Download Interactions of Foodborne Pathogens with Free‐living Protozoa

Interactions of Foodborne Pathogens with
Free-living Protozoa: Potential Consequences
for Food Safety
Mario J.M. Vaerewijck, Julie Baré, Ellen Lambrecht, Koen Sabbe, and Kurt Houf
Abstract: Free-living protozoa (FLP) are ubiquitous in natural ecosystems where they play an important role in the
reduction of bacterial biomass and the regeneration of nutrients. However, it has been shown that some species such
as Acanthamoeba castellanii, Acanthamoeba polyphaga, and Tetrahymena pyriformis can act as hosts of pathogenic bacteria.
There is a growing concern that FLP might contribute to the maintenance of bacterial pathogens in the environment.
In addition to survival and/or replication of bacterial pathogens in FLP, resistance to antimicrobial agents and increased
virulence of bacteria after passage through protozoa have been reported. This review presents an overview of FLP in
food-associated environments and on foods, and discusses bacterial interactions with FLP, with focus on the foodborne
pathogens Campylobacter jejuni, Salmonella spp., Escherichia coli O157:H7, and Listeria monocytogenes. The consequences of
these microbial interactions to food safety are evaluated.
Keywords: Acanthamoeba, bacteria–FLP interactions, food safety, foodborne pathogens, free-living protozoa (FLP),
Tetrahymena
Introduction
Food safety is a major subject of public concern. Food companies have to implement food safety systems such as GMP and
HACCP to meet the legal requirement for safe food and to gain
consumer confidence. Such systems are based on the control by reduction or, preferably, elimination of hazardous physical, chemical,
and biological agents before or during food processing. However,
the presence or even persistence of microbes, including foodborne
pathogens, in food processing industries cannot always completely
be excluded. Moreover, microorganisms often develop dense and
complex communities despite hostile treatments, such as cleaning
and sanitizing, and can survive environmental stresses such as low
temperatures and desiccation. Important microbial response mechanisms to survive these adverse conditions are induction of coldshock proteins, altered membrane compositions (Beales 2004), and
growth in biofilms, which act protectively against antimicrobial
agents (Bridier and others 2011).
During the last 4 decades, increasing attention has been paid
to the interactions of pathogenic bacteria with free-living protozoa (FLP) (amebae, flagellates, and ciliates) and to the potential
role of FLP in the maintenance of pathogens in the environment.
The first studies reported observations on engulfment and retention of mycobacteria (Jadin 1975; Krishna Prasad and Gupta
MS 20140351 Submitted 3/3/2014, Accepted 18/5/2014. Authors Vaerewijck,
Baré, Lambrecht, and Houf are with Dept. of Veterinary Public Health and Food
Safety, Ghent Univ., Belgium Author Sabbe is with Laboratory of Protistology and
Aquatic Ecology, Dept. of Biology, Ghent Univ., Belgium. Direct inquiries to author
Houf (E-mail: [email protected]).
1978) and Legionella pneumophila (Rowbotham 1980) in amebae.
In the meantime, intraprotozoan survival and replication is well
documented for L. pneumophila, the causal agent of Legionnaires’
disease (a severe pneumonia which can be lethal) and Pontiac
fever (a milder respiratory illness without pneumonia). L. pneumophila survives in biofilms but is also able to multiply in FLP
such as the amebae Acanthamoeba castellanii and Vermamoeba vermiformis (= formerly Hartmannella vermiformis) (Abu Kwaik and others 1998; Taylor and others 2009; Hilbi and others 2011; Richards
and others 2013). L. pneumophila and amebae are both found in
natural and man-made systems such as drinking water supplies,
whirlpools, cooling towers, and air conditioning devices. People
become sick by inhalation of tiny water droplets containing free L.
pneumophila bacteria or legionellae-laden vesicles expelled by amebae. Noteworthy, legionellae inside amebae are protected against
antimicrobial agents and are actively spread by motile protozoans.
Several foodborne and waterborne pathogens have been shown to
benefit from FLP in a similar way. Consequently, FLP are now not
merely considered as predators of bacteria but their potential role
in survival and protection of pathogenic bacteria is increasingly
recognized. FLP, which act as hosts of (foodborne) pathogens, are
considered as “reservoirs” for these bacteria (Brown and Barker
1999). L. pneumophila developed an escape mechanism to avoid
digestion by amebae and applies several steps of this pathway in
human alveolar macrophages, which suggests that amebae may
also serve as “biological gyms” (Harb and others 2000), “evolutionary cribs” (Greub and Raoult 2004), or “training grounds”
(Molmeret and others 2005). Finally, in some cases FLP function as “Trojan horses” where the protozoan “horse” may bring
924 Comprehensive Reviews in Food Science and Food Safety r Vol. 13, 2014
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doi: 10.1111/1541-4337.12100
Interactions of foodborne pathogens with FLP . . .
internalized bacteria within the human “Troy,” enabling bacteria to pass the 1st line of the human defense system (Barker and
Brown 1994; Greub and Raoult 2004). Because FLP can facilitate the survival, growth, and dispersal of bacterial pathogens,
host–microbe interactions, which benefit bacterial pathogens, are
increasingly considered as a potential health risk (Thomas and
Ashbolt 2011).
The aim of this review is to provide an overview of the current knowledge on interactions of foodborne pathogens with
FLP. It starts with a brief description of FLP, followed by an
overview of FLP presence in food-related environments and
on food. The possible modes of interactions between bacteria
and FLP are summarized, and an overview of known interactions of foodborne pathogens with FLP is presented. The intraprotozoan survival of 4 major bacterial foodborne pathogens
(Campylobacter jejuni, Salmonella spp., Escherichia coli O157:H7,
and Listeria monocytogenes) is discussed in detail. Where appropriate, reference is made to other, well-characterized bacteria–FLP
interactions (Legionella–Acanthamoeba, Legionella–Tetrahymena, and
Mycobacterium–Acanthamoeba) to highlight important cellular and
ecological features. Finally, the consequences of these microbial
interactions to food safety are evaluated.
Free-Living Protozoa
Definition and morphology
While there is no unambiguous definition of “protozoa”, a
general agreement is to consider protozoa as heterotrophic protists. Protists are defined by Adl and others (2005) as “eukaryotes
with a unicellular level of organization, without cell differentiation
into tissues”. FLP comprise protists which do not have an obligate
parasitic life cycle, although some species such as A. castellanii,
Balamuthia mandrillaris, and Naegleria fowleri occasionally cause
infections in humans (Marciano-Cabral and Cabral 2003; Khan
2006; Visvesvara and others 2007).
FLP are single-celled microorganisms. Two life stages are usually
distinguished: a trophozoite (vegetative cell) and a cyst (also: resting or dormant cyst). However, not all species produce cysts. The
trophozoite is the life stage in which the cell feeds and multiplies.
Sizes of trophozoites range from a few μm to a few mm. The
variety in cell shape is virtually inexhaustible and forms the basis
of morphological identification. The body surface consists of a cell
membrane but can be covered by scales or be surrounded by a test
or lorica (Sleigh 1989). FLP are mostly solitary but some species
form colonies. Most FLP are motile (for example, swimming,
crawling, or gliding), others float passively in the water column.
Several species are attached to surfaces (for example, stones, submerged plants, detritus, and aggregates) by means of a stalk or trailing flagellum. Bacterivorous FLP feed on bacteria. Other modes
of nutrition are feeding on protozoa, fungi, algae, or small invertebrates (for example, rotifers), growth on detritus, or absorption of
organic material or molecules. Omnivorous FLP use more than 1
nutritional strategy. Some FLP are mixotrophic and combine heterotrophic feeding with photosynthesis. Many FLP have at least
1 contractile vacuole, which is an osmoregulator that pumps, at
regular frequency, excess water and dissolved waste products out of
the cell. FLP multiply mostly by binary cell division although several species have sexual cycles. Cyst formation (encystment) leads
to a fundamental change in trophozoite morphology and physiology, and it can be induced by overcrowding, nutrient depletion,
accumulation of certain metabolites, or unfavorable environmental
conditions including desiccation, changes in pH, osmolarity, oxygen level, or temperature (Corliss and Esser 1974; Corliss 2001;
C 2014 Institute of Food Technologists®
Gutiérrez and others 2001; Hausmann and others 2003). Cysts
are nonmotile and generally smaller than trophozoites and usually
have a thick, often double- or multilayered wall (Corliss 2001).
The cyst wall is composed of proteins, glycoproteins, and carbohydrates such as cellulose or chitin (Corliss 2001). The cyst shape
may vary (for example, spherical, ovoid, pyriform) and several
cysts have an outer surface ornamentation (Corliss 2001), which
can aid for dispersal. Cysts can survive harsh conditions and, for
example, cysts of Acanthamoeba spp. resist gamma and UV irradiation (Aksozek and others 2002), heat (Storey and others 2004)
and disinfectants (Kilvington and Price 1990; Storey and others
2004; Coulon and others 2010), and they can remain viable for
many years (Mazur and others 1995; Sriram and others 2008).
Excystment is the transformation from cyst to trophozoite and
is triggered by favorable conditions or chemicals (Hausmann and
others 2003).
Based on morphology and locomotion, FLP are divided into
the amebae, the flagellates, and the ciliates (Figure 1). This division
is used very often for convenience, but it is strongly discouraged
by protistologists because it does not take into account the phylogenetic relationships between these organisms (Adl and others
2005). However, for the sake of simplicity and to present a general
overview, the FLP-morphogroups amebae, flagellates, and ciliates will be used throughout this review. In the rest of the text,
only organisms nonparasitic to humans (“free-living”) are covered. Indeed, each morphogroup contains a few human parasites
such as the flagellate parasites Giardia duodenalis (causing giardiasis) and Trypanosoma brucei (causative agent of sleeping sickness),
the ameba Entamoeba histolytica (amebic dysentery), and the ciliate
Balantidium coli (balantidiosis). Obviously, other obligate parasitic
protozoa, such as Plasmodium spp. (causative agents of malaria) and
foodborne pathogens such as Cryptosporidium parvum, Cyclospora
cayetanensis, Sarcocystis hominis, and Toxoplasma gondii, all organisms
classified in the past in the Sporozoa, will also not be discussed in
this review.
Amebae are characterized by the possession of pseudopodia
(Greek: false feet), which are transient cytoplasmic extensions of
the cell. Pseudopodia are used for locomotion and feeding. However, small- and medium-sized amebae often move as a whole
without forming pseudopodia (Smirnov and Brown 2004). Amebae are roughly divided into naked amebae, testate amebae, and
actinopods. Naked amebae are surrounded by a plasmamembrane.
The typical ameboid movement on surfaces results in temporal
changes of the cell shape. Some species have a temporally flagellated life stage (for example, N. fowleri). Testate amebae have
a plasmamembrane covered with a test (shell). The test is composed of organic material, inorganic salts, or agglutinated particles.
Actinopods are star-like amebae with stiffened pseudopodia radiating out of a spherical cell. Some representatives of naked amebae
are Acanthamoeba, Amoeba, Hartmannella, and Vannella. Examples of
testate amebae are Arcella and Difflugina. The actinopods include
the Heliozoa (for example, Actinophrys) and the marine Radiolaria
which have a mineral skeleton.
