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Thesis
Proteomic characterisation of the Mycobacterium marinum-containing
compartment in Dictyostelium discoideum
GUEHO, Aurélie
Abstract
Dictyostelium discoideum is a professional phagocyte using phagocytosis to feed on bacteria.
It is used as a surrogate macrophage to study phagocytosis mechanisms and as a host model
organism to study intracellular bacterial infections. Mycobacterium marinum, as its close
cousin M. tuberculosis, infects innate immune cells where it stops the maturation of its
residing-phagosome to prevent its killing and to establish a compartment where it can
replicate. In the following work, we have used the host-pathogen model system D.
discoideum-M. marinum to better understand how M. marinum manipulates the maturation of
its
containing-phagosome.
We
have
established
a
procedure
to
isolate
mycobacteria-containing compartments. Then, using isobaric labelling and mass
spectrometry, we studied quantitative proteomic modifications applied to the M.
marinum-containing compartment during the first hours of infection. We also compared the
quantitative proteomic composition of compartments containing the pathogenic strain M.
marinum to compartments containing non-pathogenic, avirulent or attenuated mycobacteria
strains.
Reference
GUEHO, Aurélie. Proteomic characterisation of the Mycobacterium
marinum-containing compartment in Dictyostelium discoideum. Thèse de doctorat :
Univ. Genève, 2013, no. Sc. 4597
URN : urn:nbn:ch:unige-372108
Available at:
http://archive-ouverte.unige.ch/unige:37210
Disclaimer: layout of this document may differ from the published version.
[ Downloaded 23/10/2016 at 12:14:02 ]
UNIVERSITÉ DE GENÈVE
FACULTÉ DES SCIENCES
Section de chimie et biochimie
Département de biochimie
Pr Thierry Soldati
Proteomic characterisation of the Mycobacterium marinumcontaining compartment in Dictyostelium discoideum
THÈSE
présentée à la Faculté des sciences de l’Université de Genève
pour obtenir le grade de Docteur ès sciences, mention biochimie
par
Aurélie GUÉHO
de
Augan (France)
Thèse No 4597
GENÈVE
Repromail
2013
Nom de l'Atelier d'Impression
2013
2
Table of content
3 0.1-Résumé.....................................................................................................................................9
0.2-Summary ................................................................................................................................11
1-Introduction .............................................................................................................................13
1.1-Innate immunity ....................................................................................................................14
1.2-Phagocytosis ........................................................................................................................16
1.3-Dictyostelium discoideum, a good model to study innate immunity ....................................17
1.4-The endocytic pathway in Dictyostelium ..............................................................................19
1.5-Phagocytosis in Dictyostelium ..............................................................................................20
1.5.1-Phagosome formation: the ingestion step ................................................................20
1.5.1.1-The phagocytic receptors ...........................................................................20
1.5.1.2-The role of actin .........................................................................................21
1.5.2-Phagosome maturation ............................................................................................24
1.5.2.1-Phagosome acidification ............................................................................24
1.5.2.2-Lysosomal enzymes trafficking .................................................................25
1.5.2.3-Reneutralisation phase and exocytosis ....................................................... 25
1.5.3-Membrane trafficking: the fusion and fission machineries .....................................26
1.5.3.1-Fusion machinery .......................................................................................26
1.5.3.1.1-CORVET .......................................................................................27
1.5.3.1.2-HOPS ............................................................................................27
1.5.3.2-Fission machinery ......................................................................................27
1.5.3.2.1-The retromer.................................................................................. 27
1.5.3.2.2-The WASH complex .....................................................................29
1.6-Intracellular pathogens ..........................................................................................................30
1.6.1-Mycobacterium tuberculosis.....................................................................................30
1.6.1.1-Pathogenesis ...............................................................................................31
1.6.1.2-Manipulation of the macrophage phagosomal pathway .............................32
1.6.1.3-Virulence factors ........................................................................................32
4 1.6.1.3.1-Virulence factors involved in macrophage manipulation ..............32
1.6.1.3.2-Virulence factors involved in intracellular growth .......................34
1.6.1.3.3-Virulence factors involved in phagosome escape and
dissemination.................................................................................................35
1.6.2-Mycobacterium marinum......................................................................................... 35
1.6.2.1-Infection of Dictyostelium .........................................................................36
1.7-Aim of the thesis ...................................................................................................................37
2-Material and methods................................................................................................................39
2.1-Material .................................................................................................................................40
2.1.1-Media ........................................................................................................................40
2.1.2-Buffers and solutions ................................................................................................40
2.1.3-Antibodies.................................................................................................................41
2.1.4-Antibiotics ................................................................................................................42
2.1.5-Kits............................................................................................................................42
2.1.6-D. discoideum cell lines............................................................................................42
2.1.7-Mycobacteria strains.................................................................................................42
2.2-Methods..................................................................................................................................43
2.2.1-Cell culture ...............................................................................................................43
2.2.1.1-D. discoideum cell culture ...........................................................................43
