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FEMS Microbiology Ecology, 92, 2016, fiw025
doi: 10.1093/femsec/fiw025
Advance Access Publication Date: 8 February 2016
Research Article
RESEARCH ARTICLE
Amoeba-resisting bacteria found in multilamellar
bodies secreted by Dictyostelium discoideum: social
amoebae can also package bacteria
Valérie E. Paquet1,2 and Steve J. Charette1,2,3,∗
1
Institut de Biologie Intégrative et des Systèmes, Pavillon Charles-Eugène-Marchand, Université Laval, Quebec
City, QC, G1V 0A6, Canada, 2 Centre de recherche de l’Institut universitaire de cardiologie et de pneumologie
de Québec, Hôpital Laval, Quebec City, QC, G1V 4G5, Canada and 3 Département de biochimie, de microbiologie
et de bio-informatique, Faculté des sciences et de génie, Université Laval, Quebec City, QC, G1V 0A6, Canada
∗
Corresponding author: 1030 avenue de la medicine, Pavillon Marchand, local 4245, Université Laval, Quebec City, QC, G1V 0A6, Canada.
Tel: +1-418-656-2131, ext. 6914; Fax: +1-418-656-7176; E-mail: [email protected]
One sentence summary: This study shows that the social amoeba Dictyostelium discoideum can package bacteria, revealing a new aspect of microbial
ecology.
Editor: Rolf Kümmerli
ABSTRACT
Many bacteria can resist phagocytic digestion by various protozoa. Some of these bacteria (all human pathogens) are known
to be packaged in multilamellar bodies produced in the phagocytic pathway of the protozoa and that are secreted into the
extracellular milieu. Packaged bacteria are protected from harsh conditions, and the packaging process is suspected to
promote bacterial persistence in the environment. To date, only a limited number of protozoa, belonging to free-living
amoebae and ciliates, have been shown to perform bacteria packaging. It is still unknown if social amoebae can do bacteria
packaging. The link between the capacity of 136 bacterial isolates to resist the grazing of the social amoeba Dictyostelium
discoideum and to be packaged by this amoeba was investigated in the present study. The 45 bacterial isolates displaying a
resisting phenotype were tested for their capacity to be packaged. A total of seven isolates from Cupriavidus, Micrococcus,
Microbacterium and Rathayibacter genera seemed to be packaged and secreted by D. discoideum based on immunofluorescence
results. Electron microscopy confirmed that the Cupriavidus and Rathayibacter isolates were formally packaged. These results
show that social amoebae can package some bacteria from the environment revealing a new aspect of microbial ecology.
Keywords: multilamellar bodies; Dictyostelium discoideum; packaged bacteria; amoeba-resisting bacteria; Cupriavidus;
Rathayibacter
INTRODUCTION
Free-living amoebae (FLAs) like Acanthamoeba spp. are mobile
unicellular protozoa that live in aquatic environments and feed
on bacteria, fungi and algae (Rodriguez-Zaragoza 1994). FLAs can
colonize many man-made infrastructures that provide a favorable environment for the proliferation of microorganisms, espe-
cially where high bacterial population densities are found. Cooling towers (Pagnier, Merchat and La Scola 2009), air conditioners (Walker et al. 1986) and drinking water distribution systems
(Thomas and Ashbolt 2011) are a few examples of man-made infrastructures where FLAs grow (reviewed in Siddiqui and Khan
2012 and Cateau et al. 2014) and regulate bacterial population
densities.
Received: 22 December 2015; Accepted: 2 February 2016
C FEMS 2016. All rights reserved. For permissions, please e-mail: [email protected]
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FEMS Microbiology Ecology, 2016, Vol. 92, No. 3
FLAs capture bacteria by phagocytosis and transfer them to
lysosomal compartments in the phagocytic pathway where they
are usually digested by enzymes (Siddiqui and Khan 2012). However, some bacteria referred to as amoebae-resisting bacteria
(ARBs) are able to avoid or withstand enzymatic degradation in
the phagocytic pathway through various mechanisms and can
survive amoeba predation and lodge inside amoebae (Loret
et al. 2008). ARBs include human pathogenic bacteria such as
Legionella, Chlamydia and Mycobacteria. It has also recently been
shown that the ARB group includes non-pathogenic bacteria
(Kebbi-Beghdadi and Greub 2014).