Flagellates possess at least 1 flagellum. If there is more than 1
flagellum, they may be unequal in length and can be differently
oriented (Patterson 1998). A flagellum is used for locomotion,
feeding, or attachment to a surface, and it emerges near or at the
anterior of the cell (Sigee 2005). The structure of a eukaryotic
flagellum differs fundamentally from a prokaryotic flagellum. A
cross-section of the eukaryotic flagellum shows the typical arrangement of 9 outer doublet microtubules that encircle 2 singlet
microtubules (9 × 2 + 2 pattern). Flagellates swim or glide, and the
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Interactions of foodborne pathogens with FLP . . .
Figure 1–Examples of morphotypes of free-living protozoa. (A) Acanthamoeba (an ameba), (B) Chilomonas (a flagellate), (C) Tetrahymena (a ciliate).
Organisms not drawn to scale. Pictures taken from http://pinkava.asu.edu/starcentral/microscope/. Drawings made by Stuart Hedley and David
Patterson, licensed to MBL (micro*scope). Reprinted with permission.
flagellum can beat with a sine wave or be held stiffly, sometimes
only moving with the tip (Patterson 1998). Other flagellates are
sessile (for example, choanoflagellates) or are temporarily attached
to the surface (for example, Bodo saltans). Many species are
very small, and cells with a size between 2 and 15 μm are usually
described as nanoflagellates, and those of less than 2 μm as
picoflagellates (Boenigk and Arndt 2002). Some flagellate groups
contain photosynthetic members (for example, Euglena). Therefore, in older literature sources, the division into zooflagellates
and phytoflagellates was made. Bodo, Cercomonas, Chilomonas,
Goniomonas, Heteromita, Notosolenus, Petalomonas, and Spumella
are all examples of flagellate genera commonly found in the
environment.
Ciliates have cilia that are structurally identical to flagella but
are shorter, occur in higher numbers over the cell body, and differ in beat pattern (Sigee 2005). Cilia are used for locomotion,
feeding, attachment, or sensing (Lynn 2008). Cilia are often arranged in distinct rows and they beat in a coordinated fashion.
The cell size of ciliates is mostly much larger compared with amebae and flagellates (most ciliates are between 20 and 200 μm in
length) (Sigee 2005). Ciliates are motile (crawling, swimming) or
are attached to a substrate. Ciliates have 2 different nuclei: a small,
diploid micronucleus necessary for reproduction and a large, polyploidy, transcriptionally active macronucleus, which controls the
organism’s phenotype (Lynn 2008). Ciliates feed by filter-feeding
or predation. Filter feeders concentrate prey through the action of
cilia, which are sometimes grouped to form special membrane-like
structures. Many ciliates have an oral cavity with a cell “mouth”
(cytostome) where food propelled through the action of cilia is
concentrated into a food vacuole. Undigested material is excreted
at the cytoproct which usually is located at the posterior side of
the cell. Predatory ciliates catch their food (for example, other ciliates) through specialized organelles such as extrusomes. A special
category is the Suctoria, a group of sessile ciliates that lack cilia
in their adult stage and capture prey with cytoplasmatic tentacles. The ciliates are a morphologically very heterogeneous group.
The textbook example of the ciliates is the slipper animal Paramecium, but others such as Colpoda, Colpidium, Cyclidium, Glaucoma,
Tetrahymena, and sessile forms such as Epistylis and Vorticella are
easily isolated from nature.
Classification and diversity of FLP
The artificial division of FLP in amebae, flagellates, and ciliates has no phylogenetic value but is often used in teaching. The
classification of microscopic eukaryotic organisms has undergone
many revisions and it was already clear from the beginning of
the 19th century that single-celled eukaryotes were a difficult
part in eukaryotic systematics (Scamardella 1999). The eukaryotes
were divided according to Whittaker into the Kingdoms Animalia, Plantae, Fungi, and Protista (Whittaker 1969). The classification of protozoa (“animal-like” protists), photosynthetic organisms other than plants and cyanobacteria (“plant-like” protists or
(micro)algae), and slime molds (“fungi-like” protists) in the Kingdom Protista served for many years. Several fundamental revisions
of Whittaker’s classification scheme have followed (for example,
Woese and others 1990; Cavalier-Smith 2004) and the idea of
protists as a separate group was abandoned. Currently, eukaryotic
organisms are classified into 5 supergroups based on ultrastructural, biochemical, and molecular phylogenetic data (Adl and others 2012): (i) the Amoebozoa (including naked and many testate
amebae, slime molds), (ii) the Opisthokonta (including Metazoa
[sponges, animals], fungi, yeasts, choanoflagellates, Mesomycetozoea, Nuclariids), (iii) the Excavata (including several flagellate
genera and the ameba group Heterolobosea), (iv) Sar, a cluster
of Stramenopiles (including the chrysophytes, diatoms, brown algae, some fungi-like organisms), Alveolata (including ciliates (Ciliophora), dinoflagellates, Apicomplexa), and Rhizaria (including
Cercozoa, Foraminifera, Radiolaria, some testate amebae), and
(v) the Archaeplastida (including Glaucophyta, plants, and red and
green algae).
The global number of protist species is still hotly debated. Recent estimates range from 16600 to 300000 species (Fenchel and
Finlay 2006; Foissner 2008), but it is not unlikely that even these
numbers are underestimates. While until recently most protist
species were considered to be cosmopolitan, there is increasing
evidence that microorganisms (including protists) display biogeographic patterns (Hanson and others 2012; Bates and others 2013).
For example, the ciliate Tetrahymena thermophila, a species that is
a commonly used as a eukaryotic model organism in genetic and
molecular biology studies and bacteria–FLP interaction studies
(see later) has recently been shown to have a distribution which
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Interactions of foodborne pathogens with FLP . . .
in grazing resistance rather than for virulence as such (Adiba and
others 2010).
Importantly, as a result of protozoan grazing, dissolved organic
matter and inorganic nutrients such as nitrogen and phosphorus
are released into the environment. These nutrients enter into the
microbial biomass and as such become available for higher trophic
levels. For example, as a result of grazing activities in soil, the
release of nutrients from consumed bacterial biomass together
with a shift to rhizobacterial communities (which are responsible
for better nitrogen fixation) both have a beneficial effect on plant
growth (Bonkowski 2004).
Finally, FLP serve as food for other organisms such as zooplankton, other small invertebrates, and fish larvae, and as such play an
Ecological importance of FLP
Bacterivorous feeding by FLP is, together with viral-mediated important role in aquatic and terrestrial food webs.
lysis, one of the main regulatory processes of bacterial biomass
(Sherr and Sherr 2002; Pernthaler 2005). Especially heterotrophic Occurrence in natural habitats and anthropogenic
nanoflagellates such as bicosoecids and Chrysophyceae (for ex- environments
ample, Spumella spp.) and small ciliates are very important bacFLP are ubiquitous in aquatic and terrestrial ecosystems. Imterial consumers in marine and freshwater ecosystems (Boenigk portant habitats include coastal waters, lakes, oceans, ponds, rivers,
and Arndt 2002). Numbers of heterotrophic nanoflagellates vary swamps, sediments, and soils. FLP are distributed all over the water
between hundreds to several tens of thousands cells/mL, de- column and can be found in waters with different nutrient and
pending on the trophic system and season (Boenigk and Arndt salt contents. In soils, FLP live in the water film around soil parti2002). Amebae are important grazers of biofilms and are prob- cles. Although FLP mainly inhabit water or moist environments,
ably, together with flagellates, the most important group of soil some species are able to survive in soils with very low moisture
protozoa (Ekelund and Rønn 1994; Foissner 1999). Protozoan content such as deserts and the dry valleys on Antarctica. They
grazing is often prey-selective, and medium-sized bacterial cells become active, often for a very limited period, under favorable
(0.4 to 1.6 μm) are preferably consumed (Pernthaler and others conditions such as a short rain period. FLP are found in extreme
1996; Hahn and Höfle 2001). Ingestion rates vary from several environments such as deep sea vents, geothermal hot springs, and
hundreds to more than 1000 bacteria/protozoan cell/h (Parry acidic and hypersaline lakes. In addition to water and soil, air
2004). However, food selection is not only determined by prey can contain viable FLP and cysts (Schlichting 1964; Kingston and
size. Physiological state of the prey (with preference for bacteria Warhurst 1969; Rivera and others 1992; Rogerson and Detwiler
in stationary phase compared to exponentially growing bacte- 1999). The above-mentioned examples of habitat types illustrate
ria) (Ayo and others 2009) and chemicals secreted by the prey that FLP can tolerate a broad range of physical (for example, tem(Hamels and others 2004) have also been shown to influence prey perature and humidity), chemical (for example, pH and oxygen
selectivity. Protozoan characteristics such as feeding forms (for content), and nutritional parameters. The tolerance range can be
example, phagocytosis and filter feeders), cell size (for example, large or small for certain parameters, and tolerance limits of several
small flagellates versus large ciliates), and behavior are important species can be broadened by gradual adaptation to new conditions
as well. Remarkably, recent feeding history of the protozoan had (Hausmann and others 2003).
an influence on the response toward prey types (Ayo and others
Besides abiotic environments, FLP are present on or in various
2009).
organisms. Insects can carry FLP on their body (Maguire 1963;
Bacteria have developed several antipredator strategies includ- Revill and others 1967a, b) and some FLP inhabit the hindgut
ing cell size reduction, modified cell morphology (for example, of insects such as termites (Ohkuma 2008). Ciliates inside the
filamentous growth), modification of cell wall characteristics (for rumen of ruminants such as cattle and sheep play an important
example, surface potential), high-speed motility, and production of role in the digestion process and contribute substantially to the
exopolymers or toxins (Hahn and Höfle 2001; Matz and Kjelle- energy and nutrient balance (Veira 1986; Williams 1986). Roots
berg 2005; Pernthaler 2005; Jousset 2012). Biofilm and micro- of plants are colonized with FLP (Bonkowski 2004), as well as the
colony formation are other well-known defense mechanisms to above-ground parts such as leaf surfaces (Ruinen 1961; Bamforth
resist protozoan grazing, and quorum-sensing is an important 1971) and bark of trees (Bartošová and Tirjaková 2008). Other
communication tool to inform neighboring bacterial cells of up- examples are mosses (Anderson 2006; Glime 2007), pitchers of
coming threats (Matz and Kjelleberg 2005). Some bacteria resist pitcher plants (Hegner 1926; Rojo-Herguedas and Olmo 1999),
digestion through escape from or modification of the phagosome and tank water of bromeliads (Foissner and others 2003).
(for example, L. pneumophila), or they survive because of the inGiven their ubiquity, it is not surprising that FLP have also
efficiency of the protozoan digestive system (Thurman and others been detected in various man-made constructions such as cars
2010).
(Simmons and others 1999), cooling towers (Yamamoto and othThe combination of selective grazing, protozoan community ers 1992; Berk and others 2006; Behets and others 2007), dental
composition, and bacterial antipredator mechanisms influences the unit waterlines (Barbeau and Buhler 2001; Singh and Coogan
morphological structure and genotypic and taxonomic composi- 2006), drinking water supplies (Thomas and Ashbolt 2011), eyetions of bacterial communities (Hahn and Höfle 2001; Jürgens wash stations (Paszko-Kolva and others 1991; Bowman and others
and Matz 2002). Moreover, it is hypothesized that grazing not 1996), homes (Marciano-Cabral and others 2003; Trzyna and othonly selects for bacteria with traits which infer resistance to graz- ers 2010; Stockman and others 2011), hospitals (Rohr and others
ing, but also for traits responsible for virulence (Adiba and others 1998), and hospital water networks (Patterson and others 1997;
2010). It was suggested that virulence is maintained for their role Thomas and others 2006), moisture-damaged buildings (Yli-Pirilä
is restricted to the eastern United States (Zufall and others 2013).