2.2.1.2-Mycobacteria culture...................................................................................43
2.2.2-Samples preparation .................................................................................................44
2.2.2.1-Latex-bead phagosomes isolation .............................................................. 44
2.2.2.2-Mycobacteria-containing compartments isolation ......................................49
2.2.3-Cell biology ..............................................................................................................54
2.2.3.1-Infections .....................................................................................................54
2.2.3.2-Acidification and proteolysis assays ...........................................................55
2.2.3.3-Phagocytosis assay ......................................................................................58
2.2.4-Biochemistry.............................................................................................................60
2.2.4.1-Quantitative mass spectrometry ..................................................................60
5 2.2.4.1.1-TMT labelling ................................................................................60
2.2.4.1.2-OGE ...............................................................................................60
2.2.4.1.3-Mass Spectrometry........................................................................ 60
2.2.4.1.4-Protein identification......................................................................61
2.2.4.1.5-Protein quantification.....................................................................61
2.2.4.2-One-dimensional SDS polyacrylamide gel electrophoresis (1D
SDS-PAGE).............................................................................................................62
2.2.4.2.1-1D SDS-PAGE...............................................................................62
2.2.4.2.2-Western blotting.............................................................................62
2.2.4.2.3-Immunodetection ...........................................................................62
2.2.5-Microscopy ...............................................................................................................63
2.2.5.1-Live imaging .....................................................................................63
2.2.5.2-Immunofluorescence.........................................................................64
2.2.5.3-EM.....................................................................................................65
3-Results.......................................................................................................................................67
3.1-Establishment of an isolation procedure for the mycobacteria-containing compartments ... 68
3.1.1-Latex-beads phagosomes isolation procedure ..........................................................68
3.1.2-Adaptation of the latex-beads phagosomes isolation procedure ..............................68
3.1.2.1-Formation of Bacteria+Beads Complexes (BBCs) .....................................69
3.1.2.2-BBCs are phagocytosed and are recognised as mycobacteria.....................72
3.1.2.3-The BBCs’mycobacteria are still alive and infectious ................................75
3.1.2.4-Isolation of BBCs-containing compartments ..............................................77
3.2-Proteomic characterisation of M. marinum-containing compartments during the
early phase of infection ................................................................................................................79
3.3.1-The M. marinum-containing compartment proteome...............................................80
3.2.2-Comparison of the early M. marinum-containing compartment proteome to the total
phagosome proteome ........................................................................................................83
3.2.3-Proteomic analysis of the temporal modification of the M. marinum-containing
compartment during the early steps of infection................................................................84
6 3.2.4-Quantitative proteomic comparison of non-manipulated phagosomes and
phagosomes manipulated by M. marinum .........................................................................88
3.3-The H+-vATPase is retrieved from M. marinum-containing compartments in a Wash
independant manner during early phase of infection ...................................................................96
3.3.1-WASH, a key factor for the H+-vATPase retrieval from phagosomes.....................97
3.3.2-WASH localises to mycobacteria-containing compartments .................................103
3.3.3-WASH is not a susceptibility factor .......................................................................105
3.3.4-The H+-vATPase retrieval from mycobacteria-containing compartments is WASHindependent ......................................................................................................................106
4-Discussion ...............................................................................................................................109
4.1-The M. marinum-containing compartment, a phagosomal compartment ............................110
4.2-The M. marinum-containing compartment poorly interacts with other endosomal
compartments .............................................................................................................................112
4.3-The role of lipids in mycobacteria infection ........................................................................112
4.4-The WASH complex is not the key player of H+-vATPase retrieval during
mycobacteria infection ...............................................................................................................113
5-Appendix.................................................................................................................................117
5.1- Role of magnesium and phagosomal P-type ATPase in intracellular bacterial killing ......118
5.2- The balance in the delivery of ER components and the vacuolar proton pump to the
phagosome depends on myosin IK in Dictyostelium .................................................................132
5.3- WASH is required for lysosomal recycling and efficient autophagic and phagocytic
digestion .....................................................................................................................................148
5.4-Phagocytosis of mycobacteria is decreased in LmpB ko cells ............................................166
6-References...............................................................................................................................169
Acknowledgments......................................................................................................................186
7 8 0.1-Résumé
Les cellules du système immunitaire inné constituent la première ligne de défense contre les
infections. Elles ingèrent les bactéries par phagocytose pour les tuer et les digérer. Dictyostelium
discoideum est un phagocyte professionnel qui utilise également la phagocytose pour se nourrir de
bactéries. Il peut être utilisé comme un substitut du macrophage pour étudier les mécanismes de la
phagocytose. D. discoideum apparait également comme un nouvel organisme modèle pour étudier les
infections bactériennes intracellulaires. Mycobacterium marinum, tout comme son proche cousin M.