ARBs can survive and grow within amoebae and may then escape by cell lysis or exocytosis as free bacteria, or by being packaged in fecal pellets, which are usually several concentric layers of lipid membranes known as multilamellar bodies (MLBs).
The secretion of packaged bacteria has been confirmed only for a
number of human pathogens (Legionella pneumophila, Salmonella
enterica, Listeria monocytogenes, Helicobacter pylori and Escherichia
coli O157:H7), but this process has been studied only with FLAs
and protozoa of the ciliate group (reviewed by Denoncourt, Paquet and Charette 2014).
Packaging provides bacteria with a number of advantages
in unfavorable conditions (Berk et al. 1998; Brandl et al. 2005;
Gourabathini et al. 2008; Raghu Nadhanan and Thomas 2014).
For example, S. enterica bacteria packaged in MLBs by the ciliate Tetrahymena are more resistant to low concentrations of calcium hypochlorite than when they are in the planktonic state
(Brandl et al. 2005). Salmonella enterica can even multiply inside
pellets.
The social amoeba Dictyostelium discoideum is a bacterial
predator that lives in damp forest floors. The virulence traits and
host–pathogen relationships of more than 20 pathogenic bacterial species have been studied using this amoeba as a model
(Cosson and Soldati 2008; Bonifait et al. 2011; Dallaire-Dufresne,
Paquet and Charette 2011). Dictyostelium discoideum is often compared to a macrophage-like organism that shares many proteins, such as lysosomal hydrolases involved in intracellular
killing, that are found in specialized phagocytic cells in mammals (Cosson and Lima 2014). Dictyostelium discoideum produces
(Mercanti et al. 2006) and secretes large amounts of MLBs when
fed digestible bacteria (Paquet et al. 2013). While no studies on
bacteria packaging by D. discoideum have been published, inert
polystyrene beads can be packaged in D. discoideum MLBs in presence of digestible bacteria (Denoncourt, Paquet and Charette
2014).
We propose that D. discoideum has also the capacity to package ARBs in MLBs. In the present study, 136 bacterial strains of
various genera and environments were tested for their capacity to resist D. discoideum predation and to determine whether
these newly identified ARBs are packaged in expelled MLBs. As
expected, some ARBs were packaged in D. discoideum MLBs and
were secreted into the extracellular milieu.
Bacteria
MATERIALS AND METHODS
Production of packaged and secreted ARBs
Amoebae
Potential packaged bacteria deduced from the bacteria/amoebae
co-culture results were mixed in a final volume of 300 μL with Ka
using the best ratio determined from previous experiments and
were plated on SM1/10 agar. Drops (5 μL) containing 100 000 D.
discoideum cells were spotted on the bacterial lawns. The plates
were allowed to dry and were incubated for 3 or 4 days at 21◦ C to
obtain large phagocytic plaques. Samples from the peripheries
of the phagocytic plaques were collected using sterile tips. The
Dictyostelium discoideum DH1-10 cells (Cornillon et al. 2000) were
grown at 21◦ C in HL5 medium supplemented with 15 μg mL−1
of tetracycline (Mercanti et al. 2006). The cells were subcultured
twice a week in fresh medium to prevent the cultures from
reaching confluence. They were also grown on bacterial lawns
as described below.
Klebsiella aerogenes was a kind gift from Pierre Cosson (Geneva
University, Switzerland), 19 bacterial isolates were provided by
Martin Filion (Moncton University, Canada) (Filion et al. 2004),
and 78 bacterial isolates were provided by Janet Martha Blatny
et al. (Norwegian University of Science and Technology, Norway)
(Dybwad et al. 2012). All the other isolates used in the present
study were from a drinking water distribution network model
(Berthiaume et al. 2014) or were obtained from ATCC or USDA.
Stock cultures were stored at −80◦ C in LB (EMD, Canada) supplemented with 15% glycerol. As needed, the stock cultures were
thawed and were inoculated on Tryptic Soy Agar (TSA) (EMD,
Canada) plates, which were incubated at 25◦ C, typically for two
days, before being used for the experiments.