The different opinions on global protist species richness and biogeography can be largely explained by the absence of a generally
accepted species concept, the occurrence of cryptic species (defined by Caron 2009 as “morphospecies of protists that contain
strains possessing different physiological abilities or mating incompatibilities”), and inadequate methodology such as undersampling
of habitats, difficulties to exhaustively characterize communities
(underreporting) and in determining dispersal rates (Schlegel and
Meisterfeld 2003; Foissner 2008; Caron 2009; Boenigk and others
2012).
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Interactions of foodborne pathogens with FLP . . .
and others 2004), spacecraft (Ott and others 2004), swimming chloride and sodium hypochlorite, both active compounds of frepools (Rivera and others 1993), and wastewater treatment plants quently used food sanitizers, were effective at concentrations commonly applied in the food industry against trophozoites of Acan(Madoni 1994; Pauli and others 2001).
thamoeba polyphaga, Tetrahymena pyriformis, and a Tetrahymena sp.
Occurrence and Diversity of FLP in Food-Related isolated from a meat-cutting plant (Vaerewijck and others 2012).
The presence of FLP in the 5 food-processing plants can be exEnvironments
The microbiology of food-related environments (places where plained by reasons such as inadequate or insufficient sanitation,
food products are produced, processed, or stored) is examined in inactivation of sanitizers by organic material, or due to uncharacthe context of (i) hygiene control, (ii) tracing the source of spoilage terized cell response mechanisms. Furthermore, various FLP form
and pathogenic microorganisms, and (iii) characterizing persistent cysts which are more resistant to disinfectants than trophozoites
strains. Hygiene monitoring generally includes analysis of micro- (Khunkitti and others 1996; Coulon and others 2010).
At home, fresh food is mostly stored in refrigerators. Results of
biological parameters such as total aerobic bacteria and hygiene
indicator organisms such as E. coli. Specific microorganisms are studies on general hygiene, occurrence of bacterial pathogens, and
determined in cases of food safety or food spoilage problems which temperature status of domestic refrigerators are regularly reported,
often requires intensive sampling of the production facility. Simi- but only 1 study has focused on the occurrence and diversity
larly, sampling sites or equipment can preemptively be monitored of FLP (Vaerewijck and others 2010). Surfaces of vegetable trays
on food pathogens. For example, L. monocytogenes will receive spe- were mostly FLP-positive, followed by discharge gutters, whereas
cial attention in plants where dairy products, meat, ready meals, interior walls were rarely FLP-positive. Vegetable trays were most
and smoked fish are processed. However, FLP are never included likely contaminated by FLP through transfer of FLP from vegetain microbiological monitoring because these microorganisms are bles (see later) to plastic surfaces. Amebae and flagellates were the
considered harmless. Consequently, data on FLP of food-related most encountered FLP while ciliates were mainly retrieved from
vegetable trays. Discharge gutters were occasionally contaminated
environments are still scarce.
Protozoan communities at farms are hardly examined, and stud- with a persistent protozoan population of amebae and flagellates.
ies are limited to broiler houses. FLP were detected in poultry Importantly, the FLP-positive status of refrigerator surfaces was
drinking water systems (Snelling and others 2005, 2006; Baré and correlated with a high aerobic plate count.
Collectively, the above-mentioned studies show that FLP are
others 2009, 2011) and to a lesser degree in litter, animal feed, and
dry areas (Baré and others 2009). Poultry water supplies showed part of the in-house microbiota of food-related environments, and
a significantly higher FLP diversity than dry environments, with that FLP diversity is high (Table 1). Surprisingly, the overall FLP
flagellates and ciliates characteristic for water and dry environ- diversity of broiler houses, meat-cutting plants, and domestic rements, respectively, and amebae appearing more or less indifferent frigerators was quite similar with representatives of the Amoebozoa
to habitat type (Baré and others 2009). Protozoan communities (Vannellida, Tubulinea, Thecamoebida), Excavata (Euglenozoa,
in broiler houses were highly diverse, with a total of 91 morpho- Kinetoplastea, Heterolobosea), Stramenopiles (Chrysophyceae),
taxa (identification of FLP based on morphology) and 22 unique Alveolata (Ciliophora), and Rhizaria (Cercomonadida) as most
phylotypes (identification of FLP based on [partial] gene sequence frequently detected taxa. Many of these FLP are common in soil
data) observed (Baré and others 2009). Of all protozoan morpho- and water habitats (Sleigh 1989; Hausmann and others 2003) and
taxa distinguished, about half were amebae, about one-quarter live in benthos or environments containing a high organic load
were ciliates, and the remainder flagellates. Interestingly, the pro- (Patterson 1998).
tozoan community structure of the broiler houses showed almost
no change during a 6-mo follow-up and remained highly habitat- Occurrence and Diversity of FLP on Food Products and
and farm-specific (Baré and others 2011). Snelling and others in Drinking Water
(2006), however, reported a much lower FLP diversity (9 phyloTo date, knowledge of quality, food safety, and microbial
types) in broiler houses, which is mainly explained by differences
in identification methodology and the number and types of sam- contamination of food products mainly concerns spoiling and
pathogenic bacteria, yeasts and molds, parasitic protozoa and
ples analyzed.
The occurrence and diversity of FLP in food-processing envi- worms, and viruses. The FLP diversity of food products, howronments is largely unexplored. To the best of our knowledge, ever, has rarely been studied.
Vegetables become contaminated with FLP through contact
there is only 1 published study on protozoan communities in food
industrial environments. Vaerewijck and others (2008) showed that with soil, irrigation water, air, rain, insects, and during indusactive communities of amebae, flagellates, and ciliates were present trial produce-washing. Amebae were recovered from mushrooms
in meat-cutting plants. FLP were detected mainly in floor drains, in and vegetables such as carrots, cauliflower, lettuce, radishes, scalstanding water on the floor, on soiled bars of cutting tables, on plas- lions, spinach, and tomatoes (Ciurea-Van Saanen 1981; Napolitic pallets, and in out-of-use knife sanitizers, but they were also de- tano 1982; Napolitano and Colletti-Eggolt 1984; Rude and othtected on surfaces which come into direct contact with meat such ers 1984; Sharma and others 2004; Gourabathini and others 2008;
as conveyer belts, working surfaces of cutting tables, and needles of Vaerewijck and others 2011) (Table 2). Flagellates were found on
a meat tenderizer device. Sampling sites rich in organic material spinach, Romaine lettuce, and butterhead lettuce (Gourabathini
(for example, floor drains, meat residues on the floor, soiled sur- and others 2008; Vaerewijck and others 2011). Ciliates were refaces) or with high moisture content (for example, floor drains and covered from mushroom surfaces (Napolitano 1982) and leafy
standing water on the floor) were more likely to be FLP-positive. vegetables such as spinach and butterhead lettuce (Gourabathini
Based on microscopy, 61 morphotaxa were found and sequencing and others 2008; Vaerewijck and others 2011). The number of
of 18S rRNA gene fragments revealed 49 unique phylotypes. The FLP on leafy vegetables was high, with flagellates being more
occurrence of FLP in meat-cutting plants suggests that FLP resisted abundant (up to more than 105 cells per mL drained water or
the daily cleaning and sanitation process. However, benzalkonium per gram leaf) than amebae and ciliates (around several hundreds
928 Comprehensive Reviews in Food Science and Food Safety r Vol. 13, 2014
C 2014 Institute of Food Technologists®
Interactions of foodborne pathogens with FLP . . .
Table 1–Overview of protist taxa detected in 3 food-related environments. Compilation of identifications obtained by microscopy and sequencing of
18S rRNA gene fragments. Data obtained from Baré and others (2009, 2010), Snelling and others (2005, 2006), and Vaerewijck and others (2008,
2010, 2011). Classification according to Adl and others (2012).
Food-related environment
Supergroup
Amoebozoa
1st rank
Tubulinea
Discosea
2nd rank
Euamoebida
Leptomyxida
Arcellinida
Flabellinia
Himatismenida
Longamoebia
Mastigamoebaea
Opisthokonta
Archamoebae
Protosteliida
Holozoa
Sar
Nucletmycea
Stramenopiles
Nuclearia
Bicosoecida
Peronosporomycetes
Chrysophyceae
Protalveolata
Ciliophora
Alveolata
Choanomonada
Rhizaria
Dinoflagellata
Cercozoa
Archaeplastida
Chloroplastida
Chlorophyta
Excavata
Discoba
Discicristata
Incertae sedis
Eukaryotaa
Centrohelida
Heterophryidae
3rd rank
Phryganellina
Echinamoebida
Vannellida
Dermamoebida
Thecamoebida
Centramoebida
Craspedida
Acanthoecida
Colpodellida
Cercomonadidae
Imbricatea
Chlorophyceae
Heterolobosea
Euglenozoa
Examples of
detected taxa
Broiler
houses
Meatcutting plants
Domestic
refrigerators
+
+
+
+
+
+
Hartmannella, Saccamoeba
Leptomyxa
Cryptodifflugia sp.
Echinamoeba
Vannella spp., Ripella
Platyamoeba placida
Cochliopodium sp.
Mayorella sp.
Thecamoeba sp.
Acanthamoeba spp.
Mastigamoeba/Mastigella
Protostelium mycophaga
Monosiga
Diaphanoeca grandis,
Stephanoeca diplocostata
Nuclearia
Adriamonas
Saprolegnia parasitica
Spumella spp.
Colpodella spp.
Colpoda, Cyclidium,
Tetrahymena, Glaucoma,
Vorticella
Glenodinium
Cercomonas, Heteromita,
Spongomonas minima
Chloromonas sp., Polytoma
uvella
Vahlkampfia
Petalomonas, Peranema,
Notosolenus, Bodo
Oxnerella sp.
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
a Incertae sedis Eukaryota is not considered as a supergroup. Incertae sedis Eukaryota contains organisms with uncertain affiliation within the protists.
Table 2–Identified FLP species detected on vegetables and mushrooms. Identification according to taxonomic convention of that time.
Vegetables
Morphogroup
Lettuce, scallions, radishes
Amebae
Mushrooms (Agaricus bisporus)
Amebae
Oak leaf lettuce, Boston lettuce
Ciliates
Amebae
A. rhysodes, A. palestinensis, Naegleria
gruberi
Acanthamoeba spp., Hartmannella spp.,
Vannella spp.
Colpoda sp., holotrichs, hypotrichs
Acanthamoeba spp., Hartmannella spp.
Amebae
A. polyphaga, A. rhysodes, A. castellanii
Amebae
Amebae
A. culbertsoni, A. rhysodes, H. vermiformis
A. palestinensis
Ciliate
Ciliate
Amebae
Glaucoma sp.
C. steinii
Cochliopodium sp., Mayorella vespertiloides,
Ripella platyopodia, Ripella platyopodia,
Vannella simplex, H. vermiformis,
Saccamoeba sp., Vahlkampfia sp.
Goniomonas truncata, Bodo saltans, Bodo sp.,
Cercomonas spp., Notosolenus sp.,
Peranema trichophorum, Petalomonas sp.,
Rhynchomonas nasuta, Allantion
tachyploon, Spumella spp.