tuberculosis, infecte les cellules du système immunitaire inné. Elle est capable d’arrêter la maturation
du phagosome où elle réside pour éviter d’être tuée et digérée, et pour établir un compartiment où elle
pourra se répliquer. Dans le travail présenté ici, nous avons utilisé le système hôte-pathogène modèle
D. discoideum-M. marinum pour mieux comprendre comment M. marinum manipule le compartiment
où elle réside pour le faire bifurquer de la voie normale de maturation d’un phagosome.
En établissant une procédure pour isoler des compartiments contenants différentes souches de
mycobactéries (pathogènes, non- pathogènes, non-virulentes ou atténuées), nous avons pu étudier le
protéome du compartiment contenant M. marinum pendant les premières heures de l’infection. Ce
protéome n’est pas différent de celui d’un phagosome en terme de composition globale. Cependant, en
utilisant un marquage isobarique couplé à de la spectrométrie de masse, nous avons pu étudier les
modifications protéomiques quantitatives subies par le compartiment contenant M. marinum pendant
les 6 premières heures de l’infection. Nous avons également pu comparer de façon quantitative la
composition protéomique de compartiments contenant la souche pathogène M. marinum à des
compartiments contenant différentes souches mycobactériennes non-pathogènes, avirulentes ou
atténuées à 1 et 6 hpi. Déjà à des temps précoses de l’infection, le compartiment contenant M.
marinum est déplété en protéines impliquées dans l’établissement d’un environnement de dégradation
comme la H+-vATPase ou les enzymes lysosomiales. Il semble que M. marinum bloque la maturation
du compartiment où elle réside en empêchant son interaction avec d’autres compartiments. En effet,
les quantités diminuées, dans le compartiment contenant M. marinum, de petites GTPases et de leurs
régulateurs, de la machinerie de fusion ainsi que de protéines impliquées dans le tri indiquent que ce
compartiment interagit peu avec les compartiments des voies endosomiale et de sécrétion. De plus, les
quantités amoindries de protéines impliquées dans le métabolisme lipidique dans le compartiment
contenant M. marinum ont confirmé que le métabolisme lipidique des cellules hôtes est affecté au
cours d’infections mycobactériennes. Le fait que les protéines liant l’actine soient enrichies dans le
compartiment contenant M. marinum a mis en évidence l’importance du réseau d’actine autour des
compartiments contenant des mycobactéries pendant les premières heures de l’infection.
La comparaison protéomique quantitative de compartiments contenant différentes souches de
mycobactéries a révélé que la H+-vATPase est déplétée des compartiments contenant la souche
pathogène M. marinum. Cependant, une étude précédente du laboratoire, a montré que la H+-vATPase
n’est délivrée que de façon transitoire dans les compartiments contenant M. marinum. Il a été montré
que le complexe WASH induit le recyclage de la H+-vATPase depuis les phagosomes, permettant
ainsi leur reneutralisation. Nous avons examiné si ce complexe était également responsable de la
reneutralisation des compartiments contenant des mycobactéries, et si il pouvait expliquer les quantités
diminuées de H+-vATPase dans les compartiments contenant M. marinum. Malgré le recrutement du
9 complexe WASH sur les compartiments contenant des mycobactéries, la délétion du gène WshA, qui
code pour la sous-unité WshA du complexe WASH, n’a pas d’impact sur le cours de l’infection de M.
marinum. De plus, la mesure du pH des compartiments contenant différentes souches de
mycobactéries a démontré que WASH n’était pas impliqué dans la reneutralisation de ces
compartiments.