Predation resistance assay
Bacterial isolates grown on TSA plates were resuspended in 3 mL
of LB, and the OD at 595 nm was adjusted to 1. The resuspended
bacteria (300 μL) were plated on three different nutrient media
(HL5: bacto peptone (Oxoid) 14.3 g L−1 , yeast extract 7.15 g L−1 ,
maltose monohydrate 18 g L−1 , Na2 HPO4 .2H2 O 0.65 g L−1 , KH2 PO4
0.5 g L−1 and bacto agar 20 g L−1 ); SM: bacto peptone 10 g L−1 ,
yeast extract 1 g L−1 , KH2 PO4 2.2 g L−1 , K2 HPO4 1 g L−1 , MgSO4 1
g L−1 and bacto agar 20 g L−1 ); or SM1/10 (the ingredients for SM
were all diluted 1/10 except for the bacto agar). The plates were
allowed to dry under sterile conditions to obtain bacterial lawns.
The tetracycline from the amoeba cell culture maintenance
was removed by medium replacement, and the D. discoideum
cells were resuspended in fresh HL5 with no antibiotic before
counting them in a hemacytometer chamber. Serial dilutions
were prepared in HL5 medium to obtain the following D. discoideum cell concentrations: 500 000; 50 000; 5000; 500, 50 and
5 cells per 5 μL. The bacterial lawns were spotted with 5 μL of
the serial D. discoideum dilutions. The plates were allowed to dry
and were incubated at 21◦ C for 7 days. They were examined visually for plaque formation on days 1, 3 and 7. The isolates that
did not allow the growth of amoebae were considered as ARBs.
Bacteria/amoebae co-cultures
The identified ARBs were co-cultured alone or were mixed in a
final volume of 300 μL with digestible K. aerogenes (Ka), which is
known to stimulate the production of MLBs (Paquet et al. 2013)
and with 30 prewashed D. discoideum cells. The mixtures were
spread on SM agar plates. Serial Ka:ARB ratios ([99:1], [9:1] [1:1],
[1:9] and [1:99], in a total volume of 300 μL), based on an OD adjusted to 1, were used to determine the best conditions for D.
discoideum growth on bacterial co-cultures. The plates were incubated at 21◦ C for 14 days and were examined visually for phagocytic plaque formation, bacterial colonies within the phagocytic
plaques, or all other anomalous growth on days 3, 9 and 14.
Paquet and Charette
samples were gently diluted in fresh SM1/10 medium and were
processed for immunofluorescence (IF) or transmission electron
microscopy (TEM) as described below.
Immunofluorescence
The samples containing suspended cells and material from the
peripheries of phagocytic plaques were allowed to adhere to
glass coverslips for 3 h and were then fixed in 4% paraformaldehyde for 30 min. The coverslips were rinsed with PBS 1X (1.9 mM
NaH2 PO4 + H2 O; 8.1 mM Na2 HPO4 + 2 H2 O; 154 mM NaCl, pH
7.4) containing 40 mM NH4 Cl to stop the fixation and then with
PBS 1X. The cells were permeabilized for 2 min with methanol
at −20◦ C, and the coverslips were rinsed with PBS 1X and then
with PBS 1X containing 0.2% bovine serum albumin (PBS-BSA)
at room temperature for at least 5 min to block non-specific
binding sites. The adherent cells were then incubated for 45
min with the H36 antibody (Mercanti et al. 2006) diluted 1:1000
in PBS-BSA and then with Alexa 568-coupled anti-mouse IgG
secondary antibody (diluted 1:400; Invitrogen, Canada) and 2.5
μg mL−1 of DAPI (4,6-di-amidino-2-phenylindole diluted in PBSBSA) for 30 min at room temperature in the dark. The coverslips
were washed at least three times with PBS-BSA between each
step. The coverslips were mounted on glass slides using Prolong
Gold (Invitrogen). Images were acquired using an Axio Observer
Z1 microscope equipped with an Axiocam camera (Carl Zeiss,
Canada).
Transmission electron microscopy
Samples from the bacteria/amoebae co-cultures and material
from the peripheries of the phagocytic plaques were collected
using sterile tips and were fixed for 3 h in 0.1 M sodium cacodylate buffer (pH 7.3) containing 2% glutaraldehyde and 0.3% osmium tetroxide. They were washed three times with sodium cacodylate buffer and were dehydrated for 5 min in 30% ethanol, 5
min in 50% ethanol, 5 min in 70% ethanol, 10 min in 95% ethanol
and 1 h in 100% ethanol. The samples were then embedded in
Epon resin and were incubated overnight at 37◦ C followed by
3 days at 60◦ C. Very thin slices (60–80 nm) were cut and were
stained for 8 min with 0.1% lead citrate and then for 5 min with
3% uranyl acetate. They were then examined using a transmission electron microscope (JEOL 1230) at 80 kV.