Aspidisca lynceus, Cinetochilum
margaritaceum, Chilodonella uncinata, C.
steinii, Colpoda sp., Cyclidium glaucoma,
Glaucoma scintillans, Glaucoma sp.,
Paramecium putrinium, Platyophrya sp.,
Tachysoma pellionellum, Trachelophyllum
sp., Vorticella convallaria complex,
Vorticella sp.
Bodo saltans, Cercomonas spp., Spumella
cylindrica, Spumella-like flagellates
C. steinii
Carrots, radishes, tomatoes, mushrooms,
cauliflowers, spinach
Carrots
Romaine lettuce
Spinach
Butterhead lettuce
Flagellates
Ciliates
Ready-to-eat lettuce (butterhead lettuce)
Flagellates
Ciliate
C 2014 Institute of Food Technologists®
Identified species
Reference
Ciurea-Van Saanen
(1981)
Napolitano (1982)
Napolitano and
Colletti-Eggolt (1984)
Rude and others (1984)
Sharma and others (2004)
Gourabathini and others
(2008)
Vaerewijck and others
(2011)
Vaerewijck and others
(2011)
Vol. 13, 2014 r Comprehensive Reviews in Food Science and Food Safety 929
Interactions of foodborne pathogens with FLP . . .
of cells per mL drained water or per gram leaf) (Gourabathini
and others 2008; Vaerewijck and others 2011). Washing or rinsing lettuce leaves followed by spin-drying in a household salad
spinner reduced the protozoan number only by a maximum 1 log
unit (Vaerewijck and others 2011). FLP were also recovered from
ready-to-eat lettuce, which suggests that they survived mild treatments such as industrial washing (Vaerewijck and others 2011).
No information was found on FLP diversity on fruit surfaces, although it can be assumed that they are likely to be present due
to contamination by air, insects, rain, or during industrial processing. Trypanosomatid flagellates were found inside edible fruits
such as plums, oranges, mandarins, grapes, pineapples, and apples
(Camargo 1999; Catarino and others 2001). Trypanosomatids such
as Phytomonas serpens and Phytomonas mcgheei are widespread in
fruit and already 33 species of fruit have been reported to be
harboring flagellates (Camargo 1999). Trypanosomatids have also
been isolated from tomatoes, beans, and maize seeds. The role
of trypanosomatid flagellates in fruit is unknown. Transmission of
flagellates occurs by insects (phytophagous hemipterans), which
feed on fruit juice after penetration of their stylet bundle in plant
tissues. The penetration causes only local mechanical damage.
No studies have been published on FLP-diversity of meat and
meat products yet. FLP were recovered from surfaces which came
into contact with meat such as conveyer belts, needles of a meat
tenderizer device, and working tables (Vaerewijck and others
2008). It is not unlikely that cross-contamination of FLP between
both surfaces takes place. However, studies are necessary to determine the transfer rate and to determine if meat juice or meat
surfaces are suitable environments for FLP survival and multiplication. Similarly, to the best of our knowledge, FLP communities on
edible fish and shellfish have not been examined, although harmless FLP can be found on shells or gills of crustaceans (so-called
epibiotic protozoa) (Fernandez-Leborans 2010) and on the skin of
fish. Most studies have focused on fish and shellfish with disease
caused by infectious species (reviewed in Scholz 1999; Dyková
and Lom 2004; Morado 2011).
Contrary to food products, more information has been published on the occurrence of FLP in drinking water supply systems
and tap water at home. The occurrence and diversity of freeliving amebae in drinking water production plants and distribution systems was recently reviewed by Loret and Greub (2010)
and Thomas and Ashbolt (2011). Raw water such as surface water
naturally contains FLP, and ameba counts up to a few thousands
of cells per liter surface water have been recorded, though reduced after flocculation, sedimentation, and filtration (Hoffmann
and Michel 2001; Thomas and others 2008). Generally, a modal
log removal of 1 to 2 log has been observed (Thomas and Ashbolt 2011) prior to disinfection of the treated water. After this
treatment, however, the number of amebae increased in water
distribution systems and in domestic water supplies (Loret and
Greub 2010; Thomas and Ashbolt 2011). Residual organic matter,
biofilm formation, sediments in pipelines, and decline of chlorine
concentration in drinking water are favorable for bacterial regrowth and, hence, their predators. Dead-end legs in distribution
systems, domestic water storage tanks, and premise plumbing offer
even better conditions for amebal growth (Thomas and Ashbolt
2011). Although most of the studies focused on amebae, flagellates and ciliates are also common inhabitants of engineered water
systems (Poitelon and others 2009; Valster and others 2009; Otterholt and Charnock 2011). Not only drinking water supplied by
public water services contains FLP, bottled mineral water has also
tested positive for amebae (Desmet-Paix 1974; Dive and others
1979; Rivera and others 1981; Salazar and others 1982; Fluviá and
others 1983; Penland and Wilhelmus 1999), flagellates (Rivera and
others 1981), and ciliates (Salazar and others 1982). The occurrence of FLP in mineral water is not surprising since FLP inhabit
aquifers (Novarino and others 1997; Risse-Buhl and others 2013).
The number of FLP in groundwater of aquifers increases when
organic material is introduced (Ramirez and others 2006).
Similar to food-related environments, the same conclusion can
be drawn that FLP are part of the microbiota of several foods. Most
published papers report on free-living amebae mainly because research attention was focused on this group of microorganisms.
Especially in older studies, some researchers warned about the
possible health effects after eating lettuce (Napolitano and CollettiEggolt 1984) or drinking bottled mineral water (Rivera and others
1981; Salazar and others 1982) contaminated with acanthamoebae,
a group of amebae, which includes some opportunistic pathogens
(see next section). However, the infection route for Acanthamoeba
spp. was unknown in the past. Currently, oral intake of FLP is not
considered of risk, except for marine toxins produced by some
flagellates (see later). In 2003, the U.S. Environmental Protection Agency announced that no regulatory action was appropriate
or necessary for 9 drinking water contaminants, including Acanthamoeba spp. (US Environmental Protection Agency 2003), and
this opinion was reaffirmed in 2009 (US Environmental Protection Agency 2009). According to the World Health Organization
(WHO), “normal uses of drinking-water lack significance as a
source of infection,” and the WHO therefore stated that “setting a health-based target for Acanthamoeba spp. is not warranted,”
as published in the WHO Guidelines for drinking water quality
(World Health Organization 2011). Finally, the Food and Drug
Administration (FDA, U.S.A.) included Acanthamoeba in its 2nd
edition of the Bad Bug Book (Food and Drug Administration
2012) but emphasized that acanthamoebae were neither implicated in gastrointestinal illness, nor that food was responsible for
diseases such as keratitis and granulomatous amebic encephalitis (see next section). The FDA concluded that food analysis on
Acanthamoeba spp. was not necessary.
A few marine dinoflagellate species (for example, Alexandrium
spp. and Gymnodinium spp.) and several representatives of the diatom genus Pseudo-nitzschia produce neurotoxins (Etheridge 2010;
James and others 2010). Shellfish such as mussels, oysters, and clams
are filter feeders and can accumulate toxins in their tissues. Consumption of contaminated shellfish can cause life-threatening food
intoxication, known as paralytic shellfish poisoning (Etheridge
2010; James and others 2010). In Europe, a limit value of 800 μg
paralytic shellfish poison/kg (measured in the whole body or any
edible part separately) has been set by the EU in Commission Regulation (EC) No 853/2004 (European Union 2004). Food safety
authorities and producers take samples to analyze the presence of
this contaminant. Food poisoning due to eating of commercially
sold bivalves contaminated with marine toxins will therefore be
unlikely.
Acanthamoeba and Tetrahymena: Protagonists in
Bacteria–FLP Interaction Studies
Postingestional survival of bacterial (foodborne) pathogens has
been described for several amebae (Acanthamoeba spp., V. vermiformis, Naegleria spp., Vahlkampfia spp.), ciliates (Cyclidium sp.,
Colpoda sp., Glaucoma sp., Tetrahymena spp.), and the slime mold
(or social ameba) Dictyostelium discoideum. However, most of the
bacteria–FLP interaction studies are performed with members
of the genera Acanthamoeba and Tetrahymena. They have the
930 Comprehensive Reviews in Food Science and Food Safety r Vol. 13, 2014
C 2014 Institute of Food Technologists®
Interactions of foodborne pathogens with FLP . . .
advantage that they are easy to handle, are culturable to high numbers at low costs, are genetically well characterized, and generate
no ethical problems in contrast to animal models.
Acanthamoeba spp. (supergroup Amoebozoa) are flattened,
naked amebae with thorn-like pseudopodia called acanthapodia
(Figure 1). Sizes of trophozoites vary between 12 and 35 μm in
dia, while the cysts are smaller, between 5 and 20 μm in diameter, depending on the species (Khan 2006). Currently, more than
25 Acanthamoeba species have been described, with A. castellanii,
A. polyphaga, A. rhysodes, and A. culbertsoni as frequently studied
species. Based on cyst morphology and size, acanthamoebae were
traditionally divided into 3 groups. However, cyst morphology
may vary upon culture conditions and therefore molecular-based
techniques are increasingly used. 18S rRNA gene sequencing distinguishes 18 genotypes (T1 to T18) (Siddiqui and Khan 2012;
Qvarnstrom and others 2013) with each genotype exhibiting 5%
or more difference between each other. Acanthamoeba spp. are
ubiquitous in aquatic and terrestrial ecosystems but are also frequently isolated from man-made water systems such as tap water,
shower heads (Stockman and others 2011), dental unit waterlines
(Barbeau and Buhler 2001; Trabelsi and others 2010; Hassan and
others 2012), and are recovered from bottled drinking water and
vegetables. Because Acanthamoeba spp. are widespread, human exposure to them is not unlikely. Of 55 healthy individuals tested,
more than 80% had antibodies against A. polyphaga in their blood
(Chappell and others 2001). In 2 other studies, seroprevalence up
to 100% against A. castellanii was recorded (Cursons and others
1980; Brindley and others 2009). Acanthamoeba spp. have been
isolated from human feces, with prevalence ranging from 0.35%
to 10% (de Moura and others 1985; Mergeryan 1991; Zaman and
others 1999), and Acanthamoeba spp. have been cultured from nasal
mucosa from healthy individuals (Červa and others 1973; Michel
and others 1982; Badenoch and others 1988). A few Acanthamoeba
species, including A. castellanii and A. polyphaga, are opportunistic pathogens. Most of the isolates associated with human illness
are grouped in genotype T4. Two major pathologies are recognized: keratitis is an eye infection, mainly related to the use of
acanthamoebae-contaminated contact lens solutions, and amebic
granulomatous encephalitis, which mainly affects immunocompromised patients and is mostly lethal. Fortunately, both infections
are rare. For keratitis, the number of cases is estimated to be 0.01 to
1.49 per 10000 contact lens wearers (Khan 2006). Granulomatous
encephalitis is a secondary infection and the approximate ratio is
estimated to be 1.57 deaths per 10000 HIV/AIDS deaths in the
United States, but it might be higher in countries with warmer
climate (Khan 2006). The morphology of acanthamoebae and the
way prey is engulfed is similar to that of macrophages, which enables researchers to compare cellular features and mechanisms of
internalization between both cell types (Van Waeynberghe and
others 2013). The genome sequence of A. castellanii has been
published by Clarke and others (2013).
