993;
Sudhadham et al., 2008; Russo et al., 2010; Kondori et al., 2011) and with their presence in household
DWs, they could represent a potential source for indoor infections (Zupančič et al., 2016).
Bacterial interactions play a major role in shaping and maintaining the diversity within bacterial
communities (HilleRisLambers et al., 2012) and also influence the balance between cooperating and
competing phenotypes (Nadell et al., 2016). Studies have elucidated the coexistence patterns among
microbial groups from a variety of ecosystems using microbial correlation networks (Eiler et al., 2012;
Kittelmann et al., 2013; Zhalnina et al., 2013). However, little is known on whether these coexistence
patterns reflect the actual biogenic relationships and interactions in-situ. In this study, we analysed the
co-occurrence patterns between the different bacterial taxa in the four DW samples. Positive and
negative correlations of bacterial taxa were accounted to the genus level. DW1 and DW4 had higher
number of positive correlations compared to DW2 and DW3. Also, when screened for synergistic
multispecies biofilm, it was found that DW1 and DW4 had higher numbers of four-species
combinations interacting synergistically leading to an overall increase in biomass.
Biofilm levels of the 4-species consortia when further examined and compared to the levels of biofilm
production of each isolate under monospecies conditions, it was revealed that P. aeruginosa and A.
junii, isolated from DW1 and DW4, respectively, were found to contribute as best biofilm producers
that included poor or non-biofilm producing isolates, increasing the overall biofilm formation within
the included consortia. Likewise, co-association networks revealed that the genera Pseudomonas and
Acinetobacter had higher number of positive correlations suggesting a potential to cooperate with
other bacterial genera. These observations could support our previous evidence where DW1 and DW4
had higher percentage of synergistically interacting 4-species biofilm. Likewise, the co-association
networks revealed that the genera Exiguobacterium and Micrococcus had higher numbers of negative
correlations signalling competition or exclusion to other bacteria. In DW2 and DW3, most
combinations included isolates belonging to the genera Exiguobacterium and Micrococcus and the
observed number of synergistic 4-species consortia were much lower than what was seen in DW1 and
DW4. Interestingly, these findings demonstrate an observed trend between the correlation detection
technique and in-vitro multispecies biofilm assessments, where co-occurrence of bacterial members
within these ecological systems could contribute to multispecies biofilm formation.
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126
Synergy impacts bacterial composition in multispecies biofilms and their overall biomass (Burmølle et
al., 2014). Such multispecies biofilms are tolerant against antimicrobials compared to their
monospecies equivalents (Lee et al., 2014; Schwering et al., 2013). We have characterised the
interactions within different bacterial species and how they impact each other during biofilm
development, both under mono and mixed species cultures. Later, the ability of selected mixed
bacterial consortia to incorporate the polyextremophile E. dermatitidis, (Zalar et al., 2011; Poyntner et
al., 2016) the prevalent fungal species in DW systems, was assessed. Fungi and bacteria play
important roles in promoting the survival of their interacting partners (Frey-Klett et al., 2011). Such
complex biofilms can be beneficial to all microbial partners, but can be detrimental to the human host
(Ghannoum, 2016, Kalan et al., 2016; Minerdi et al., 2008). Our results show that when bacterial
consortia were supplemented with E. dermatitidis, the biomass production and the numbers of bacteria
were stimulated together with the growth of fungal partner in the mixed biofilm. The observations that
the bacterial community of DWs facilitating the growth of an opportunistic pathogenic fungus and
mixed bacterial-fungal biofilm established on commonly used industrial surfaces (EPDM 18 and PP)
complementing their persistence and growth; represent significant findings with scientific and applied
implications. Though these observations are similar to the results obtained in other studies
investigating mixed species biofilms like Candida albicans an opportunistic pathogenic fungi (Pammi
et al., 2013; Seneviratne et al., 2007; Harriott and Noverr, 2011), it should be noted that the studies
were made using one fungi and single bacteria in co-cultures; whereas, in this study, we present the
establishment of an opportunistic black yeast pathogen into mixed bacterial consortium comprising of
four species. Further, the formation of bacterial and fungal biofilms on dishwasher related
environments emphasize the importance of interactions played between different microbial species and
their change in population dynamics across kingdoms during biofilm development.
In summary, our main findings include the existence of synergistic interactions observed during
biofilm formation between bacteria isolated from different DWs where, A. junii and P. aeruginosa
were recognized as the best biofilm producers and important contributors to synergy in biofilm. This
finding corresponds with network based co-occurrence analysis where these two bacterial genera in
dishwasher systems, account to most positive correlations observed. In addition, mixed bacterial
biofilms could incorporate the opportunistic yeast pathogen, E. dermatitidis and facilitate its
establishment on rubber seals and other surfaces. The enhancement of trans-kingdom biofilm
formation on rubber used in DWs suggests that microbes surviving these environments have been
selected by their ability to engage in synergistic biofilm formation. With our study, we have shown
that our experimental model has the capacity to reveal new and unique features of these complex and
dynamic microbial communities. Additionally, our observations and methodology could have
important implications for future design and maintenance of house-hold and medical appliances, as
these systems could present as a source of domestic cross-contamination and human infections.
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127
Acknowledgements
Our acknowledgements go to all the people who kindly provided samples from their dishwashers. We
also thank Karin Vestberg for her assistance with NGS and prof. Børge Diderichsen for careful and
critical reading of the manuscript.
Ethics approval and consent to participate
In this study, field sampling was performed, and to our knowledge, no endangered or protected species
were involved. All of the samples studied here were obtained from the discussed sampling areas, for
which permission was obtained from the owners.
Conflict of Interest
The authors declare that they have no conflicts of interest.
Funding
This research was funded by the Ministry of Higher Education, Science and Technology of the
Republic of Slovenia, as a Young Researcher grant to JZ (grant no. 382228-1/2013). We also thank
the Slovenian Research Agency (Infrastructural Centre Mycosmo, MRIC UL) and the Danish Council
for Independent Research grant: 1323 00235 for providing financial support. The funding bodies had
no influence on the design of the study and collection, analysis, and interpretation of data and in
writing the manuscript.
Authors’ contributions
SJS, NGC and MB designed the study. JZ and PKR performed the experiments. JZ and PKR analysed
the data. JZ, PKR, KH, NGC, MB, and SJS compiled the manuscript.
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Supplementary information
Table S1: Bacterial and fungal isolates obtained as pure cultures from DW associated biofilm formed
on 1 cm2 rubber seals. 16S rRNA 16S ribosomal RNA gene; LSU 26S ribosomal RNA gene; ITS
internal transcribed spacer 1, 5.8S ribosomal RNA gene, and internal transcribed spacer 2; tef,
translation elongation factor 1-alpha (EF1a) gene; FOSC Fusarium oxysporum species complex.
Strain ID represents the isolate identification after deposition (as ‘EXF’ for fungal and ‘EXB-L’ for
bacterial isolates) at the Microbial Culture Collection Ex (MRICUL EX).