3
RESULTS AND DISCUSSION
Predation resistance assay
Dictyostelium discoideum is probably the simplest system for assessing bacterial virulence (Hilbi et al. 2007; Froquet et al. 2009).
Because medium richness may have an impact on the results
of predation resistance assays (Froquet et al. 2007; Filion and
Charette 2014), our assays were performed using three different
media of varying composition and richness (HL5, SM, SM1/10 ).
Phagocytic plaques, which are bacteria-free zones due to
amoeba grazing, are produced when amoebae are spotted on
lawns of digestible bacteria (Fig. 1). Phagocytic plaques were
not observed in the presence of ARBs or were observed only
for the highest D. discoideum cell concentrations (Fig. 1C and
D) (Filion and Charette 2014). Ka is used routinely in many
phagocytic experiments to feed D. discoideum, which is why
we used it as a positive control for amoeba predation (Fig. 1B)
(Froquet et al. 2009).
We considered that the isolates were ARBs when 500 or
fewer D. discoideum cells were unable to produce phagocytic
plaques on the bacterial lawn for at least one of the media
tested. For example, it is the case for Cupriavidus sp. and Microbacterium sp. isolates shown in Fig. 1C and D. Isolates that allowed the growth of the amoebae with an initial inoculum of
500 D. discoideum cells per drop or less were considered sensitive to amoeba predation and were rejected for subsequent
experiments.
A total of 136 bacterial isolates were screened with the
amoeba predation assay to identify those that were potential
ARBs. All the experiments were performed twice, and 45 isolates
were considered as D. discoideum resisting bacteria and, as such,
potential candidates for the packaging process (see Table S1,
Supporting Information).
The newly discovered ARBs were not specific to one phylum but belonged to various clades distributed throughout the
prokaryotes, which was in agreement with a study by Moliner,
Fournier and Raoult (2010). Table 1 presents the ARBs discovered
in the present study. Our results suggested that the adaptation
of bacteria to avoid digestion during phagocytosis is widespread
in bacteria. Moreover, the term ARB cannot be generalized and
be applied to an entire genus or species since bacteria from the
same genus or species did not display the same resistance to
predation (Table 1).
Figure 1. Predation resistance assay. (A) Serial dilutions of D. discoideum cells (500 000 to 5 cells/5 μL) were spotted counter clockwise on bacterial lawns on HL5 agar
plates. The plates were incubated for 7 days. The negative control (HL5 medium only) was spotted in the middle of the lawn. (B) Klebsiella aerogenes is sensitive to
predation by amoebae. It was used as a positive control for amoeba predation. Cupriavidus sp. (C) and Microbacterium sp. (D) were resistant to predation and were
considered as potential ARBs.
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FEMS Microbiology Ecology, 2016, Vol. 92, No. 3
Table 1. Taxonomic grouping of new ARBs identified by the predation assay.
Gram
Classa
Positive
Actino
Order
Actinomycetales
Familia
Microbacteriaceae
Micrococcaceae
Bacilli
Negative
Alpha
Beta
Gamma
a
Lactobacillales
–
Rhizobiales
Burkholderiales
Nocardiaceae
Streptomycetaceae
Promicromono-sporaceae
Paenibacillaceae
Staphylococcaceae
Leuconostocaceae
–
Rhizobiaceae
Burkholderiaceae
Enterobacteriales
Comamonadaceae
Oxalobacteraceae
Enterobacteriaceae
Pseudomonadales
Xanthomonadales
Pseudomonadaceae
Xanthomonadaceae
Micrococcales
Bacillales
Genera
Species
No. of
isolates
tested
Microbacterium
Rathayibacter
Kocuria
Micrococcus
Rhodococcus
Streptomyces
Cellulosimicrobium
Paenibacillus
Staphylococcus
Weissella
–
Sinorhizobium
Burkholderia
Cupriavidus
Comamonas
Duganella
Escherichia
Serratia
Pseudomonas
Luteibacter
Microbacterium sp.
Rathayibacter tritici
Kocuria sp.
Micrococcus luteus
Rhodococcus sp.