Tetrahymena spp. (supergroup SAR, Alveolata, Ciliophora) are
pear-shaped ciliates (Figure 1) with a length of 50 to 60 μm
and a width of approximately 30 μm. At the anterior end of the
cell, tetrahymenae have an oral apparatus, which consists of membranelles (groups of cilia) and an undulating membrane. Tetrahymenae are commonly found in freshwater. The genus Tetrahymena
has over 40 species including several species which are morphologically and physiologically indistinguishable (Simon and others
2008). Barcoding using cox-1 sequences has proven to be an effective approach to identify Tetrahymena spp. (Lynn and StrüderKypke 2006; Chantangsi and others 2007; Kher and others 2011).
C 2014 Institute of Food Technologists®
In contrast with Acanthamoeba spp., most Tetrahymena spp. do not
form cysts (an exception is T. rostrata). Exposure to ethanol might
induce cyst formation in T. pyriformis (Nilsson 2005), but encystment is generally considered as nonexisting for this species. T.
pyriformis and T. thermophila are popular test organisms in bioassays to determine the ecotoxicological effect of chemicals (Sauvant
and others 1999; Gerhardt and others 2010). In addition, T. thermophila is used in many cellular and molecular biology studies.
The genome sequence of T. thermophila is published (Eisen and
others 2006).
Although Acanthamoeba spp. and Tetrahymena spp. are easily
isolated from soil and freshwater samples, strains obtained from
culture collections such as the American Type Culture Collection (ATCC) and the Culture Collection for Algae and Protozoa
(CCAP) are usually used in bacteria–FLP interaction studies, especially A. castellanii, A. polyphaga, T. pyriformis, and T. thermophila.
Interestingly, some culture collection strains grow axenically (without other microorganisms in the growth medium) to high numbers
in chemically defined media or peptone broths.
Intracellular Life of Bacteria in FLP
While most bacteria are digested by FLP, some species have
developed postingestional survival strategies. The term “amebaresistant bacteria” was proposed by Greub and Raoult (2004) to
categorize bacteria, which resist digestion by free-living amebae.
Two groups are distinguished: (i) obligate endobionts and (ii) bacteria with a transient stay in the protozoan host cell. The latter
process implies that an intracellular association with a protozoan
is not part of the life cycle and that bacteria normally live freely
in the environment (for example, soil, water, biofilms) or other
hosts (animals, humans). The type of association with the protozoan host can be neutral, mutualistic (both partners benefit; syn.:
symbiosis), commensalistic (1 partner benefits and the other is unaffected), or parasitic (1 partner benefits and the other is harmed).
The association type is not static and can change depending on
external factors (for example, temperature and food depletion).
Obligate endobionts which often belong to the Alphaproteobacteria, Betaproteobacteria, Gammaproteobacteria, Bacteroidetes, and
Chlamydiae will not be discussed here because the focus of this
review lies on interactions of foodborne pathogens with FLP. For
more information on endobionts, the reader is referred to several
review papers (Görtz 2001; Hausmann and others 2003; Horn
and Wagner 2004; Görtz 2006; Gast and others 2009; Nowack
and Melkonian 2010; Schweikert and others 2013). It should be
noted, however, that endobionts in FLP are not uncommon. For
example, Fritsche and others (1993) reported that 24% of their
examined acanthamoebae harbored endobionts.
A number of human (opportunistic) pathogens (for example,
Burkholderia cepacia, Chlamydophila pneumoniae, Coxiella burnetii,
Francisella tularensis, Helicobacter pylori, L. pneumophila, and Mycobacterium spp.) are able to survive in FLP after uptake (for reviews, see Greub and Raoult 2004; Thomas and others 2010).
Bacteria enter the protozoan cell via phagocytosis, whether or
not receptor-mediated. Remarkably, bacteria such as Aeromonas
hydrophila (Rahman and others 2008) and C. jejuni (AxelssonOlsson and others 2005; Baré and others 2010) bind to specific
sites on the Acanthamoeba cells before uptake, or are collected at
the uroid region (= the posterior end of the locomotive form
of the trophozoite) as seen in L. monocytogenes, Salmonella enterica serovar Typhimurium, Bacillus cereus, and Cronobacter sakazaki
(Doyscher and others 2013). After internalization, bacteria remain
in or escape the phagosome. The fate of internalized, nondigested
Vol. 13, 2014 r Comprehensive Reviews in Food Science and Food Safety 931
Interactions of foodborne pathogens with FLP . . .
Figure 2–Overview of different postingestional survival mechanisms of bacteria in free-living protozoa. (A) Uptake of bacterial cell. (B) Survival of
bacterial cell in a protozoan. (C) Survival and multiplication of bacteria in a protozoan. (D) Multiplication and release of bacteria after lysis of the
protozoan. (E) Release of bacteria through expelled vesicles or fecal pellets. (F) Survival of bacteria in cysts or cyst walls.
bacteria is divided into 3 outcomes (Barker and Brown 1994) zoa but are in fact not true or facultative intracellular pathogens.
Food- and waterborne pathogens which benefit when cocultured
(Figure 2):
with Acanthamoeba spp. are Arcobacter butzleri, A. hydrophila, B.
(i) bacteria which survive without multiplication;
cereus, Shigella spp., S. aureus, V. cholerae, Vibrio parahaemolyticus,
(ii) bacteria which multiply without lysis of the protozoan cell;
and Yersinia enterocolitica (Table 3). Interactions of 4 major food(iii) bacteria which multiply and cause lysis of the protozoan cell.
borne pathogens (C. jejuni, Salmonella spp., E. coli O157:H7, and
In addition to trophozoites, survival in Acanthamoeba cysts was L. monocytogenes) with FLP are discussed below in detail.
reported for F. tularensis (El-Etr and others 2009), L. pneumophila
(Kilvington and Price 1990), several Mycobacterium spp. (Stein- C. jejuni
ert and others 1998; Adékambi and others 2006; Mura and others
C. jejuni cocultured with A. castellanii or A. polyphaga attached
2006; Ben Salah and Drancourt 2010), E. coli (Walochnik and oth- to certain sites of the amoebal cell membrane, resulting in large
ers 1999), and the bacteria Staphylococcus aureus and Vibrio cholerae aggregations of C. jejuni at these sites (Axelsson-Olsson and oth(Table 3). Bacteria such as L. pneumophila are located in cysts while ers 2005; Baré and others 2010). At moderately acidic conditions
mycobacteria are found in the cyst wall. After excystment bacteria (pH 4 and 5), increased adhesion, and internalization was observed
are released into the environment. Survival in cysts enables the (Axelsson-Olsson and others 2010b). At pH 4, about 80% of the
bacteria to survive long periods and harsh conditions such as des- A. polyphaga trophozoites had C. jejuni cells internalized or on the
iccation and disinfection (Kilvington and Price 1990; Adékambi cell surface, compared to about 37% at pH 7. This high percentage
and others 2006).
was obtained when C. jejuni cells were added after acidification
of the coculture medium, suggesting that an acidic environment
Interactions of 4 Major Foodborne Pathogens (C. je- might trigger internalization of C. jejuni into A. polyphaga. Prejuni, Salmonella spp., E. coli O157:H7, and L. monocy- exposure of C. jejuni to environmental stresses such as heat, nutrient starvation, and hyperosmolarity reduced survival in A. casteltogenes) with FLP
Bacteria–FLP interactions of several foodborne pathogens have lanii (Bui and others 2012a). Noteworthy, C. jejuni invasiveness
been reported. However, not all studies confirm intracellular sur- toward Caco-2 cells could not be extrapolated toward amebal cells
vival or replication. The growth of bacteria on metabolic waste since low-invasiveness strains survived better in coculture with A.
products or cellular material of dead protozoan cells is often con- castellanii than high-invasive strains (Baré and others 2010). After
sidered as beneficial for foodborne pathogens. Noteworthy, several uptake, C. jejuni aggregated in amebic vacuoles. Baré and others
foodborne pathogens (for example, E. coli O157:H7, S. aureus, and (2010) found that C. jejuni cells were located in amebic vacuoles
Vibrio spp.) survive transiently in FLP and even multiply in proto- of A. castellanii but that they were not always colocated with acidic
932 Comprehensive Reviews in Food Science and Food Safety r Vol. 13, 2014
C 2014 Institute of Food Technologists®
Interactions of foodborne pathogens with FLP . . .
Table 3–Described interactions between food- and waterborne pathogens and FLP. Interactions of C. jejuni, Salmonella spp., E. coli O157:H7, and L.
monocytogenes with FLP are presented in the text.
Food/waterborne
pathogen
Bacteria
Arcobacter butzleri
Aeromonas
hydrophila
Protozoan
A. castellanii
Acanthamoeba sp.
A. castellanii
A. polyphaga
T. thermophila
B. cereus
A. castellanii
A. polyphaga
A. polyphaga
Shigella dysenteriae
A. polyphaga
A. castellanii
Shigella flexneri
A. castellanii
Shigella sonnei
A. castellanii
A. castellanii
S. aureus
A. polyphaga
A. castellanii, H.
vermiformis
A. polyphaga
Vibrio cholerae
V. cholerae O1, V.
cholerae O54
A. castellanii
A. polyphaga,
Naegleria gruberi
A. castellanii
V. cholerae O139
A. castellanii
V. cholerae O1, V.
cholerae O139
Vibrio mimicus
V. parahaemolyticus
A. polyphaga
A. castellanii
A. castellanii
Y. enterocolitica
Cafeteria
roenbergensis,
Rhynchomonas
nasuta,
Ochromonas sp., T.
pyriformis,
Strombidium sp., A.
castellanii, D.
discoideum
A. polyphaga
A. castellanii
Parasites
C. parvum
T. gondii
Acanthamoeba sp.
Acanthamoeba sp.,
Thecamoeba
quadrilineata
Paramecium
caudatum
A. castellanii
Described effect
Reference
Survival of arcobacters in vacuoles for 10 d. Survival
in vacuoles not fused with lysosomes.
No survival but digestion
Fernández and others (2012);
Medina and others (2014)
Preston and others (2001)
Survival in trophozoites
Intracellular replication (24 h after uptake) of
aeromonads followed by digestion of bacteria
Survival in ciliate depends on virulence A. hydrophila
strain
Survival in trophozoites and cysts
Proliferation of B. cereus in presence of A. polyphaga.
Decrease in number of amebae due to
cytopathologic affect B. cereus?
Growth of amebae in presence of B. cereus.
Extracellular numbers of B. cereus decreased over
1 log after 72 h of co-culture.
Partial lysis of amebae after 5 d co-culture
Increase in S. dysenteriae number with 2 log in
coculture with amebae. Bacteria were present in
cysts.
Amebae enhanced longer survival of bacteria. Killing
of amebae at 37 °C but not at 30 °C. Inhibition of
growth amebae at 30 °C.
Prolonged incubation resulted in gradual change in
morphology of amebae and eventually killing of
amebae
Increase in S. sonnei number with 1 log in coculture
with amebae. Bacteria were present in cysts.
After 24 h coculture, staphylococci found in
phago-lysosomes (50%) and cytoplasma (2%) of
amebae.
Part of the staphylococci survived passage through
the amebic cells. Staphylococci found in fecal
pellets.
Grow of staphylococci (> 3 log cycle after 24 h) in
amebic cells without lysis of the amebae.