10 0.2-Summary
Innate immune cells such as macrophages are the first line of defence against infections. They ingest
bacteria by phagocytosis to kill and digest them. Dictyostelium discoideum is a professional
phagocyte, which also uses phagocytosis to feed on bacteria. It can be used as a surrogate macrophage
to study phagocytosis mechanisms. D. discoideum has also emerged as a new host model organism to
study intracellular bacterial infections. Mycobacterium marinum, as its close cousin M. tuberculosis,
infects innate immune cells. It is able to stop the maturation of the phagosome where it resides to
prevent its killing and to establish a compartment where it can replicate. In the following work, we
have used the host-pathogen model system D. discoideum-M. marinum to better understand how M.
marinum manipulates its containing-phagosome to divert it from the normal phagosome maturation
pathway.
By establishing a procedure to isolate compartments containing different mycobacteria strains
(pathogenic, non-pathogenic, avirulent or attenuated), we could study the proteome of the M.
marinum-containing compartment during the first hours of infection. This proteome is not different
from a phagosome proteome in term of composition. However, using isobaric labelling and mass
spectrometry, we could study quantitative proteomic modifications applied to the M. marinumcontaining compartment during the 6 first hours of infection. We could also compare the quantitative
proteomic composition of compartments containing the pathogenic strain M. marinum to
compartments containing non-pathogenic, avirulent or attenuated mycobacteria strains at 1 and 6 hpi.
Already at very early stage of infection, the M. marinum-containing compartment is depleted of
proteins involved in the establishment of a degradative environment such as the H+-vATPase or
lysosomal enzymes. M. marinum seems to block the maturation of its containing-compartment by
preventing its interaction with other compartments. Indeed, decreased amounts of small GTPases and
their regulators, of fusion machinery and of proteins involved in sorting in the M. marinumcontaining-compartment indicated that this compartment poorly interacts with the other compartments
of the endosomal and secretory pathways. Furthermore, the decreased abundance of proteins involved
in lipid metabolism in the M. marinum-containing compartment confirmed that the host cell lipid
metabolism is affected during mycobacteria infection. The enrichment of actin-binding proteins at the
M. marinum-containing compartment highlighted the importance of the actin-network around
mycobacteria-containing compartments during the first hours of infection.
The quantitative proteomic comparison of mycobacteria-containing compartments also revealed that
the H+-vATPase is depleted from compartments containing the pathogenic strain M. marinum.
However, a previous study in the lab showed that the H+-vATPase is transiently delivered to the M.
marinum-containing compartment. The WASH complex has been shown to retrieve the H+-vATPase
from phagosomes allowing their reneutralization. We investigated whether it was also responsible for
the reneutralization of mycobacteria-containing compartments and whether it could explain the
decreased level of H+-vATPase in M. marinum-containing compartments. Despite the recruitment of
the WASH complex to mycobacteria-containing compartments, the deletion of wshA, which encodes
for the WshA subunit of the WASH complex, did not have an impact on the M. marinum infection
course. Furthermore, pH measurement of compartments containing different mycobacteria strains
revealed that WASH was not necessary for the reneutralization of these compartments.
11 12 1-Introduction
13 1.1-Innate immunity
The immune system protects the body against pathogen invasion. It is divided into two branches: the
innate and the adaptive immunity. The innate immune system is composed of the macrophages, the
dendritic cells, the natural killer (NK) cells and the neutrophils. The adaptive immune system is
mainly composed of the B lymphocytes, the CD4+ and CD8+ T lymphocytes (Messaoudi, Estep et al.
2011).
Figure 1: The immune system is divided into innate and adaptive immunity (Messaoudi, Estep et al. 2011). The
figure shows the cellular components of each immune branch.
Immune cells are segregated into these groups according to the way they recognize pathogens.
Adaptive immune cells have a highly diverse set of receptors, which are specifically designed for
specific antigens. They can also develop immunological memory, allowing better immunological
response efficiency when they are re-exposed to the same antigen. However, the adaptive immune
response is slower than the innate immune response. Innate immune cells, like macrophages,
neutrophils and dendritic cells, are the first line of defense against pathogen invasion. For example,
neutrophils are really motile and can migrate to an infection site, where they are actually the first cells
to arrive. They have a fixed set of receptors called pattern recognition receptors (PRR), which
recognize PAMPs (Pathogen-Associated Molecular Patterns) or more generally MAMPS (MicrobeAssociated Molecular Patterns) (McCullough and Summerfield 2005, Messaoudi, Estep et al. 2011).