Closest Relative Phylum DNA based
identification
method
Strain ID
EXF- / EXB
L-
Accession
number of the
closest relative
DW1
Bacillus cereus Firmicutes 16S rRNA EXB L-1175 KC969074
Bacillus pumilus Firmicutes 16S rRNA EXB L-1225 AJ494726
Bacillus sp. Firmicutes 16S rRNA EXB L-1177 HM989921
Bacillus thuringiensis Firmicutes 16S rRNA EXB L-1173 JX035937
Brachybacterium
paraconglomeratum
Actinobacter
ia
16S rRNA EXB L-1160 FJ172038
Chryseobacterium sp. Bacteroidete
s
16S rRNA EXB L-1165 JN545042
Comamonas testosteroni Proteobacter
ia
16S rRNA EXB L-1141 AY247415
Enterobacter cancerogenus Proteobacter
ia
16S rRNA EXB L-1132 FJ009375
Enterobacter hormaechei Proteobacter
ia
16S rRNA EXB L-1135 KP303395
Enterobacter sp. Proteobacter
ia
16S rRNA EXB L-1129 KM979225
Klebsiella oxytoca Proteobacter
ia
16S rRNA EXB L-1137 CP011636
Kurthia gibsonii Firmicutes 16S rRNA EXB L-1146 AB271738
Leucobacter sp. Actinobacter
ia
16S rRNA EXB L-1152 KC550185
Lysinibacillus fusiformis Firmicutes 16S rRNA EXB L-1140 DQ333300
Ochrobactrum
pseudintermedium
Proteobacter
ia
16S rRNA EXB L-1130 KF026284
Pseudomonas aeruginosa Proteobacter
ia
16S rRNA EXB L-1125 KR911837
Pseudomonas alcaligenes Proteobacter
ia
16S rRNA EXB L-1113 AF390747
Pseudomonas putida Proteobacter
ia
16S rRNA EXB L-1149 KJ735915
Sphingobacterium spiritivorum Bacteroidete
s
16S rRNA EXB L-1227 EF090267
Stenotrophomonas maltophilia Proteobacter
ia
16S rRNA EXB L-1167 KP185140
Candida parapsilosis Ascomycota LSU EXF-9745 KJ481229
Candida pararugosa Ascomycota LSU EXF-9751 GU904205
Clavispora lusitaniae Ascomycota LSU EXF-9744 KF728663
Exophiala phaeomuriformis
genotype 1
Ascomycota ITS EXF-9735 KP034987
Fusarium oxysporum species
complex
Ascomycota tef EXF-9737 KP761169
Meyerozyma guilliermondii Ascomycota LSU EXF-9759 KJ481231
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Rhodotorula mucilaginosa Basidiomyco
ta
LSU EXF-9755 KC442283
DW2
Acinetobacter lwoffii Proteobacter
ia
16S rRNA EXB L-1215 LN774665
Acinetobacter sp. Proteobacter
ia
16S rRNA EXB L-1191 AY486382
Aerococcus sp. Firmicutes 16S rRNA EXB L-1205 EU376006
Bacillus cereus Firmicutes 16S rRNA EXB L-1223 KP988025
Enterobacter sp. Proteobacter
ia
16S rRNA EXB L-1204 KM979225
Pseudescherichia vulneris Proteobacter
ia
16S rRNA EXB L-1211 JQ958880
Exiguobacterium aestuarii Firmicutes 16S rRNA EXB L-1196 FJ462716
Exiguobacterium panipatensis Firmicutes 16S rRNA EXB L-1201 EF519705
Exiguobacterium sp. Firmicutes 16S rRNA EXB L-1196 EU159578
Kocuria rhizophila Actinobacter
ia
16S rRNA EXB L-1199 AY030315
Kocuria salsicia Actinobacter
ia
16S rRNA EXB L-1221 GQ352404
Lactococcus lactis subsp. lactis Firmicutes 16S rRNA EXB L-1213 KR732324
Leclercia sp. Proteobacter
ia
16S rRNA EXB L-1203 JX949970
Micrococcus luteus Actinobacter
ia
16S rRNA EXB L-1190 KF993675
Micrococcus sp. Actinobacter
ia
16S rRNA EXB L-1212 EU379020
Pseudomonas psychrotolerans Proteobacter
ia
16S rRNA EXB L-1186 KM019821
Pseudomonas sp. Proteobacter
ia
16S rRNA EXB L-1220 AM945563
Rothia sp. Actinobacter
ia
16S rRNA EXB L-1189 EU135638
Candida parapsilosis Ascomycota LSU EXF-9760 KJ481228
Exophiala dermatitidis genotype
A
Ascomycota ITS EXF-9487 DQ826738
Exophiala dermatitidis genotype
C
Ascomycota ITS EXF-9463 JF766671
Rhodotorula mucilaginosa Basidiomyco
ta
LSU EXF-9756 KP087899
DW3
Bacillus cereus Firmicutes 16S rRNA EXB L-1263 KC969074
Bacillus circulans Firmicutes 16S rRNA EXB L-1279 KM349203
Exiguobacterium aestuarii Firmicutes 16S rRNA EXB L-1244 FJ462716
Exiguobacterium arabatum Firmicutes 16S rRNA EXB L-1278 JF775422
Exiguobacterium panipatensis Firmicutes 16S rRNA EXB L-1260 EF519705
Exiguobacterium profundum Firmicutes 16S rRNA EXB L-1270 KM873375
Exiguobacterium sp. Firmicutes 16S rRNA EXB L-1269 EU159578
Microbacterium
hydrocarbonoxydans
Actinobacter
ia
16S rRNA EXB L-1250 JQ954857
Microbacterium sp. Actinobacter
ia
16S rRNA EXB L-1272 FR774577
Micrococcus luteus Actinobacter
ia
16S rRNA EXB L-1261 KJ733861
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DW4
Acinetobacter junii Proteobacter
ia
16S rRNA EXB-L-1308 EU862296
Acinetobacter sp. Proteobacter
ia
16S rRNA EXB-L-1324 EU705470
Haematomicrobium sanguinis Actinobacter
ia
16S rRNA EXB-L-1326 EU086805
Bacillus amyloliquefaciens Firmicutes 16S rRNA EXB-L-707 JX036499
Bacillus cereus Firmicutes 16S rRNA EXB-L-1176 GU568201
Bacillus horneckiae Firmicutes 16S rRNA EXB-L-1313 FR749913
Brachybacterium
paraconglomeratum
Actinobacter
ia
16S rRNA EXB-L-1311 FJ172038
Brevibacillus sp. Firmicutes 16S rRNA EXB-L-1330 GQ927158
Brevibacterium casei Actinobacter
ia
16S rRNA EXB-L-1336 HM012705
Brevibacterium sanguinis Actinobacter
ia
16S rRNA EXB-L-1305 AJ564859
Exiguobacterium aestuarii Firmicutes 16S rRNA EXB-L-1327 FJ462716
Exiguobacterium panipatensis Firmicutes 16S rRNA EXB-L-1316 EF519705
Exiguobacterium profundum Firmicutes 16S rRNA EXB-L-1335 KF269103
Exiguobacterium sp. Firmicutes 16S rRNA EXB-L-1331 EF519705
Microbacterium paraoxydans Actinobacter
ia
16S rRNA EXB-L-1310 DQ350825
Staphylococcus saprophyticus Firmicutes 16S rRNA EXB-L-1314 AB697718
Candida parapsilosis Ascomycota LSU EXF-9764 EU056283
Exophiala dermatitidis genotype
A
Ascomycota ITS EXF-9777 DQ826738
Exophiala dermatitidis genotype
A2
Ascomycota ITS EXF-9778 FJ387565
Exophiala phaeomuriformis
genotype 1
Ascomycota ITS EXF-9779 KP034987
Rhodotorula mucilaginosa Basidiomyco
ta
LSU EXF-9762 AF335986
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Figure S1: Heat map of most abundant (A) bacterial and (B) fungal genera identified by 16s rRNA
and ITS gene based sequencing done in a previous study (Raghupathi et al., 2017). The scaled heat
maps were generated using log-transformed bacterial and fungal abundances and clustered based on
‘coniss’. The heatmaps were created using various R packages: gplots, vegan, rioja and Rcolorbrewer
available for Rgui 3.2.0.