Streptomyces luridiscabiei
Cellulosimicrobium funkei
Paenibacillus larvae
Staphylococcus sp.
Weissella confusa
–
Sinorhizobium sp.
Burkholderia sp.
Cupriavidus sp.
Comamonas koreensis
Duganella zoogloeoides
Escherichia coli
Serratia grimesii
Pseudomonas sp.
Luteibacter anthropi
8
1
17
23
6
1
1
1
9
1
1
2
3
5
1
1
3
1
13
1
No. of ARB
isolates
3
1
2
9
6
1
1
1
1
1
1
1
3
4
1
1
2
1
4
1
Actino = Actinobacteria; Alpha = Alphaproteobacteria; Beta = Betaproteobacteria; Gamma = Gammaproteobacteria.
Figure 2. Triple co-cultures. Example of potential ARB isolates co-cultured with digestible bacteria (Ka) and 30 D. discoideum cells on SM agar. (A) A lawn of Ka was used
as positive control for phagocytic plaque formation (clear zones in the bacterial lawn; black arrow). (B) A lawn of co-cultured Ka and Luteibacter anthropic [ratio 1:9]. After
the same incubation time, the amoebae were unable to farm the bacterial lawn, and the plaques (black arrow) were much smaller than those of the negative control.
This bacterial species was not retained for subsequent analyses. (C) A lawn of co-cultured Ka and Cupriavidus sp. [ratio 1:9]. Pigmented colonies corresponding to the
Cupriavidus sp. can be seen in the middle of the phagocytic plaques (upper black arrow). Pigmented colonies can also seen around the plaques (lower black arrow).
This isolate was considered as an ARB.
Triple co-cultures
The 45 newly identified ARBs were co-cultured with digestible
bacteria (Ka) and D. discoideum. The goal of this experiment was
to assess the growth of amoebae on digestible bacteria (Ka) in
the presence of ARBs to determine whether the ARBs were toxic
for the amoebae, making it impossible for them to produce packaged bacteria. All the phagocytic plaques with a profile similar
to the positive control, that is, with a large bacteria-free zone
(black arrow, Fig. 2A) due to extensive amoeba growth, were rejected. Similarly, co-cultures where no amoeba growth occurred,
as for the negative control, were also rejected. For example,
all the Ka:Luteibacter anthropic ratios produced small phagocytic
plaques compared to the plaques produced by amoebae grown
only on Ka, suggesting that L. anthropic was toxic to the amoebae or markedly limited their growth (black arrow, Fig. 2B). Conversely, the presence of bacterial colonies in the middle of graz-
ing plaques (black arrow at top, Fig. 2C) or substantial growth
of the ARB around phagocytic plaques (black arrow at the bottom, Fig. 2C) indicated that the ARB was resistant to predation
and had no obvious toxicity for D. discoideum. One possibility is
that the bacteria passed through the phagocytic pathway and
were expelled as packaged bacteria, which then began to grow
and form colonies. Three Cupriavidus and 17 other isolates displayed this profile (Table 2). Thus based on the unusual growth
pattern of amoebae on their lawns, 20 isolates were considered
as ARBs and were retained in order to determine whether they
were packageable.
Bacteria packaging by D. discoideum
The next step was to determine whether D. discoideum cells
were able to package ARBs. Based on previous packaging assays by Gourabathini et al. with E. coli O157:H7 and the ciliate
Paquet and Charette
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Table 2. ARBs identified after co-culture assays as potential candidates for bacteria packaging.
Strains
Ratio KA:ARB
Cupriavidus basilensis
1:1
Cupriavidus sp.
Micrococcus luteus (Norway)
Micrococcus luteus US4
9:1
9:1
1:9
Rathayibacter tritici
Rhodococcus erythropolis US1
Rhodococcus erythropolis US2
Rhodococcus fascians US1
Rhodococcus fascians US2
Cupriavidus necator US1
9:1
1:9
9:1
9:1
1:1
1:1
Duganella zoogloeoides
Kocuria kristinae
Microbacterium oxydans US1
Micrococcus luteus
Micrococcus luteus 8 4 14 × 2
Micrococcus luteus D 1 6 × 2
Micrococcus luteus US3
Rhodococcus erythropolis
Rhodococcus pyridinovorans
Cellulosimicrobium funkei
1:1
1:9
9:1
9:1
1:9
1:9
1:9
1:1
1:9
1:1
Tetrahymena pyriformis (Gourabathini et al. 2008), packaged bacteria released on a rich medium are able to grow inside the package
and break out. Indeed, packaged bacteria are likely a transitory
state, allowing the bacteria to survive in harsh conditions (Berk
et al. 1998; Marciano-Cabral and Cabral 2003) until they are released into an environment that is more favorable for bacterial
growth. Packaged ARBs were not observed during the triple coculture experiments using rich medium even after a long period
of time probably due to growth of potentially packaged bacteria.