After recognition of the pathogen, their role is to ingest it to kill and digest it via a process called
phagocytosis. This process is divided in several steps, which will be further described in the following
sections. A pathogen ingested by phagocytosis resides in an intracellular membranous compartment
called phagosome. Then, the phagosome matures in order to establish a bactericidal and degradative
environment in its lumen. Pathogen degradation products (peptides) can then be loaded onto MHC
class II molecules and exported to the cell surface. However, innate immune cells can also load
pathogen peptides on MHC class I molecules normally used for the presentation of self-antigens or
14 antigens from intracellular pathogens, such as viruses. This process is called “cross-presentation”
(Desjardins, Houde et al. 2005, Jutras and Desjardins 2005, McCullough and Summerfield 2005,
Savina and Amigorena 2007). For this, large pathogen peptides have to be exported from partially
degradative compartments (not yet mature phagosomes) in the cytosol where they are further digested
by the proteasome. Antigen peptides are then transported to ER where they are also loaded onto MHC
class I and exported at the plasma membrane. It has also been shown that soon after phagocytosis, the
phagosome fuses with the ER to form a phagosome-ER mix compartment. After digestion by the
proteasome, the antigen peptides can also be transported back to this immature ER-phagosome where
the conditions are permissive for loading onto MHC class I molecules (Guermonprez, Saveanu et al.
2003, Jutras and Desjardins 2005). With those processes, the innate immune cells become the socalled antigen presenting cells (APC). The ability of a cell to present antigen is dependent of its
degradative efficiency. The more a cell has phagosomes with strong degradative conditions, the less
efficient it is to load antigens on MHC molecules. Dendritic cells are particularly efficient in
presenting antigens. Once presented at the innate immune cell surface, antigens can be presented to
adaptive immune cells leading to their activation. This antigen presentation then links innate and
adaptive immunity (Desjardins, Houde et al. 2005, Jutras and Desjardins 2005, McCullough and
Summerfield 2005, Savina and Amigorena 2007).
Figure 2: Antigen processing and loading onto MHC molecules in phagosomes (Jutras and Desjardins 2005).
Pathogen peptides generated by pathogen degradation can be loaded on MHC class II molecules and exported to
the cell surface. Pathogen peptides can also be presented by MHC class I molecules by a process called “cross
presentation”. Large pathogens peptides are exported in the cytosol were they are further digested by the
proteasome. They are then transported to the ER where they are loaded on MHC class I molecules and exported
to the plasma membrane. After proteasome degradation, they can also be transported back to immature ERphagosomes where the conditions are permissive for loading onto MHC class I molecules.
15 1.2-Phagocytosis
Phagocytosis is the process by which phagocytic cells ingest large particles (>200 nm). Once ingested
by the cell, the particle resides in a closed phagosome. The classical ingestion process is described as
the “zipper model”, meaning that the plasma membrane of the phagocytic cell is closely apposed to
the particle to ingest by a ligand-receptor interaction (Haas 2007). Indeed, the first step of
phagocytosis consists in the recognition of the particle by receptors present on the cell surface. This
binding event activates signaling pathways in order to recruit proteins involved in F-actin formation at
the binding site, underneath the membrane. Actin polymerisation allows the deformation of the plasma
membrane and the extension of the circular, cup-like lamellipod around the particle (Haas 2007,
Bozzaro, Bucci et al. 2008). Once the particle is totally surrounded by a membrane, the membrane
closes. The particle is then in a phagosome.
The phagosome follows a well organized maturation program that allows the killing and
digestion of the ingested particle. Maturation is a highly dynamic membrane trafficking process,
which involves fusion with endosomes but also protein sorting and membrane recycling by fission
events. By its interaction with the endosomal pathway, the phagosome acquires proteins involved in
the establishment of a bactericidal and degradative environment in its lumen. However, it is not clear
if the phagosome completely fuses with those different endosomes. It seems that it is more a “kiss and
run” process (Haas 2007, Flannagan, Cosio et al. 2009). The lipid bilayers of the two organelles
transiently fuse allowing a short exchange of contents. Then the vesicle rapidly disconnects from the
phagosome. The maturation process can be seen as sequential fusions with first early endosome
vesicles, then late endosome vesicles and finally lysosomal vesicles. Indeed, those different endosome
populations are well segregated and fusion between them follow strict rules (Haas 2007, Flannagan,
Cosio et al. 2009). Early endosomes can fuse homotypically and with newly formed phagosomes and,
a bit more rarely, with late endosomes. However, they cannot fuse with lysosomes. Late endosomes
can fuse homotypically and less frequently with lysosomes. So, by fusing with an early endosome, the
newly formed phagosome becomes an early phagosome, favorising homotypic fusions with other early
endosomes. The early phagosome can also fuse with late endosomes, again progressively favorising
homotypic fusions with later endosomes, until it becomes a late phagosome. Finally, the late
phagosome can also fuse with lysosomes and become a phagolysosome. At each step of the
maturation, some components are also retrieved from the phagosome by fission. They can be recycled
back to the plasma membrane or to the endosome population they were originally coming from
(Flannagan, Cosio et al. 2009).