Survival in A. castellanii trophozoites and cysts
Survival of vibrios for 24 h in amebae. Survival within
N. gruberi cysts.
V. cholerae O1 grew and survived in trophozoites,
survival in cysts. V. cholerae O54 could only be
found in trophozoites.
Intra-amebic growth and survial of vibrios in the
cytoplasm of throphozoites. Survival in cysts.
Growth and survival of bacteria in amebae for 2 wk
Rahman and others (2008)
Anacarso and others (2011)
Li and others (2011); Pang and
others (2012)
Yousuf and others (2013)
Yli-Pirilä and others (2006)
Huws and others (2008)
Evstigneeva and others (2009)
Saeed and others (2009)
Saeed and others (2012)
Jeong and others (2007)
Saeed and others (2009)
Huws and others (2006)a
Pickup and others (2007)
Anacarso and others (2011)
Cardas and others (2012)a
Thom and others (1992)
Abd and others (2004)
Abd and others (2005)
Sandström and others (2010)
Survival and growth in amebae. Survival in cysts.
Longer survival when cocultured with amebae,
probably due to diffusible factor produced by
amebae. Bacteria do not reside in amebae.
Cytotoxic toward the 7 protists.
Abd and others (2010)
Laskowski-Arce and Orth (2008)
Intracellular replication (24 h after uptake) of
yersiniae followed by digestion of bacteria
Enhancement of survival of yersiniae under
nutrient-rich conditions at 25°C and under
nutrient-poor conditions at 37°C. Intracellular
survival depended on nutrient availability and
temperature.
Anacarso and others (2011)
Amebae served as vehicle for oocysts.
Uptake of oocysts. Proliferation of cryptosporidia in
amebae could not be detected.
Gómez-Couso and others (2007)
Scheid and Schwarzenberger
(2011)
Oocysts were egested in fecal pellets, facilitating in
this way dispersal of oocysts in the environment.
Uptake and survival of oocysts in amebae.
Stott and others (2003)
Matz and others (2011)
Lambrecht and others (2013)
Winiecka-Krusnell and others
(2009)
a Performed experiments with methicillin-resistant S. aureus (MRSA).
C 2014 Institute of Food Technologists®
Vol. 13, 2014 r Comprehensive Reviews in Food Science and Food Safety 933
Interactions of foodborne pathogens with FLP . . .
organelles, suggesting potential bacterial interference with digestive processes. Interestingly, numerous vacuoles containing aggregated viable C. jejuni were egested in the surrounded culture
media by A. polyphaga (Olofsson and others 2013). Coculture
experiments are often performed at 25 °C and 37 °C to simulate
temperatures inside broiler houses or human cells, respectively. C.
jejuni is not able to multiply at 25 °C, but several studies have
shown that C. jejuni survived longer at 25 °C in a medium with
Acanthamoeba spp. compared to a medium without the amebae
(Axelsson-Olsson and others 2005; Snelling and others 2005; Baré
and others 2010; Bui and others 2012b). Cocultivation of C. jejuni with A. polyphaga at 37 °C resulted in rupture of the amebae
and subsequent release of the bacteria into the medium (AxelssonOlsson and others 2005). Intracellular multiplication at 37 °C was
not confirmed by Baré and others (2010) and Bui and others
(2012b) who used A. castellanii in their coculture experiments.
Baré and others (2010) did not find an increase of either bacteria
or amebae, instead a delayed decline and an increased long-term
survival of C. jejuni was observed. Bui and others (2012b) postulated that oxygen depletion by A. castellanii in the medium caused
growth of C. jejuni rather than intracellular replication or growth
on amebal byproducts. Bui and others (2012b) also reported that
no C. jejuni cells could be seen in A. castellanii cysts.
C. jejuni survived longer (up to 36 h) in the presence of T.
pyriformis compared to when tetrahymenae were absent (Snelling
and others 2005). However, replication in ciliates was not determined, although living bacteria were observed in food vacuoles.
Coculture experiments were also performed with other FLP such
as the amebae A. rhysodes, V. vermiformis, Naegleria americana, and
the flagellates Euglena gracilis and Cercomonas sp. (Axelsson-Olsson
and others 2010a; Bui and others 2012c). Prolonged survival of
C. jejuni occurred when cocultured, but replication in these FLP
was not observed, except for A. rhysodes. First and others (2012)
showed that C. jejuni cells survived for 5 h in the ciliate Colpoda
sp., but replication was not determined.
Amebic coculture was proposed as an isolation and enrichment technique for C. jejuni, C. coli, C. lari, and C. hyointestinalis
(Axelsson-Olsson and others 2007). In the protocol, a sample of
environmental or clinical origin is added to a confluent layer of axenically grown A. polyphaga cells. The coculture plate is incubated
for 24 to 48 h at 37 °C to allow enrichment of campylobacters.
The presence of Campylobacter spp. is determined by plating a small
volume of the coculture on blood agar. No genetic loss of C. jejuni
strains was observed when this protocol was followed as revealed
by MLST and flaA typing (Griekspoor and others 2013).
Salmonella spp.
S. enterica serovar Typhimurium resided for at least 4 d in the
contractile vacuole of A. polyphaga and multiplied up to 200 cells
per vacuole (Gaze and others 2003), but this multiplication occurred in less than 1% of the amebae. Salmonella pathogenicity
island 2 was essential in the survival of S. enterica serovar Typhimurium in A. polyphaga (Bleasdale and others 2009). Anacarso
and others (2011) reported intracellular replication of S. enterica
serovar Enteritidis in A. polyphaga, with the first 48 h no detection
of viable salmonellae and a 2 log increase after 72 h. A preferential
uptake of S. enterica serovar Dublin over that of serovar Enteritidis
or Typhimurium by different Acanthamoeba spp. was observed, and
bacteria were most efficiently internalized by A. rhysodes (TezcanMerdol and others 2004). After uptake, salmonellae were located
in membrane-bound vacuoles in which some bacteria replicated.
After 16 h of infection, A. rhysodes cells detached from the surfaces.
A. rhysodes was killed after intracellular growth of S. enterica serovar
Dublin (Tezcan-Merdol and others 2004), most probably caused by
actin degradation by SpvB-dependent ADP-ribosylation (TezcanMerdol and others 2005). S. enterica serovar Typhimurium, S. enterica serovar Typhi, and S. enterica serovar Choleraesuis also caused
cell death of A. rhysodes (Feng and others 2009). Feng and others (2009) found that genes from Salmonella pathogenicity islands
and virulence plasmid were upregulated within A. rhysodes, while
flagella genes of salmonellae were down-regulated. The survival of
S. enterica serovar Typhimurium in A. polyphaga, however, was not
confirmed by Huws and others (2008) who stated that differences
between their results and those of other studies might be attributed
to other bacterial and amebal strains, and also to infection assay and
experimental conditions. Similarly, the study of Douesnard-Malo
and Daigle (2011) showed that S. enterica serovar Typhi was not
able to multiply in amebae. However, they found a longer survival
(up to 10 wk) of salmonellae when cocultured with A. castellanii
compared to free cells, which survived for 10 d under the same
conditions without acanthamoebae.
S. enterica serovar Thompson survived digestion by Tetrahymena
spp. and was subsequently expelled through vesicles (3.1 to 8.0
μm in dia) at the cytoproct of the ciliate (Brandl and others 2005;
Gourabathini and others 2008). Each vesicle contained about 10
bacteria. Salmonellae in vesicles survived for a longer period than
free cells in the same suspension. No cell lysis of tetrahymenae
was detected during the experiments. When internalized, genes
involved in anaerobiosis, virulence, and antibiotic or antimicrobial
resistance were upregulated. Acid resistance genes were strongly
upregulated suggesting an important role in resistance to digestion
(Rehfuss and others 2011).
Glaucoma sp. entrapped S. enterica serovar Thompson in vesicles,
and per cell between 81 and 124 vesicles were expelled (Gourabathini and others 2008). Experiments with an axenic Cercomonas
sp. showed that S. enterica serovar Typhimurium survived longer in
coculture compared to the medium where the flagellate was absent (Bui and others 2012c). Intracellular replication, however, was
not investigated. Coculture of S. enterica serovar Thompson with
Colpoda steinii showed that salmonellae were digested by this ciliate
(Gourabathini and others 2008). The viability of S. enterica serovar
Typhimurium in the rumen ciliate Entodinium caudatum decreased
105 min after engulfment (de la Fuente and others 2010).
E. coli O157:H7 and other pathogenic E. coli
E. coli O157:H7 survived and replicated in A. polyphaga (Barker
and others 1999), although in some trophozoites digestion was
observed. These authors also showed that E. coli O157:H7 was located in outer walls of A. polyphaga cysts. During internalization of
E. coli O157:H7 in A. castellanii, several virulence genes (including
Shiga toxin (Stx) genes and type III secretion system components)
and genes involved in the response to various stressful conditions
(including iron deprivation and oxidative stress) were upregulated
(Carruthers and others 2010). Chekabab and others (2012) showed
that Pho regulon was required for E. coli O157:H7 growth when
cocultured with A. castellanii, while Shiga toxins, however, were
not involved in the interaction. Stx, however, was responsible for
the killing of A. castellanii trophozoites (Chekabab and others 2013;
Arnold and Koudelka 2014). Remarkably, only Stx produced by
internalized bacteria was cytotoxic (Arnold and Koudelka 2014).
Noteworthy, another pathogenic E. coli, the neuropathogenic E.
coli K1, which causes meningitis, is able to survive and multiply in
A. castellanii trophozoites (Alsam and others 2006; Jung and others
2007). The outer-membrane protein A, lipopolysaccharides, and
934 Comprehensive Reviews in Food Science and Food Safety r Vol. 13, 2014
C 2014 Institute of Food Technologists®
Interactions of foodborne pathogens with FLP . . .
type III secretion system were crucial for survival of E. coli K1 in A.
castellanii (Alsam and others 2006; Jung and others 2008; Siddiqui
and others 2011). E. coli K1 also survived encystment (Jung and
others 2007; Siddiqui and others 2011). The enteropathogenic E.
coli O127:H6 strain E2348/69 did not survive predation by A.
polyphaga (Huws and others 2008).
E. coli harboring the Stx2 gene such as E. coli O157:H7 and
an E. coli K-12 modified with Stx2-encoding prophage had an
increased survival in food vacuoles of T. pyriformis compared to
those strains where Stx2 was missing (Meltz Steinberg and Levin
2007). Not only the O157 type of E. coli resisted digestion, also
other serotypes such as O126:NM, O111:NM, and O144:H25
survived digestion by a Tetrahymena sp. (Smith and others 2012).
Two hours after the ciliates were fed with bacteria, fecal pellets
containing viable E. coli bacteria were egested. These fecal pellets had a net-like matrix around the bacteria (Smith and others
2012). Contrary to the aforementioned studies, Nelson and others
(2003) found that their E. coli O157:H7 strain was digested when
fed to T. pyriformis. Also, 2 other studies showed that another
Tetrahymena species, T. thermophila, was killed when fed with E.
coli bearing the Stx-encoding prophage (Lainhart and others 2009;
Stolfa and Koudelka 2013). Release of the Stx toxin prior to digestion of the bacteria was the key to survive digestion (Stolfa and
Koudelka 2013). It was recently hypothesized that Stx production
is a form of “bacterial altruism” where a few bacteria are sacrificed to kill the protozoan predator in order to save the remaining
bacterial population (Mauro and Koudelka 2011; Łoś and others
2013). Hydrogen peroxide production triggered prophage induction, sufficient to produce Stx. However, the authors stated that
not all protozoan species are susceptible to the Stx toxin, and that
various forms of interactions can occur between FLP and E. coli
O157:H7.