ChryseobacteriumDysgonomonasXanthobacterAzospiraBrevibacteriumPseudomonasStenotrophomonasOchrobactrumRoseomonasRhodoplanesCellulosimicrobiumGordoniaAcinetobacterParacoccusBrevundimonasRhizobiumMycobacteriumPaenibacillusLegionellaPeredibacterBdellovibrioPseudoxanthomonasRhodobacterExiguobacteriumBacillusMicrococcusStaphylococcusAerococcusKocuriaRothiaEnterococcusLactococcusEscherichia/ShigellaPetrobacterNesterenkonia
DW1 DW2 DW3 DW4Trichosporon
Clavispora
Candida
Pichia
Cryptococcus
Fusarium
Acremonium
Sporobolomyces
Saccharomyces
Alternaria
Aspergillus
Aureobasidium
Rhodosporidium
Filobasidium
Penicillium
Debaryomyces
Cladosporium
Wallemia
Exophiala
Rhodotorula
DW1 DW2 DW3 DW4
A B
-3 -1 1 3
Relative abundance
-3 -1 1 3
Relative abundance
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Figure S2: Bacterial co-occurrences by network analysis and its parameters. The type of interaction of
one bacterial genus to the other bacterial genera from network based analysis and the network
parameters. ‘Green’ connectors indicate ‘positive correlations’ signalling cooperation and ‘red’
connectors indicate ‘negative correlations’ signalling mutual exclusion. The significant networks (p <
0.01) were generated using log-transformed bacterial abundance and bacterial taxa that were present in
sample (n > 2), classified to the genus level and represented in the isolate collection (Table 1) of
dishwashers.
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Figure S3: Microbial biofilm formation of Consortia 2 on EPDM and PP materials. A) Biofilm
establishment on three EPDM rubber types and B) Biofilm establishment on three PP types after 24,
48 and 120 hours of incubation at 25 °C. The biofilm establishedment were absorbance (OD590)
measurements quantified by 1% CV staining. The error bars denote the mean ± S.E.M from three
biological trails.
24 48 120 24 48 120 24 48 120-0.2
0.0
0.2
0.4
0.6
0.8
1.0
A
EPDM 17 EPDM 18 EPDM 19
Bacterial biofilmBacterial-Fungal biofilmFungal biofilm
Time (h)
Abs
590
24 48 120 24 48 120 24 48 120-0.2
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
Bacterial biofilm
Bacterial-Fungal biofilm
Fungal biofilm
B
Time (h)
Abs
590
PP Type 1 PP Type 2 PP Type 3
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Table S2: Biofilm formed by single and combinations of 4-species observed in samples DW1, DW2,
DW3 and DW4. Biofilm quantification in 10%LB was done after 24 hours of incubation at 25°C by
crystal violet staining and absorbance measured at 590 nm. Fd is the ratio of (Abs 590 multispecies
biofilm – Std Err) / (Abs 590 best single species+ Std Err) or (Abs 590 multispecies biofilm + Std Err)
/ (Abs 590 best single species - Std Err) calculated across the three biological trials
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Table S3: Biofilm formed by single and combinations of four species isolated from sample DW4. Biofilm quantification was done after 24 and 120 hours of
incubation at 25°C by crystal violet staining and absorbance measured at 590 nm. Fd1 is the ratio of (Abs 590 multispecies bacterial biofilm – St Err) / (Abs
590 best single bacterial species + St Err) or (Abs 590 multispecies bacterial biofilm + St Err) / (Abs 590 best single bacterial species - St Err). Fd2 is is the
ratio of (Abs 590 multispecies bacterial-fungal biofilm – St Err) / (Abs 590 best single bacterial species co-cultured with fungi + St Err) or (Abs 590
multispecies bacterial-fungal biofilm + St Err) / (Abs 590 best single bacterial species co-cultured with fungi - St Err). Fd3 > 1 (Fd3 =Fd2/Fd1) determines the
total biofilm induction in the presence of E. dermatitidis cells. The combinations 22, 23, 24, 27 (Consortia 1) and 22, 24, 25, 27 (Consortia 2) had Fd3 value >
1 in all the three biological trails signifying an increase in the biofilm production due to the addition of E. dermatitidis cells. These two consortia were further
investigated for bacterial and fungal cell number quantification and biofilm formation on EPDM and PP materials.
142
Raghupathi PK, Liu W, Sabbe K, Houf K, Burmølle M, Sørensen SJ. Synergistic Interactions within
a Multispecies Biofilm Enhance Individual Species Protection against Grazing by a Pelagic Protozoan.
Front Microbiol. 2018 Jan 9; 8: 2649. Doi: 10.3389/fmicb.2017.02649.
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Synergistic interactions within a multispecies biofilm enhance individual species protection
against grazing by a pelagic protozoan
Prem K Raghupathi1, 3
, Wenzheng Liu3, Koen Sabbe
2, Kurt Houf
1, Mette Burmølle
3, Søren J.
Sørensen3
1Department of Veterinary Public Health and Food Safety, Faculty of Veterinary Medicine, Ghent
University, Merelbeke, Belgium; 2 Laboratory of Protistology and Aquatic Ecology, Department of
Biology, Faculty of Sciences, Ghent University, Ghent, Belgium; 3Section for Microbiology,
Department of Biology, University of Copenhagen, Universitiesparken, Copenhagen, Denmark.
Biofilm formation has been shown to confer protection against grazing, but little
information is available on the effect of grazing on biofilm formation and protection in
multispecies consortia. With most biofilms in nature being composed of multiple bacterial
species, the interactions and dynamics of a multispecies bacterial biofilm subject to grazing by a
pelagic protozoan predator were investigated. To this end, a mono and multispecies biofilms of
four bacterial soil isolates, namely Xanthomonas retroflexus, Stenotrophomonas rhizophila,
Microbacterium oxydans and Paenibacillus amylolyticus, were constructed and subjected to
grazing by the ciliate Tetrahymena pyriformis. In monocultures, grazing strongly reduced
planktonic cell numbers in P. amylolyticus and S. rhizophila and also X. retroflexus. At the same
time, cell numbers in the underlying biofilms increased in S. rhizophila and X. retroflexus, but
not in P. amylolyticus. This may be due to the fact that while grazing enhanced biofilm formation
in the former two species, no biofilm was formed by P. amylolyticus in monoculture, either with
or without grazing. In four-species biofilms, biofilm formation was higher than in the best
monoculture, a strong biodiversity effect that was even more pronounced in the presence of
grazing. While cell numbers of X. retroflexus, S. rhizophila and P. amylolyticus in the planktonic
fraction were greatly reduced in the presence of grazers, cell numbers of all three species
strongly increased in the biofilm. Our results show that synergistic interactions between the
four-species were important to induce biofilm formation, and suggest that bacterial members
that produce more biofilm when exposed to the grazer not only protect themselves but also
supported other members which are sensitive to grazing, thereby providing a “shared grazing
protection” within the four-species biofilm model. Hence, complex interactions shape the
dynamics of the biofilm and enhance overall community fitness under stressful conditions such
as grazing. These emerging inter- and intra-species interactions could play a vital role in biofilm
dynamics in natural environments like soil or aquatic systems.
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Introduction
In recent years, protozoa-bacteria interactions have received increasing attention in studies ranging
from ecology to consumer health and diseases. Free-living protozoa are commonly found in natural
environments like soils and aquatic habitats (Ekelund et al., 2001; Foissner, 1999; Pernthaler, 2005;
Pfister et al., 2002) and in anthropogenic environments like swimming pools (Rivera et al., 1993),
drinking water systems (Thomas & Ashbolt, 2011), kitchens (Chavatte et al., 2014) and health care
facilities (Cateau et al., 2014; Singh & Coogan, 2005). Various studies have also reported the
presence of bacterial biofilms in such environments (Besemer et al., 2012; Bryers, 2008; Burmølle et
al., 2011). Though most studies emphasize that the main role played by the protozoa lies in control of
the bacterial populations by predation (Arndt et al., 2003; Brown & Barker, 1999; Jürgens & Güde,
1994; Logares et al., 2012), another potential impact involves the induction of biofilm formation by
bacterial communities (Joubert et al., 2006) to avoid grazing.