On the other hand, starvation media (Smith et al. 2010), which
contains only few nutriments to prevent bacterial growth have
been also tried, but they induce the multicellular development
of amoebae despite the presence of digestible bacteria (data not
shown). Again, no packaged bacteria were seen because active
vegetative D. discoideum cells are required for the packaging process to occur.
The stimulation of bacteria packaging and secretion was also
studied using diluted nutrient agar (SM1/10 ) to avoid rapid bacterial growth following exocytosis that could break up the packages. We observed amoebae on mixed bacterial lawns of digestible bacteria and ARBs (see ratios and strains in Table 2).
Samples collected at the peripheries of the phagocytic plaques
were examined by IF with the H36 antibody (Mercanti et al. 2006)
and by TEM.
A sample containing potential packaged bacteria had to
display combined DAPI and H36 antibody-positive staining for
structures smaller than amoebae but bigger than free-living bacteria (data not shown) due to packaging of bacteria. DAPI would
reveal the presence of bacteria in the structures. On its side, H36
antibody has been shown in a previous study to be a specific
marker of MLBs by binding to a protein still not characterized
(Paquet et al. 2013). The magenta arrows in Fig. 3 point to bacteria
packages measuring 2–3 μm in diameter, and the black arrow indicates a D. discoideum cell. Of the 20 potential candidates tested
by IF, three Cupriavidus isolates, two Micrococcus luteus isolates
Observations and comments
Based on the morphology and color of the
colonies at the center and periphery of the
phagocytic plaques
A few fruiting bodies, with colored spores at
the top.
Several colonies within the phagocytic
plaques.
Unusual growth on agar.
and one isolate each of Rathayibacter tritici and Microbacterium
oxydans presented features suggesting that they were packaged
by D. discoideum.
The same co-culture protocol was performed on several samples to formally confirm the presence of expelled packaged bacteria by TEM. For the control condition shown on Fig. 4, D.
discoideum produced (white arrow, Fig. 4B) and secreted empty
MLBs (black arrow, Fig. 4C) in the presence of digestible bacteria on SM1/10 . However, D. discoideum produced fewer MLBs on
SM1/10 than on rich HL5 medium (Paquet et al. 2013). Despite
this, Cupriavidus sp. and R. tritici were found inside secreted MLBs
when they were co-cultured with amoeba and digestible bacteria (Fig. 4E, F and I). The TEM observations revealed that some of
the tested bacteria could be packaged by D. discoideum.
Interestingly, R. tritici accumulated inside the amoebae, with
up to 50 undigested bacteria visible inside each D. discoideum cell
(Fig. 4H). It is not clear whether the accumulation was due to
rapid bacterial growth inside the amoebae, the inhibition of the
exocytic process, or a combination of both. While the mechanism involved is not known, this result suggested that bacteria
can also survive in harsh environments by residing inside amoebae. The intracellular survival in protozoa of many bacteria has
been described in the past (reviewed in Denoncourt, Paquet and
Charette 2014). Many bacteria of the genus Rathayibacter are phytopathogens of terrestrial plants (Hahn et al. 2003; Schaad and
Schuenzel 2010), and it is likely that amoebae and these soil bacteria interact.
We showed that the packaging of bacteria is possible by D.