Intraprotozoan survival of E. coli O157:H7 is currently mainly
demonstrated for species of the genera Acanthamoeba and Tetrahymena. E. coli O157:H7 resisted digestion of a Glaucoma sp. isolated
from Romaine lettuce. It was subsequently released in vesicles expelled from the ciliate (Gourabathini and others 2008). In another
study, a ciliate (Vorticella microstoma) isolated from dairy lagoon
wastewater failed to digest E. coli O157:H7 during the 13 d of
coculture (Ravva and others 2010). E. coli O157:H7 did not appear to be ingested by rumen ciliate protozoa such as Entodinium
spp. (Burow and others 2005; Stanford and others 2010), while a
potential protective role was suggested for Dasytricha spp. (Stanford
and others 2010). Finally, it was suggested that FLP in composts
might contribute to the reduction of E. coli O157:H7, but the authors did not determine which protozoan species were responsible
for the reduction (Puri and Dudley 2010).
It is worth mentioning that survival of E. coli in FLP is not
only restricted to pathogenic types. Siegmund and others (2013)
showed that E. coli bacteria (a modified K12 strain) were able to
leave food vacuoles and, hence, evade digestion by T. pyriformis.
However, Nilsson (1987) reported that most E. coli bacteria were
digested 2 h after uptake.
L. monocytogenes
L. monocytogenes survived in A. castellanii trophozoites but did not
survive encystment (Ly and Müller 1990a, b). Intracellular replication was observed by Anacarso and others (2011) who found
that the number of listeriae in A. polyphaga increased up to 4
log after 72 h. However, survival or replication of L. monocytogenes in Acanthamoeba spp. has not been confirmed in most of the
other studies (Akya and others 2009a, 2010; Doyscher and others
C 2014 Institute of Food Technologists®
2013). These authors assumed that L. monocytogenes in coculture
with acanthamoebae survived on products released by amebae,
such as metabolites or fragments of dead cells (Zhou and others
2007; Huws and others 2008; Akya and others 2009a; Fieseler and
others 2014). Akya and others (2009a) did not find evidence of
long-term bacterial survival or multiplication within phagosomal
vacuoles or within the cytoplasm of the amebae. Instead, rapid
digestion (4 h after uptake) of L. monocytogenes was observed in
A. polyphaga (Akya and others 2009b, 2010). Expression of listeriolysin O, an important protein necessary to escape vacuoles in
mammalian cells, could not be demonstrated (Zhou and others
2007). Interestingly, Doyscher and others (2013) showed that L.
monocytogenes cells were concentrated in so-called “backpacks” at
the uroid regions. These backpacks contained tens to hundreds of
aggregated L. monocytogenes cells. Backpacks were phagocytosed
and listeriae were killed and digested in food vacuoles. Backpack
formation only took place when L. monocytogenes was cultured at
30 °C prior to coculture and not when cultured at 37 °C. Based
on this observation, Doyscher and others (2013) concluded that
bacterial motility was a prerequisite for backpack formation.
During a 6-d coculture, ingested L. monocytogenes multiplied in
T. pyriformis and subsequently lysed the T. pyriformis cells (Ly and
Müller 1989, 1990a, b). Pushkareva and Ermolaeva (2010) showed
that listeriolysin O played a role in the survival of L. monocytogenes
in T. pyriformis and induced protozoan encystment. However, this
latter observation seems doubtful because T. pyriformis normally
does not encyst. The observed cysts were probably rounded, dead
cells as often seen in old or exhausted cultures. Replication of L.
monocytogenes in a Tetrahymena sp. was not confirmed by Brandl
and others (2005) who found that a small fraction of the listeriae
survived and was expelled in vesicles. Coculture with T. pyriformis
and the same L. monocytogenes strain failed to reveal the presence
of such vesicles (Gourabathini and others 2008).
Glaucoma sp. released small vesicles containing listeriae, although
the number was significantly lower compared to E. coli O157:H7
and S. enterica serovar Thompson (Gourabathini and others 2008).
No significant decrease in L. monocytogenes numbers was observed
when cocultured with a Cercomonas sp., from which Bui and others (2012c) concluded that listeriae were not a food source and
could be cytopathogenic for the flagellates. However, intracellular
replication was not determined. L. monocytogenes was digested by
C. steinii (Gourabathini and others 2008) and 2 other Colpoda isolates, but some bacteria remained undigested and were released in
fecal pellets (Nadhanan and Thomas 2014).
Environmental and Biological Factors
Influencing Intraprotozoan Survival
Most coculture experiments have been performed in vitro, and at
least 3 parameters (temperature, bacterial virulence, and host susceptibility) determined the results. Replication of L. pneumophila in
acanthamoebae is temperature dependent, with multiplication at
35 °C and digestion or limited intra-amoebal growth of legionellae at 20 °C (Anand and others 1983; Ohno and others 2008).
Intracellular growth and replication of C. jejuni in A. polyphaga
was observed at 37 °C but not at 25 °C (Axelsson-Olsson and
others 2005). Temperature control can therefore be important to
minimize or exclude intraprotozoan growth. However, low temperatures do not exclude that (foodborne) pathogens grow on
protozoan metabolites or waste products. Nevertheless, low temperatures remain important to decrease growth rates as most of
the foodborne pathogens cannot replicate at temperatures below
10 °C. Surprisingly, the number of ingested bacteria together with
Vol. 13, 2014 r Comprehensive Reviews in Food Science and Food Safety 935
Interactions of foodborne pathogens with FLP . . .
temperature can determine the outcome. In a study by Kikuhara
and others (1994), it was found that when 30 L. pneumophila
bacteria were ingested by T. thermophila, intracellular replication
occurred at 35 °C but not at 28 °C or 32 °C (only survival).
However, when 10 bacteria were ingested, bacteria were digested
at all temperatures.
The fate of internalized bacteria is furthermore dependent on
bacterial virulence and susceptibility of its protozoan host. Goy and
others (2007) showed that pathogenic M. kansaii strains grew better
in A. castellanii than nonpathogenic strains. In another study, 1 clinical isolate of L. pneumophila showed a 1200% increase in cell count
per ameba, while another clinical isolate failed to grow in amebae
and environmental strains gave intermediate values (Molmeret and
others 2001). Remarkably, L. pneumophila lost its ability to multiply
in A. polyphaga after 50 passages on Legionella-selective agar plates
(Nagl and others 2000). Conversely, the pathogenicity of bacteria toward 1 protozoan species cannot be extrapolated to other
protozoa. Different outcomes were observed when Acanthamoeba
and Hartmannella were infected with various L. pneumophila strains
(Harb and others 1998; Dey and others 2009; Buse and Ashbolt
2011).
Concerns Related to Bacteria–FLP Interactions
tunately, it was not clear from the study whether the acanthamoebae were heavily infected or when internalization of salmonellae
had occurred, or whether water samples were Salmonella-positive.
Similarly, A. hydrophila, B. cereus, and Yersinia pseudotuberculosis were
recovered from amebae isolated from lakes and soil using a coculture assay (Pagnier and others 2008; Evstigneeva and others 2009).
Isolation of foodborne pathogens directly from FLP supports the
hypothesis that FLP can act as a reservoir for these bacteria.
Dispersal of bacteria by FLP in the environment
Internalized bacteria can be spread by motile protozoa, expelled
vesicles, or fecal pellets. Gaze and others (2003) demonstrated
ameba-mediated translocation of S. enterica on agar plates, and
egested salmonellae formed microcolonies along amebal tracks.
Salmonellae were transported several millimetres per day on the
agar surface where they were normally unable to move. The dispersal of bacteria by migrating amebae or swimming ciliates in the
environment is therefore not unlikely. Long-term survival (up to
6 mo) of L. pneumophila in A. castellanii vesicles has been demonstrated (Bouyer and others 2007), and vesicles seemed to be resistant to freeze-thawing cycles of −70 °C to +35 °C (Berk and
others 1998). Olofsson and others (2013) showed that A. castellanii
egested vacuoles containing living C. jejuni cells in the surrounding medium. Similarly, the ciliates T. pyriformis, Tetrahymena sp.,
and Glaucoma sp. are able to expel vesicles loaded with foodborne
pathogens such as S. enterica serovar Thompson, E. coli O157:H7
and, to a lesser extent, L. monocytogenes (Brandl and others 2005;
Gourabathini and others 2008; Smith and others 2012). Up to
50 S. enterica cells per vesicle were recorded (Brandl and others
2005), with dozens of vesicles produced per ciliate (Gourabathini
and others 2008). Production of S. enterica-loaded vesicles by a
Tetrahymena sp. was not only observed in vitro but also on cilantro
leaves, which indicates that FLP play a role in the ecology of foodborne pathogens on produce (Gourabathini and others 2008). As
shown for A. castellanii vesicles, long-term survival of bacteria in
ciliate vesicles or fecal pellets was observed. For example, L. pneumophila inside T. tropicalis-produced pellets remained viable for 90
d (Koubar and others 2011).
Foodborne pathogens and FLP can cooccur in the same foodrelated environments. For example, C. jejuni, Salmonella bacteria,
and FLP were isolated from broiler houses (Snelling and others
2005; Baré and others 2009). From a public health perspective, the
transient stay of pathogenic bacteria in FLP is of concern for several
reasons. First, foodborne pathogens such as C. jejuni, Salmonella,
E. coli O157:H7, and L. monocytogenes are able to multiply in or
in the vicinity of Acanthamoeba spp., a group of amebae which
are widespread in nature and in man-made systems (see previous
sections). So instead of a decrease in bacterial number by digestion,
an increase of bacteria takes place. FLP can serve as a reservoir
and as a vehicle of bacteria and, because of protozoan motility,
bacteria are thus spread further into the environment. Internalized
bacteria are protected against antimicrobial agents, and an increased
virulence after passage through a protozoan has been observed for
several bacteria. Finally, some protozoan species serve as “gene
melting pot” where exchange of genetic elements takes place, FLP as vector or Trojan horse of bacteria
Rowbotham (1980) hypothesized that L. pneumophila could be
which may lead to the development of new bacterial phenotypes.
inhaled by vesicles or infected amebae and subsequently cause
Legionnaires’ disease. Vesicles expelled by A. castellanii and A.
FLP as reservoir of bacteria
Because foodborne pathogens and FLP share the same en- polyphaga have a size between 2 and 10 μm in dia and theoretvironments, it can be hypothesized that FLP can act as hosts ically could contain several hundreds of L. pneumophila bacteria
and contribute to the contamination or persistence of foodborne (Rowbotham 1986; Berk and others 1998). This infection route
pathogens in food-related environments. For example, the occur- was demonstrated by Brieland and others (1997) who found that
rence and persistence of FLP in broiler houses has been demon- amebae with internalized legionellae served as infectious particles
strated (Baré and others 2009, 2011). The combination of FLP and caused pulmonary disease in a murine model.