Biofilm formation represents a surface attached mode of life (Donlan, 2002) that can contain multiple
species of archaea, bacteria, fungi and algae (Flemming et al., 2016). Biofilms offer physical
protection through the secreted polymeric matrix (Joubert et al., 2006) creates a protective
microhabitat against predation (DePas et al., 2014, Darby et al., 2002; Matz et al., 2005). Close
interactions between bacteria and protozoa in biofilms are also thought to give rise to a series of
adaptations in bacterial communities by promoting horizontal gene transfer events, quorum sensing
abilities and induce bacterial protein secretion systems (Darby et al., 2002; Matz et al., 2004)
enhancing their survival, dynamics and coexistence (Matz & Kjelleberg, 2005).
Grazing by protozoa has been reported to stimulate micro-colony formation, alter mass transfer of
nutrients and induce biofilm development by stimulating bacterial layer thickness (Böhme et al.,
2009; Kaminskaya et al., 2007; Matz et al., 2004; Weitere et al., 2005; Wey et al., 2008). Other
studies however argue that protozoa do not induce biofilm formation (Huws et al., 2005) but instead
show a marked preference for grazing on attached or aggregated bacterial cells or only change biofilm
community structure (Caron, 1987; Huws et al., 2005; Sibbald & Albright, 1988; Wey et al., 2008).
Furthermore, studies have also shown that the grazed or consumed bacterial cells can become adapted
to resist uptake or digestion and are even capable of intracellular replication within the protozoan host
cells (Lambrecht et al., 2015; Rowe & Grant, 2006; Taylor et al., 2009). Although feeding
interactions between protozoa and planktonic bacteria are well understood (Jürgens & Matz, 2002;
Matz & Jürgens, 2005; Roberts et al., 2011) only few studies have attempted to assess the impact of
grazing on biofilms at the multi-bacterial level. In multispecies biofilm settings, interactions between
different bacteria play an important role in determining the structure, function and dynamics of the
biofilms and it has been suggested that they contribute to defence mechanisms of bacterial biofilms
against predators (Matz, 2011; Wey et al., 2008). Moreover, it has been shown that interspecific
interactions within the mixed bacterial communities in the presence of a grazing protist promoted co-
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aggregation of bacterial members and enhanced complex biopolymer degradation pathways leading to
an overall increase in carbon transfer efficiency (Corno et al., 2013, 2015). Mixed biofilms have in
other cases been shown to offer the harboured species protection against antibacterial compounds and
enhanced capabilities of invasion and virulence within host organisms (Burmølle et al., 2016).
Different protozoan members have different impact on the microbial communities (Brown & Barker,
1999; Paisie et al., 2014). In soils, protozoa present themselves as a diverse group of flagellates,
ciliates and naked amoebae (Bonnet et al., 2005; Ekelund & Rønn, 1994). Like flagellates, ciliates
display a substantial diversity in motility, morphology and feeding strategies (Dopheide et al., 2011)
and are considered to be important predators of bacteria. Hence, there is a need to unravel different
prey-predator interactions and their impact on mixed species bacterial biofilm, as mixed biofilms are
the predominant lifestyle in most ecosystems (Battin et al., 2003; Costerton, 2007; Mielich-Süss &
Lopez, 2015). Grazing on diverse biofilms is likely to shape the existing complex interactions within
the biofilm communities (Hansen et al., 2017; Wen et al., 2010) or alter the feeding traits of protozoa.
Examples include Gram negative bacteria being more vulnerable to grazing than Gram positive
bacteria (Rønn et al., 2002) or altered feeding responses of protozoa to one bacterial group over
another (Dopheide et al., 2011).
The aim of the present study is to assess whether individual biofilm bacterial species gain enhanced
protection by other members in multispecies consortia under grazing pressure. Therefore, we
examined the effect of grazing by the ciliate T. pyriformis on biofilm formation and population
dynamics in a consortium composed of four bacterial soil species X. retroflexus, S. rhizophila, M.
oxydans and P. amylolyticus. These four strains when combined have been shown to act
synergistically resulting in increased biofilm development (Ren et al., 2015). Ciliates were shown to
be effective bacterial grazers with often extremely high ingestion rates (Iriberri et al., 1995), making
them a specialised subgroup within the protist (Parry, 2004). Under such extreme grazing pressure, we
hypothesize that multispecies biofilms will generate a protective effect compared to single species
biofilms. We used a qPCR protocol developed previously for these model consortia (Ren et al., 2014)
to quantify the species-specific impact of protozoan grazing.
Materials and Methods
Soil isolates and Protozoa culture conditions
The bacterial species Xanthomonas retroflexus (JQ890537), Stenotrophomonas rhizophila
(JQ890538), Microbacterium oxydans (JQ890539) and Paenibacillus amylolyticus (JQ890540) stored
in the culture collection of the Section of Microbiology, University of Copenhagen, were subcultured
from frozen glycerol stocks onto TSA plates (Tryptic Soy Agar, Sigma-Aldrich, Germany). The plates
were incubated at 24°C for 48 hours. Single colonies were inoculated into 5 ml TSB (Tryptic Soy
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Broth, Sigma-Aldrich, Germany) media and incubated with shaking at 180 rpm for 24 h at 24 °C
when required. These strains were used as bacterial prey for the protozoan predator.
A Tetrahymena pyriformis (Tp) culture (Culture Collection of Algae and Protozoa, CCAP nr
1630/w1) was provided by Department of Veterinary Public Health and Food Safety, Ghent
University, Belgium. Axenic cultures of this protozoan were maintained in 25 cm2 culture flasks with
20 ml PPY medium [proteose peptone yeast extract; 20 g Proteose peptone (Merck KgaAm
Germany), 2.5 g yeast extract (Merck KgaAm Germany) in 1L H2O; autoclaved]. Weekly
maintenance of the ciliate cultures at 24 °C was done by aseptically transferring 5 ml of the culture
into 15 ml fresh PPY medium incubated. For biofilm grazing experiments, T. pyriformis cells in
exponential phase (after 48 h at 24°C) were washed twice in PAS (Page’s amoeba saline) solution
followed by centrifuging at 850g after which the cells were re-suspended into 10 ml TSB media.
Biofilm cultivation and grazing experiments
Biofilm cultivation experiments were performed in 96-well cell culture plates (cat. no. 655180,
Grenier Bio-one, Germany). The four selected strains were screened for biofilm formation as single
species (monospecies) and in three/four-species combination (multispecies) as described (Ren et al.,
2015) both in the presence and absence of protozoa. Briefly, bacterial cell cultures in exponential
growth phase (OD600 between 0.3 - 0.6) were selected and adjusted to a start OD600 of 0.15 in TSB
media for all cultures. For monospecies biofilms, aliquots of 150 µl of cell culture and for three- and
four-species biofilms, respectively, 50 or 37.5 µl of each bacterial strain were added into the wells so
that the final inocula were 150 µl in all the settings. To the wells that were to be grazed, an volume of
1.5 µl containing ~approx. 1000 cells T. pyriformis cells in TSB media were added. The plates were
incubated at 24°C for 12, 24 and 96 h. Wells containing only 150 µl TSB media and TSB media with
T. pyriformis cells served as blank/control. Three wells each time served as one technical replicate and
this was repeated at five different times.