discoideum amoeba model and that the phenomenon is not restricted to specific genera. Indeed, both Gram-negative and positive bacteria from various environments, including soil and
water, were trapped inside the MLBs. Moreover, the outcome
of various isolates from a same genera or even a same species
regarding packaging is fairly variable. For example, 23 strains
of M. luteus were tested using the predation assay and 9 were
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FEMS Microbiology Ecology, 2016, Vol. 92, No. 3
Figure 3. Immunofluorescence of bacteria packaged by D. discoideum. Material from the peripheries of phagocytic plaques on lawns of co-cultured bacteria (see ratio in
Table 2) on SM1/10 agar spotted with D. discoideum were processed for IF and were observed under an epifluorescence microscope. For each ARB tested, the differential
interference contrast (DIC) is shown on the left while DAPI (blue), which targets the DNA of bacteria and amoebae, and the H36 antibody (red), which targets MLBs and
the amoeba membrane, staining are presented on the right. Dictyostelium discoideum (black arrow in A) produced and secreted a few packaged Cupriavidus sp. (magenta
arrow) into the extracellular milieu. The bacteria shown on the images (A and B) were coated and recognized by the H36 antibody. In (C) and (D), only a fraction of the
M. luteus and R. tritici cells were in H36-positive structures.
identified as ARBs, two of which were packaged in MLBs based
on the IF results. A total of 13 Pseudomonas strains were also
tested using the predation assay. While four displayed an ARB
phenotype, none was packaged in MLBs. These results indicated that bacterial adaptive evolution with respect to protozoa is complex, as has been shown by the farming of different
strains of Burkholderia sp. by non-farmer D. discoideum (DiSalvo
et al. 2015). Given this, it would be difficult to predict whether a
given bacterial isolate can be packaged or can resist predation
by a specific protozoan without in vitro testing. It would thus be
interesting to determine whether the same ARBs are packaged
by different wild-type strains of D. discoideum or other protozoa.
Lastly, the present study showed that some ARBs are packaged in MLBs and are secreted by D. discoideum in laboratory
conditions. Amoeba/bacteria interactions are ubiquitous in natural as well as in man-made environments such as in municipal drinking water storage tank sediments (Lu et al. 2015), the
floating and fixed biofilms of spring recreation areas (Hsu
et al. 2011), and the surface water of warm water systems and
cooling towers (Kuiper et al. 2006). As such, it is likely that
Paquet and Charette
7
Figure 4. Transmission electron microscopy of bacteria packaged and secreted by D. discoideum. The peripheries of phagocytic plaques from co-cultured bacteria (see
ratio in Table 2) on SM1/10 agar spotted with D. discoideum were processed and were observed by TEM. (A), (D), and (G) Bacteria grown alone on rich medium. (B) and
(C) Dictyostelium discoideum produces (white arrow) and secretes (black arrow) MLBs with digestible bacteria on SM1/10 . No Ka were seen inside the MLBs. (E), (F), and (I)
Cupriavidus sp. and R. tritici were packaged by D. discoideum and were exocytosed into the extracellular milieu. H. More than 50 undigested R. tritici can be seen inside
a D. discoideum cell.
bacteria packaging occurs in real conditions, not just in the
laboratory.
SUPPLEMENTARY DATA
Supplementary data are available at FEMSEC online.
CONCLUSION
The resistance to predation of 136 bacterial isolates was assessed using a standardized D. discoideum predation assay. A total of 45 of these isolates displayed an ARB phenotype and were
co-cultured with digestible bacteria to stimulate MLB production. Twenty potential candidates were retained based on this
screening. The bacteria packaging of seven isolates by D. discoideum was suggested by IF and confirmed for two isolates by
TEM. This is the first study to show that D. discoideum can package bacteria. These results open the way to a better understanding of the role of ARBs in microbial ecology and their persistence
in many environments.
ACKNOWLEDGEMENTS
We are grateful to P. Cosson (University of Geneva, Switzerland)
for the antibodies and bacterial strains. We warmly thank the
teams of J. M. Blatny (FFI, Norway) and M. Filion (University of
Moncton, Canada) as well as the USDA, who provided many
bacterial strains. We thank A. Denoncourt and A. Vincent (Université Laval, Canada) for their critical reading of the manuscript
and Richard Janvier (Plateforme de microscopie, IBIS,
Université Laval, Canada) for acquiring the transmission
electron microphotographs.
8
FEMS Microbiology Ecology, 2016, Vol. 92, No. 3
FUNDING
This work was supported by grants to SJC from the Fonds
de la Recherche du Québec – Nature et Technologies (FRQNT)
[2014-PR-173418], the Chaire de pneumologie de la fondation
J.-D. Bégin de l’Université Laval, the Fonds Alphonse L’Espérance
de la fondation de l’IUCPQ, and the Establishment of young researchers - Juniors 1 program of the Fonds de la Recherche du
Québec en Santé (FRQS) [20004].
Conflict of interest. None declared.
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