With regard to foodborne pathogens, a similar result was obin water supplies of broiler houses and their ability to internalize
campylobacters might therefore form an unexplored and under- tained with C. jejuni. In an animal experiment, chickens fed with
rated part within the transmission route of C. jejuni to poultry. In C. jejuni-infected A. castellanii were already after 1 d colonized
another study, amebae isolated from fields not used for grazing by with C. jejuni (Snelling and others 2008).
cattle or not fertilized with animal manure were positive for Mycobacterium avium subsp. paratuberculosis (White and others 2010). Increased virulence of bacteria after passage through
The authors suggested that infected amebae might be of impor- protozoan hosts
tance in the understanding of the epidemiology of paratuberculoIntra-amebal growth of L. pneumophila enhanced invasion of
sis. M. avium subsp. paratuberculosis not only causes paratuberculosis bacteria in murine macrophages and human monocytes (Cirillo
in cattle but has also been related to Crohn’s disease in humans. and others 1994), and amebal-grown legionellae were more inSalmonellae were recovered from 2 Acanthamoeba species isolated fectious in mice compared to buffered charcoal yeast extract agarfrom water and mud from a Polish lake, suggesting that amebae can grown legionellae (Cirillo and others 1999). Similarly, a higher virbe a reservoir of Salmonella spp. (Hadaś and others 2004). Unfor- ulence to mice was reported for intra-amebally grown M. avium
936 Comprehensive Reviews in Food Science and Food Safety r Vol. 13, 2014
C 2014 Institute of Food Technologists®
Interactions of foodborne pathogens with FLP . . .
compared to mycobacteria grown in broth (Cirillo and others
1997).
A hypervirulent phenotype of S. enterica serotype Typhimurium
phage type DT104 was obtained after passage though rumen protozoa as shown by a tissue culture invasion assay and a bovine
infection model (Rasmussen and others 2005). The high invasiveness was related to SO13 gene expression which on its turn
activated hilA overexpression (Carlson and others 2007). Protozoamediated hypervirulence of S. enterica serotype Typhimurium in
calves and pigs was also observed after passage through A. castellanii
(Carlson and others 2007; Xiong and others 2010).
FLP as gene melting pots
Acanthamoeba spp. can serve as “gene melting pots,” “evolutionary cribs,” or “incubators” where lateral gene exchange occurs between eukaryotic hosts, bacteria, and viruses (Moliner and
others 2010; Bertelli and Greub 2012). In contrast with obligate
endobionts, which often show genome size reduction, genome
sizes of facultative intraprotozoan bacteria such as L. pneumophila
are larger compared to their intracellular relatives as a result of receipt of genetic material from other bacteria and amebal hosts.
Genome analysis of L. pneumophila revealed that several genes
encode for eukaryotic-like proteins, suggesting that they play a
role in the intracellular life cycle through mimicking host proteins, which enables the bacterium to invade different eukaryotic hosts (Lurie-Weinberger and others 2010; Gomez-Valero and
Buchrieser 2013). Similarly, 8 open-reading frames in 15 mycobacterial genomes were probably acquired via lateral gene transfer from Firmicutes, Beta- and Gammaproteobacteria and might
have been mediated within protozoan hosts (Lamrabet and others 2012). Conversely, 2.9% of the proteome (450 genes) of A.
castellanii is acquired through lateral gene transfer (Clarke and authors 2013). However, there is currently no indication that gene
transfers have occurred between foodborne pathogens and amebal
hosts. For example, genome analysis showed limited gene acquisition and gene loss within the genus Listeria (den Bakker and others
2010).
From a public health point of view, the potential spread of
antibiotic resistance mediated by protozoan cells is of particular
interest. Transfer of plasmids encoding for kanamycin resistance
and extended-spectrum β-lactamases between 2 E. coli strains was
demonstrated under lab conditions in food vacuoles and fecal pellets of T. pyriformis (Schlimme and others 1997; Matsuo and others
2010; Oguri and others 2011). Exchange of plasmids encoding ceftriaxone resistance from Klebsiella to Salmonella was demonstrated
in the rumen of calves and sheep (McCuddin and others 2006;
Brewer and others 2011).
Protection of internalized bacteria against antimicrobial
agents
Bacteria inside cysts and trophozoites are protected against antimicrobial agents such as disinfectants and antibiotics. Internalized
L. pneumophila were recovered from A. polyphaga cysts exposed to
50 mg/L free chlorine for 18 h (Kilvington and Price 1990).
Mycobacterium spp. within A. polyphaga cysts survived when exposed to 15 mg/L free chlorine for 24 h (Adékambi and others
2006). But also bacteria in trophozoites were protected and survived higher concentrations of biocides compared to free bacteria
(King and others 1988; Whan and others 2006; Garcı́a and others
2007). Remarkably, L. pneumophila grown in A. castellanii showed
a higher susceptibility to chlorine compared to legionellae grown
C 2014 Institute of Food Technologists®
in V. vermiformis when they left the amebae (Chang and others
2009).
The time required for 99% reduction was greatly increased when
C. jejuni, S. enterica serovar Typhimurium, and Y. enterocolitica were
ingested by T. pyriformis (King and others 1988). C. jejuni was 20fold more resistant to free chlorine at 4 mg/L when ingested by
T. pyriformis. At 1 mg/L free chlorine, C. jejuni, S. enterica serovar
Typhimurium, and Y. enterocolitica were more than 50-fold more
resistant when ingested by T. pyriformis while free bacteria were
killed in 1 min when exposed to 0.5 mg/L free chlorine (King
and others 1988). Snelling and others (2005) showed that C. jejuni inside A. castellanii and T. pyriformis was protected against an
iodophor disinfectant applied to sanitize broiler houses. The disinfection resistance was greatly influenced by the age of protozoa
and the coculture time, and a maximum survival of C. jejuni was
obtained when protozoa were 3 d old and at a coculture time of 12
h. Finally, S. enterica in Tetrahymena sp. vesicles survived exposure
to 2 ppm calcium hypochlorite for 10 min (Brandl and others
2005). Also, L. monocytogenes located in Colpoda fecal pellets were
protected against 10% sodium hypochlorite for 24 h (Nadhanan
and Thomas 2014).
Besides disinfectants, antibiotic resistance was observed when
bacteria were located in trophozoites or after they had left the protozoan hosts. M. avium located in A. castellanii showed resistance
against the antibiotics rifabutin, azithromycin, and clarithromycin
(Miltner and Bermudez 2000). It remains unclear if the protection
was due to decreased uptake of the antibiotics through the protozoan cell membrane, metabolic inactivation of the compounds
in the amebae, or selection of a resistant bacterial phenotype.
Intra-amebal grown L. pneumophila were more resistant against the
antibiotics rifampin, ciprofloxacin, and erythromycin compared to
broth-grown legionellae, with even a 1000-fold greater resistance
toward rifampin (Barker and others 1995). However, antibiotic
resistance was lost when amoebal-grown L. pneumophila was cultured afterwards in a nutrient-rich medium. L. pneumophila inside
Tetrahymena tropicalis pellets were more resistant against the antibiotic gentamicin, and this resistance was conserved in legionellae
released from pellets aged for 90 d (Koubar and others 2011). Similarly, L. monocytogenes in Colpoda sp. fecal pellets were protected
against gentamicin (Nadhanan and Thomas 2014).
Resuscitation of viable but nonculturable bacteria
Viable but nonculturable bacteria can be resuscitated when cocultured with Acanthamoeba spp. This has already been observed
for L. pneumophila (Steinert and others 1997; Garcı́a and others
2007) and for C. jejuni (Axelsson-Olsson and others 2005).
Conclusions and Future Research Recommendations
During the last 2 decades interactions of foodborne pathogens
with FLP have been an emerging field of research. Ingested bacteria may survive or even multiply within FLP. Internalized bacteria
are protected against antimicrobial agents and potentially can survive sanitation processes. Consequently, there is a growing concern
that FLP might be responsible for contamination or persistence
of foodborne pathogens in food-related environments. However,
contrary to L. pneumophila, research on interactions of foodborne
pathogens with FLP is still in its infancy. More information is
needed to evaluate the role of FLP in food-related environments
and on food, and the potential health risk, which can arise from
interactions of foodborne pathogens with FLP.
First, the diversity and dynamics of protozoan communities
in food-related environments needs further exploration. Only a
Vol. 13, 2014 r Comprehensive Reviews in Food Science and Food Safety 937
Interactions of foodborne pathogens with FLP . . .
handful of studies have demonstrated that protozoan communities are present but it is unknown whether typical “in-house”
protozoan populations exist and, whether protozoan compositions remain stable over time. In addition, nothing is known
on how FLP respond to desiccation, daily cleaning, and sanitation; and, as a result of these treatments, whether selection for
more resistant FLP occurs. If so, what are the underlying mechanisms for this resistance: cyst formation, species with special cell
characteristics, ineffectiveness of the sanitizers toward FLP, and
so on?
In addition to a profound knowledge of FLP diversity, the conditions in which foodborne pathogens interact with FLP are essential. There are currently conflicting results: some researchers
observe survival and/or replication, while others find no intracellular viability of bacteria in protozoans. Variations in coculture assays (temperature, multiplicity of infection, contact time
between organisms, coculture medium, recovery, quantification,
and viability determination of survivors, actual intracellular multiplication compared with saprophytic growth) and organisms (bacterial strains and protozoan species and strains) make comparisons
between results difficult. In a 1st phase, standardized methods
are necessary to unravel and understand the cellular mechanisms of survival and replication, and under which conditions
these interactions take place. In a next step, coculture experiments should be performed which mimic food-processing or
food-producing environments, including the presence of other
organisms, and at ambient temperatures of food-processing and
food-producing locations. Finally, most interaction studies have
been performed with Acanthamoeba and Tetrahymena species so
far. Although these organisms were found in food-related environments and on foods, there is paucity on information about
the ability of other FLP to act as hosts to foodborne pathogens.
Freshly isolated FLP should be tested on internalized foodborne
pathogens, for example, by means of fluorescent in situ hybridization (FISH). Noteworthy, as several of these environmental isolates likely harbor endobionts, it is interesting to examine in
which way they determine the fate of internalized foodborne
pathogens.
The protective role of cysts for internalized foodborne
pathogens against food sanitizers has not been determined yet.
Currently, only E. coli O157:H7, S. aureus, Shigella spp., and Vibrio spp. have been shown to be localized in Acanthamoeba cysts.
However, no experiments were performed to determine which
disinfectant concentrations are necessary to destroy bacteria inside
cysts, and how long (days, months, years) foodborne pathogens
remain viable inside cysts.
The main focus of this review was on the survival of foodborne
pathogens in trophozoites and cysts. However, the opposite might
be also interesting to investigate: what is the effect of grazing
activities of FLP on microbial communities in food-processing
areas, and can it be turned into a positive direction, to eliminate
foodborne pathogens and spoilage bacteria with the aid of selected
protozoan strain(s)?
In summary, more research is necessary (i) to further characterize FLP communities throughout the food production chain
and (ii) to understand the mechanisms of intraprotozoan survival
of foodborne pathogens, and also to determine whether internalization is a coincidence or an unknown, yet integral, part of the
ecology of these bacteria. Today, too much information is lacking
to estimate health risks. In the case that microbial risk assessments
demonstrate that survival and replication of foodborne pathogenic
bacteria in FLP present a significant health risk, redefinition of
the microbiological quality standards (for example, drinking water
and food-processing environments) with regard to FLP needs to be
considered, and sanitation protocols would need to be reevaluated
to be effective against FLP and internalized foodborne pathogens.
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