Quantification of biofilm and planktonic fractions
Biofilm formation was assayed and quantified using the traditional crystal violet (CV) method as
previously described (Ren et al., 2014). The biofilm attached to the wells was then washed twice
gently with 160 µl 1X PBS (phosphate buffer saline) solution and stained with 180 µl 1% (w/v) CV
solution. After 20 min of staining, the CV solution was removed by pipette, and the stained biofilm
was gently washed five times with 200µl PBS solution. The remaining CV dye retained by the biofilm
was de-stained into 200 µl 96% ethanol for 30 minutes. Biofilm formation was then quantified by
measuring the absorbance of de-stained CV at 590nm using EL340 BioKinetics reader (BioTek
Instruments, USA) and expressed as biofilm forming index (BFI) according to the equation BFI=(AB-
CW)/G (Niu & Gilbert, 2004) where, AB: OD590 of attached microorganisms, CW: OD590 control
wells and G: OD600 of cells in planktonic fraction. Biodiversity (BD) effect was calculated as the
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difference between the observed biofilm yield (biofilm of mixed cultures) and the expected yield
(average of the monoculture yields) (Loreau & Hector, 2001; Vanelslander et al., 2009). Biofilm fold
(Fd) i.e. the observed increase in biofilm formation due to grazing is the ratio between OD590 of grazed
three-species biofilm and OD590 of non-grazed three-species biofilm.
Quantification of the biofilm and planktonic fractions was performed by plating. 100 µl of the
planktonic fraction from the wells after 24 and 96 h incubation was suspended in 900 µl 1X PBS
solution. Once the planktonic fractions were removed, the wells with attached biofilm were gently
washed twice with 160 µl 1X PBS solution. The wells were then filled with 200 µl 1X PBS and
mixed thoroughly by pipetting. Serial dilutions in 900 µl 1X PBS were performed and 100 µl of the
dilutions were plated onto TSA plates by spread plating after which the plates were allowed to dry
completely at room temperature. Drying restricts the movement of T. pyriformis on plates. The plates
were then incubated for 48 hours at 24° C. Single colonies formed after incubation were counted and
the results were calculated in CFU (colony forming units). Grazing fold i.e. the percentage reduction
in planktonic fraction due to grazing was expressed by 100 × [CFU (culture) – CFU (culture+Tp) / CFU
(culture)]. Changes in cell counts from biofilm fraction were expressed by log (CFU (culture+Tp) / CFU
(culture)).
Ciliate growth on bacterial cultures
We determined the ciliate numbers of T. pyriformis grown on the four bacteria separately
(monospecies) and as a mixture (four-species) for up to 96 h at regular intervals in microtiter plates.
The protozoa cells were counted using a Sedgewick-Rafter chamber and an inverted microscope (40X
magnification) as described previously (Gittleson, Stephen M and Ganapathy, 2011) with minor
modifications. The wells containing the suspension of bacteria and protozoa were homogenized by
pipetting and 150µl of the cell suspension was fixed in 1% (w/v) Lugol’s iodine solution to a final
volume of 1.2 ml in dH2O. The contents were then immediately transferred to the counting chamber
and the cells were allowed to settle for few minutes. The change in protozoa cell numbers over time
was expressed using ∆N = log10 (Nt – N0) / t. To visualize the changes in protozoa numbers over time
in co-culture with bacteria, 50µl spots of the fixed suspension were made on glass slides and
micrographs were taken at different time points using Zeiss Axioplan II, Carl Zeiss with a 10X
objective.
16S rRNA based fluorescent in-situ hybridization (FISH) and confocal imaging to investigate
grazing
To visualize the effects of grazing by the protozoan and the internalization of bacteria within the food
vacuoles of T. pyriformis, FISH was performed with 16S rRNA gene probes targeting the specific
bacteria (Liu et al., 2017). 50 μl spots of co-culture suspensions (bacteria and protozoa) after 24 h
were collected after thorough pipetting to homogenize the suspension. The collected cells were then
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left to air dry on a glass slide. The above step was repeated 5 times (5 × 50 µl) with the aim to collect
more cells. The attached cells were coated with 0.5% (w/v) agarose by immersing the slides into a
tube containing 45 ml molten agarose and fixed using 4% PFA (paraformaldehyde) at 4 °C. Samples
were dehydrated and the hybridization protocol was performed according to (Amann, 1995; Daims,
2009) with 30% formamide concentration. After hybridization, the slides were washed in cold water
and dried at room temperature. The slides were stored in the dark and visualized under confocal
microscopy (Point-scanning confocal and multiphoton microscope SP5-X MP, Leica Microsystems).
Images were processed using Leica Application Suite X.
qPCR quantification of bacterial cell numbers in multispecies setting
The biofilm formation assay was conducted both in the presence and absence of T. pyriformis in 96-
well microtiter plates as described above. After 24h, the planktonic fractions were collected in
Eppendorf tubes and the biofilm fraction was rinsed twice with weak phosphate buffer to remove
loosely attached cells. Three replicate wells were prepared for each treatment. The cell numbers of
the four strains in multispecies planktonic and biofilm fractions with and without protozoa were
quantified by SYBR Green qPCR using standard curves generated by serial 10-fold dilutions of
plasmid DNA using the species specific primers and thermal profile setup previously reported (Ren et
al., 2014). All samples were run in triplicate and a no template control was included in each run.
Bacterial DNA was extracted using FastDNA™ SPIN Kit for soil (MP Biomedicals, Germany)
according to manufacturer’s instruction.
Results
T. pyriformis grazing promotes biofilm formation and reduces the number of bacteria in the
planktonic fractions
Monocultures and four-species mixed cultures of X. retroflexus, S. rhizophila, M. oxydans and P.
amylolyticus were tested for biofilm formation in the absence and presence of protozoa (Fig. 1). T.
pyriformis grazing on monospecies cultures of X. retroflexus and S. rhizophila resulted in significantly
enhanced biofilm formation (paired t-test, P<0.05) whereas M. oxydans and P. amylolyticus
monocultures did not form biofilms neither in the presence nor in the absence of T. pyriformis.
Biofilm formation was enhanced in the four-species mixtures, and was even more strongly induced in
these mixtures in the presence of grazing for up to 96 h (n=5, paired t-test, P<0.05), suggesting a
strong biodiversity effect (Fig. S1, supplementary information). Moreover, biofilm formation in the
mixtures was higher than in the best performing monoculture both in the absence and presence of
grazing.
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Figure 1: Biofilm forming index (BFI) of mono and mixed species cultures subject to T. pyriformis
(Tp) grazing and non-grazed cultures at 12, 24 and 96 h. The data points indicate the biofilm mean ±
standard error of the mean (SEM) obtained from five biological replicates.* P<0.05 **P<0.01.
Planktonic fractions of three of the four bacterial species were effectively grazed upon in
monoculture, but less so in the four-species co-culture. P. amylolyticus was the most intensively
grazed species at 24 h, whereas after 96 h S. rhizophila monocultures were the most highly grazed
followed by P. amylolyticus and X. retroflexus monocultures. Among all the strains, M. oxydans was
the least preferred prey, and S. rhizophila and P. amylolyticus were the most favored prey (Fig. 2).
These grazing experiments verified the ability of T. pyriformis to feed on planktonic bacteria. In the
four-species mixed cultures, overall grazing by T. pyriformis on the planktonic community was
reduced compared to the monospecies cultures observed by the low grazing fold values at 24h and
96h (Fig. 2).
12 24 96 12 24 96 12 24 96 12 24 96 12 24 96
-2
0
2
4
6
8
X. retroflexus S. rhizophila
M. oxydans P. amylolyticus
4-species mix
** **
**
* * *
**
**
**
Without grazer
Presence of grazer
Time (h)
Bio
film
Form
ing I
ndex
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Figure 2: Percentage of mono and multispecies planktonic cultures grazed by T. pyriformis after 24 h
and 96 h compared to the non-grazed cultures. The data points indicate the percentage reduction in
cell numbers (%) ± SEM obtained from five biological replicates.
In the biofilm fraction, cell numbers of X. retroflexus and S. rhizophila increased at 24 and 96 h in the
grazed relative to the non-grazed monocultures whereas the cell numbers of M. oxydans and P.
amylolyticus decreased with grazing compared to the non-grazed monocultures (Fig. 3). This
underscores the inability of M. oxydans and P. amylolyticus to form a biofilm in monoculture. In the
four-species culture, total cell numbers increased both at 24 h and 96 h compared to the non-grazed
biofilm (Fig. 3 and Fig. S2, supplementary information).
0
10
20
30
40
X.retroflexus S.rhizophila M.oxydans P.amylolyticus 4-species mix
24 h96 h
gra
zin
g f
old
(%
)
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Figure 3: Change in viable cell numbers from the biofilm fractions of mono and multispecies cultures
grazed with T. pyriformis after 24 h and 96 h obtained from plating. The data points indicate the
change in cell numbers of grazed biofilm fraction relative to the non-grazed biofilm fraction ± SEM
obtained from two biological replicates.
Growth of T. pyriformis on bacterial cultures
The growth of T. pyriformis cells on all bacterial isolates cultured as both mono and mixed planktonic
cultures was followed over time (Fig. 4). The change in cell numbers over time demonstrated that S.
rhizophila and P. amylolyticus were suitable prey for the protozoa (Fig. 4B and 4D) and that TSB
media can support the axenic growth of protozoa (Fig. 4F). Growth on M. oxydans was not
pronounced (Fig. 4C); while X. retroflexus monocultures had a negative impact on the growth of the
protozoa at 96 h (Fig. 4A). Our results thus indicate that T. pyriformis may prefer to graze on S.
rhizophila and P. amylolyticus. The numbers of protozoa grazing on the four-species mixed cultures
represent a smoother curve over time indicating that the protozoa can adapt to an available prey in
multispecies bacterial environments (Fig. 4E).
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Figure 4: T. pyriformis growth curves. The data points indicate the change in protozoan cell numbers
with respect to time (t = 0h) that were grown in co-culture with the bacterial isolates (A –D) as
monocultures and (E) as four-species mixed culture. (F) Change in protozoan cell numbers under
axenic conditions over time in TSB media. The data shown are mean ± SEM from three biological
replicates.
To visualize the change in protozoan numbers over time, micrographs showing T. pyriformis cells
raised on both mono and multispecies bacterial cultures are shown (Fig. S3, supplementary
information). The protozoan population raised on the four-species mixtures remained viable for up to
96 h. However, in monospecies cultures; it can be seen that the protozoan cell numbers increased
from 24 h and reached a maximum at 96 h when co-cultured with S. rhizophila and P. amylolyticus
whereas the protozoa population declined from 24 h to 96 h in co-culture with X. retroflexus and M.
oxydans.
Grazed bacterial prey within the food vacuoles of T. pyriformis
To visualize grazing on monocultures and mixed cultures, a 16S rRNA gene based FISH was
performed, similar to a previous study (Jezbera et al., 2005), after 24 hours of grazing and samples
were visualized by laser scanning confocal microscopy. It was confirmed that T. pyriformis can
consume the bacteria in all tested monospecies settings, however, at seemingly different rates as
indicated by the number of food vacuoles formed within the ciliates (Fig. 5).
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Figure 5: Grazing by protozoa on monospecies bacterial cultures. FISH based staining and confocal
imaging shows X. retroflexus (FL: A, BF: E and OL: I), S. rhizophila (FL: B, BF: F and OL: J), M.
oxydans (FL: C, BF: G and OL: K) and P. amylolyticus (FL: D, BF: H and OL: L) cells, cultured as
monospecies, localized within the food vacuoles (indicated by the arrows) of T. pyriformis cells after
24 h of grazing. ‘FL’ denotes fluorescence, ‘BF’ denotes bright-field and ‘OL’ denotes overlay
images respectively.
In co-cultures of T. pyriformis with X. retroflexus or S. rhizophila monocultures, the bacteria were
abundantly present within the food vacuoles of T. pyriformis cells (Fig. 5 A and B) showing that these
bacterial strains are readily consumed. P. amylolyticus cells were also found to be localized within the
food vacuoles of grazers but not as abundantly as compared to X. retroflexus and S. rhizophila (Fig.
5D). Most protozoan cells appeared to form cysts when co-cultured with M. oxydans (Fig. 5C), but
some bacterial cells were found to be internalized within T. pyriformis indicating that the protozoa
were able to consume M. oxydans cells. In the case of grazing on four-species mixed cultures (Fig.
6A-H), most food vacuoles were dominated by X. retroflexus indicating that at 24 h most protozoan
cells prefer to graze on X. retroflexus. This was in accordance with the fact that this bacterium
previously was shown to dominate the 24 h mixed biofilm population (Ren et al., 2014) and thus
could be readily available for the grazers.
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Figure 6: Grazing for 24 h by protozoa on the four-species mixed cultures. FISH based staining and
confocal imaging shows the distribution of the different bacterial species in and around the protozoan
cells. Applying the fluorescence filter channels, it is observed that X. retroflexus is abundantly present
within the food vacuoles (indicated by the arrows) of T. pyriformis (A). S. rhizophila (B) is detected
to a lesser extent whereas M. oxydans (C) and P. amylolyticus (D) cells are not visibly present in the
food vacuoles. Figures (E) and (F) depict the overlay and bright-field images, respectively. Figures
(G) and (H) were included to phase out the dominating fluorescence signals from X. retroflexus and
visualize the other bacterial members in the biofilm consortia around the ciliate.
Biofilm formation by X. retroflexus is vital to the overall biofilm development
From the above results, biofilm formation in the presence of T. pyriformis was further assessed to
better understand the dynamics. To this end, either the least preferred prey M. oxydans or the best
biofilm producer X. retroflexus were excluded three-species consortia (Fig. 7). Biofilm formation
(biofilm-fold Fd) was enhanced when X. retroflexus remained in the consortium together with S.
rhizophila and P. amylolyticus, indicating that the interaction between these three members is vital for
biofilm stability. However, in the absence of X. retroflexus and in the presence of M. oxydans, the
consortium was effectively grazed, although there seemed for this consortium to be a gradual
adaptation to predation (as evidenced by increased biofilm formation) over time.
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Figure 7: Biofilm formation in the presence of T. pyriformis in three-species bacterial consortia. X.
retroflexus is vital for biofilm development. Biofilm fold was calculated as the ratio of Abs590 [(three -
species biofilm cultured with grazer cells + SEM) – (three -species biofilm as control – SEM)] to
Abs590 (three -species biofilm cultured with grazer cells+ SEM).
Impact of grazing on the population dynamics of individual bacterial species in multispecies
biofilm and planktonic consortia
In order to determine the cell numbers of the individual species within the multispecies consortium,
16S rRNA gene based q-PCR quantification was applied according to a previously developed protocol
(Ren et al., 2014). The results showed that in the multispecies biofilm fraction, cell numbers of X.
retroflexus, S. rhizophila and P. amylolyticus increased in the presence of grazers compared to the
control biofilms that were not grazed. The ~2.5 fold increase in cell numbers of X. retroflexus and P.
amylolyticus and 1.7 fold increase in S. rhizophila cell numbers suggest that synergistic interactions
between these species were enhanced in the presence of grazing, resulting in increased cell numbers
in the biofilm. The cell numbers of M. oxydans in the biofilm remained unaffected either in the
presence or absence of grazers (Fig. 8A).
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Figure 8: Impact of grazing by T. pyriformis on the population dynamics of the individual bacterial
species in the multispecies consortia as assessed by qPCR. Cell numbers of the individual bacterial
members in (A) the multispecies biofilm fraction and (B) the multispecies planktonic fraction after 24
h of grazing.
In the planktonic fraction without grazing, a similar trend in cell numbers compared to the non-grazed
biofilm was seen with X. retroflexus, P. amylolyticus and S. rhizophila being the dominant species
(Fig. 8B). However, the planktonic cell numbers of these species decreased in the presence of T.
pyriformis indicating an effect of grazing on these planktonic fractions. In contrast, the cell numbers
of M. oxydans increased, which possibly can be a result of grazing preference of the ciliate in the
mixed communities and/or higher nutrient or space availability for M. oxydans cells as the other
members of the consortia were grazed upon.
Discussion
In the present study, the impact of grazing by the ciliate T. pyriformis on a previously described
synergistic mixed species biofilm model consortium (Ren et al., 2015) was assessed. These bacterial
strains were isolated from a single micro-habitat and studies have reported that long-term coexistence
within a habitat can stimulate synergistic biofilm development in complex communities (Madsen et
al., 2016). Our results showed that co-culturing T. pyriformis with single-species bacterial cultures
stimulated biofilm formation in X. retroflexus and S. rhizophila strains but not in M. oxydans and P.
amylolyticus (Fig. 1). S. rhizophila and P. amylolyticus were most sensitive to grazing (Fig 2). Ciliate
abundances reached a maximum in co-culture with these strains over time indicating extensive
feeding on these strains (Fig. 4B & 4D and Fig. S1; supplementary information). The monospecies
grazing experiments thus indicate differential bacterial behaviour in response to a predator and vice-
versa. Similar observations have been reported previously where protozoa regulate the social
behaviour of the bacteria (Rønn et al., 2002; Scherwass et al., 2016) or where bacteria regulate the
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protozoan population (Kaminskaya et al., 2007). The specificity of such responses has been reported
to vary depending on the selected bacteria and protozoa (Dopheide et al., 2011; Friman et al., 2013).
In the four-species consortia, biofilm formation was enhanced even when compared to the best
performing monoculture, suggesting a strong and significant biodiversity effect which was even
further enhanced in the presence of grazing (Fig. 1, Fig. S1, supplementary information). The total
cell numbers in mixed biofilm fraction under grazed conditions were increased compared to non-
grazed mixed biofilm (Fig 3) and at the same time, qPCR results showed that the bacterial numbers of
all strains except M. oxydans increased in comparison with the non-grazed biofilm. Even the grazing
sensitive species P. amylolyticus increased in cell numbers in the mixed biofilm during grazing. X.
retroflexus dominated the grazed biofilm followed by P. amylolyticus and S. rhizophila, respectively
(Fig. 8A). This suggests strong synergistic and complex interactions between these species under
grazing pressure, resulting in a shared protection against grazing. In contrast, total bacterial numbers
in the multispecies planktonic fraction under grazing were reduced for all species, except M. oxydans
(Fig. 8B). This can be explained by the lowest grazing preference for M. oxydans in monoculture.
Protozoan cell numbers in the mixed planktonic cultures (Fig. 4E) gradually decreased with time,
possibly reflecting a lower availability of the preferred individual prey or co-aggregation of the
bacterial consortia members into composite aggregates.
In the mixed-species consortia there was an increase by ~2.5 fold in total bacterial cell numbers (all
four species combined) in the grazed biofilm compared to the non-grazed biofilm, whereas in the
planktonic fractions grazing reduced total cell numbers by ~1.8 fold, emphasizing the protective
nature of the biofilm mode of life. Evidence that grazing pressure is positively correlated with the
formation of cell clusters has come from both monospecies laboratory biofilm (Matz et al., 2004,
2005) and from natural/semi-natural multispecies biofilm (Corno et al., 2015; Rychert & Neu R,
2010; Wey et al., 2008). Grazing induced biofilm formation could reflect either an active defence
mechanism (Friman & Buckling, 2014; Matz & Kjelleberg, 2005) or a passive mechanical process
where the movement of the protozoan cells drives the bacterial cells to the substratum (Wey et al.,
2012). Also, protozoan grazing on the planktonic bacterial population could release nutrients which
stimulate the biofilm-associated cells resulting in enhanced levels of biofilm formation (Böhme et al.,
2009; Petropoulos & Gilbride, 2005). Additionally, the total bacterial productivity is shown to be
influenced under grazing where bacterial aggregates display increased carbon transfer and uptake
(Corno et al., 2013, 2015). Discrepancies found in the literature with respect to the protective nature
of biofilms against grazing (Huws et al., 2005; Jackson & Jones, 1991; Weitere et al., 2005) could be
attributed to the type of protozoa used, their feeding mechanism and the growth conditions. Studies
have shown feeding traits of grazer to influence grazing resistance in bacterial biofilms (Seiler et al.,
2017) and surface associated bacteria can be even more consumed when exposed to a specialized
grazer (Rogerson & Laybourn-Parry, 1992). Therefore, more studies with different gazers are needed
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for a comprehensive picture on the effect of grazing by protozoan on bacterial biofilm. In this study,
we determined the grazing effect on a four species biofilm using a single pleagic grazer, the precise
mechanisms that confer grazing resistance to individual species remains unknown. However, the
biofilm formation was enhanced in a more diverse biofilm composed of four species, beyond the
expected biofilm forming capacities of all monocultures, especially under grazed conditions. Thus, in
a multispecies biofilm, the observed protection due to biofilm formation could be seen as a result of
synergistic interactions or complementarity within the mixed cultures.
In addition, X. retroflexus dominated the multispecies biofilm while M. oxydans was the least
preferred prey in monoculture. However, both these species have been shown to confer synergy and
shared protection (Hansen et al., 2017; Liu et al., 2017; Ren et al., 2015). Different three-species co-
cultures, set up to investigate the role of these two bacteria in the communal protection observed in
the multispecies biofilm, showed that biofilm formation was up by 3.5 folds in the three-species
biofilm composed of X. retroflexus, S. rhizophila and P. amylolyticus in the presence of grazers; but
that the synergy was hampered when X. retroflexus was substituted by M. oxydans (Fig. 7). From
these results, it can be deduced that the intricate interactions between X. retroflexus and the other two
members is vital for enhanced biofilm formation and communal grazing resistance. Grazing-sensitive
members (S. rhizophila and P. amylolyticus) are more susceptible to grazing in the absence of key
biofilm producers such as X. retroflexus. These results demonstrate that synergistic interactions within
the multispecies communities are further enhanced under grazing pressure, as also observed by
(Corno et al., 2015), and the multispecies biofilm architecture provided grazing sensitive members
with improved protection (Burmølle et al., 2016). This emergent property of multispecies biofilms
could serve as a public goods strategy, as previously reported for antimicrobials (Lee et al., 2014),
and can thus act as a major driver for synergistic cooperative behavior.
Our findings support previous findings (Madsen et al., 2016; Ren et al., 2015) that bacteria can
increase their fitness by engaging in the formation of multispecies biofilms. We showed that in
multispecies consortium under grazing pressure, cell numbers of free floating bacteria decrease while
biofilm cell numbers increase. Our findings thus suggest that synergy in biofilm formation could have
evolved from the selective pressures under stressful environmental conditions such as grazing.
Conflict of interests
The authors declare that they have no conflicts of interest.
Funding
This study was funded by grants from The Danish Council for Independent Research; ref no: DFF-
1335-00071, ref no: DFF-1323-00235 (SIMICOM) and BOF Special Research Fund, Belgium:
01SF1614.
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Acknowledgement
We thank Karin Vestberg and Anette Hørdum Løth for their technical assistance during the
experiments.
Authors and Contributions
PR, SS, MB and KH designed the study. PR performed the experiments. PR and WL analysed the
data. PR, WL, KS, KH, MB and SS revised the manuscript. KH, KB, MB and SJS provided the final
approval to publish.
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Supplementary Information
Figure S1: Quantification of (A) biofilm and (B) planktonic fractions of mono and mixed-species
cultures in the presence and absence of grazing Tetrahymena pyriformis (Tp) after 24 and 96 hours
obtained by plating. The data points indicate the colony forming unit (CFU) ± SEM obtained from
two biological replicates.
1.00E+00
1.00E+02
1.00E+04
1.00E+06
1.00E+08
1.00E+10
1.00E+12
1.00E+14
1.00E+16
1.00E+18
1.00E+20
CF
U/m
l
24h
96h
A B
1.00E+00
1.00E+02
1.00E+04
1.00E+06
1.00E+08
1.00E+10
1.00E+12
1.00E+14
1.00E+16
1.00E+18
1.00E+20
CF
U/m
l
24h
96h
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Figure S2: Micrographs depicting the number of T. pyriformis cells in co-culture with monospecies,
mixed-species bacterial cultures and in TSB media at 0.5 h, 6 h, 12 h, 24 h and 96 h. The protozoan-
bacterial suspensions were fixed using 1% (w/v) Lugol’s iodine solution.
Figure S3: Net Biodiversity effect calculated as the difference between the biofilm formation of the
mixed culture and the biofilm yield expected on the basis of the average of the monocultures at 12, 24
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and 96 hours. The biodiversity effect significantly increased in multispecies biofilm compared to the
monocultures and was even more pronounced in the presence of protozoa (ANOVA, *P < 0.05)
suggesting a strong synergistic or complementarity effect during biofilm formation. The data point
indicates the mean BFI ± SEM obtained from five biological replicates.
0
1
2
3
4
5
6
12 24 96
Net B
D e
ffect
Time (h)
Without grazer
Presence of grazer
**
*
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CV and List of publications
Prem Krishnan Raghupathi
Date of Birth : 4. July 1987
Nationality : India
Address : Sportstraat 119, 9000 Ghent, Belgium
Phone : 0032 465651622
Email : [email protected]
Education 2014-2018
Doctor of Veterinary Sciences
Doctor of Sciences
Copenhagen University and Ghent University
2010-2012
Master of Science
Food Science and Technology
Specialized in Food Safety
Copenhagen University, Denmark
2005-2009
Bachelor of Technology
Industrial Biotechnology
Anna University Chennai, India
Work Experience
2014- 2018
Ph.D. Candidate
Faculty of Veterinary Sciences, Ghent University, Belgium
Department of Biology, Copenhagen University, Denmark
2012-2014
Research Assistant
Section for Microbiology
Institute of Veterinary Disease Biology
Copenhagen University, Denmark
2009-2010
Marketing Executive
SR Lab Products and Diagnostics
Chennai, India
[CV AND PUBLICATIONS]
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Certifications
PRINCE 2 Project Management Certificate
RM Academy, Copenhagen, Denmark
Laboratory Handling of Animals
FELASA C, University of Copenhagen, Denmark
Scientific Publications:
1. Raghupathi PK, Dumolin C, Burmølle M, Sørensen SJ, Sabbe K, Houf, K . Identification
of Free Living Protozoa (FLP) and bacterial composition on toothbrushes. In preparation
2.Zupančič J, Raghupathi PK, Houf K, Burmølle M, Sørensen SJ, Gunde-Cimerman N. Synergistic Interactions in Microbial Biofilms Facilitate the Establishment of Opportunistic Pathogenic Fungi in Household Dishwashers. Front Microbiol. 2018 Jan 30;9:21. doi: 10.3389/fmicb.2018.00021. eCollection 2018. PubMed PMID: 29441043; PubMed Central PMCID: PMC5797641. 3.Raghupathi PK, Zupančič J, Brejnrod AD, Jacquiod S, Houf K, Burmølle M, Gunde-Cimerman N, Sørensen SJ. Microbiomes in Dishwashers: Analysis of the microbial diversity and putative opportunistic pathogens in dishwasher biofilm communities. Appl Environ Microbiol. 2018 Jan 12. pii: AEM.02755-17. doi: 10.1128/AEM.02755-17. [Epub ahead of print] PubMed PMID: 29330184; PubMed Central PMCID: PMC5812945. 4.Raghupathi PK, Liu W, Sabbe K, Houf K, Burmølle M, Sørensen SJ. Synergistic Interactions within a Multispecies Biofilm Enhance Individual Species Protection against Grazing by a Pelagic Protozoan. Front Microbiol. 2018 Jan 9;8:2649. doi: 10.3389/fmicb.2017.02649. eCollection 2017. PubMed PMID: 29375516; PubMed Central PMCID: PMC5767253. 5.Raghupathi PK, Herschend J, Røder HL, Sørensen SJ, Burmølle M. Genome Sequence of Psychrobacter cibarius Strain W1. Genome Announc. 2016 May 26;4(3). pii: e00078-16. doi: 10.1128/genomeA.00078-16. PubMed PMID: 27231353; PubMed Central PMCID: PMC4882934. 6. Herschend J, Raghupathi PK, Røder HL, Sørensen SJ, Burmølle M. Genome Sequence of Arthrobacter antarcticus Strain W2, Isolated from a Slaughterhouse. Genome Announc. 2016 Mar 31;4(2). pii: e00073-16. doi: 10.1128/genomeA.00073-16. PubMed PMID: 27034477; PubMed Central PMCID: PMC4816605. 7. Herschend J, Raghupathi PK, Røder HL, Sørensen SJ, Burmølle M. Genome Sequence of Kocuria palustris Strain W4. Genome Announc. 2016 Mar 31;4(2). pii: e00074-16. doi: 10.1128/genomeA.00074-16. PubMed PMID: 27034478; PubMed Central PMCID: PMC4816606. 8. Raghupathi PK, Herschend J, Røder HL, Sørensen SJ, Burmølle M. Genome Sequence of Kocuria varians G6 Isolated from a Slaughterhouse in Denmark. Genome Announc. 2016 Mar 31;4(2). pii: e00076-16. doi: 10.1128/genomeA.00076-16. PubMed PMID: 27034480; PubMed Central PMCID: PMC4816608.
[CV AND PUBLICATIONS]
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9.Raghupathi PK, Herschend J, Røder HL, Sørensen SJ, Burmølle M. Draft Genome Assembly of Two Pseudoclavibacter helvolus Strains, G8 and W3, Isolated from Slaughterhouse Environments. Genome Announc. 2016 Mar 31;4(2). pii: e00077-16. doi: 10.1128/genomeA.00077-16. PubMed PMID: 27034481; PubMed Central PMCID: PMC4816609. 10.Herschend J, Raghupathi PK, Røder HL, Sørensen SJ, Burmølle M. Draft Genome Sequences of Two Kocuria Isolates, K. salsicia G1 and K. rhizophila G2, Isolated from a Slaughterhouse in Denmark. Genome Announc. 2016 Mar 31;4(2). pii: e00075-16. doi: 10.1128/genomeA.00075-16. PubMed PMID: 27034479; PubMed Central PMCID: PMC4816607. 11.Røder HL, Raghupathi PK, Herschend J, Brejnrod A, Knøchel S, Sørensen SJ, Burmølle M. Interspecies interactions result in enhanced biofilm formation by co-cultures of bacteria isolated from a food processing environment. Food Microbiol. 2015 Oct;51:18-24. doi: 10.1016/j.fm.2015.04.008. Epub 2015 Apr 30. PubMed PMID: 26187823. 12.Alagesan K, Raghupathi PK, Sankarnarayanan S. Amylase inhibitors: Potential source of anti-diabetic drug discovery from medicinal plants. International Journal Of Pharmacy & Life Sciences. 2012 February 01; 3(2):1407-1412
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