The recognition of a particle by a receptor induces the assembly of the NADPH oxidase
complex at the membrane. This complex produces reactive oxygen species (ROS). After phagosome
closure, these toxic compounds, potentially bactericidal, accumulate in the phagosome (Bokoch and
Zhao 2006, Bylund, Brown et al. 2010). Late endosomes deliver the H+-vATPase to the phagosome.
This protein complex pumps protons inside the phagosome in an ATP dependant manner. This leads
to the acidification of the phagosome. Upon fusion with lysosomes, the phagosome finally acquires
hydrolases that are active at low pH and allow the digestion of the ingested particle (Jutras and
Desjardins 2005, Haas 2007, Bozzaro, Bucci et al. 2008, Flannagan, Cosio et al. 2009)
16 Figure 3: Phagosome maturation pathway (Haas 2007). To establish a degradative environment in their lumen,
phagosomes mature by sequential fusion with early endosomes, late endosomes and lysosomes to form early
phagosomes, late phagosomes and finally phago-lysosomes.
Phagocytosis is a very ancient process not originally used for immunity purposes. It is also
observed in unicellular organisms such as the social amoeba Dictyostelium discoideum. Protozoan
generally use phagocytosis to ingest and digest bacteria for nutrition (Desjardins, Houde et al. 2005).
Phagocytosis in Dictyostelium will be further described in the following parts.
1.3-Dictyostelium discoideum, a good model to study innate immunity
Dictyostelium discoideum is a social amoeba that lives on the forest soil. It uses phagocytosis to feed
on bacteria. It is a very motile cell able to chase its prey before its ingestion. Dictyostelium has the
particularity to have two different stages in its life cycle: a growth phase or vegetative phase, and a
developmental phase. At the vegetative stage, Dictyostelium cells grow as single cells. But when
bacteria are no longer available in their environment, the Dictyostelium cells start to secrete and to
respond to the chemoattractant cAMP. They become “social”. They aggregate and adhere to each
other to form a multicellular organism. The so-formed slug can migrate on surfaces towards light and
along temperature gradients. It can then form a fruiting body, harbouring the spores, which can
germinate to start a new life cycle (Bozzaro 2013).
The Dictyostelium genome has been fully sequenced (Eichinger, Pachebat et al. 2005). It is
haploid and composed of 6 chromosomes encoding 12,000 protein-coding genes. The simplicity of
this genome, combined to efficient homologous recombination allow the easy generation of knock-out
mutants, as well as a multitude of strains expressing fluorescent reporter-tagged proteins.
17 Figure 4: Dictyostelium life cycle (Bozzaro 2013). Dictyostelium life cycle is divided in two phases, a vegetative
phase and a development phase. During vegetative phase, cells feed on bacteria by phagocytosis and grow as
individual cells. In starvation conditions, cells aggregate to form a multicellular organism. The formed slug can
migrate and form a fruiting body harbouring a spore. The spore can germinate to start a new life cycle.
Dictyostelium has been used for decades to study its phagocytic capacity. As a professional
phagocyte, it can not only ingest bacteria but also yeast, dead cells and inert particles like latex beads.
It usually performs phagocytosis at higher rates than neutrophils and macrophages. Furthermore, even
if phagocytosis is used by Dictyostelium for feeding purposes, the molecular mechanisms of
phagocytosis are well conserved between innate immune cells and Dictyostelium (Desjardins, Houde
et al. 2005, Jin, Xu et al. 2009). As demonstrated by recent proteomic studies on isolated latex-bead
phagosomes, 70% of the mammalian phagosomal proteome is also found in the Dictyostelium
phagosome proteome (Boulais, Trost et al. 2010).
The study of the interactions of Dictyostelium with bacteria started more than 30 years ago. In
1978, Depraitere and Darmon identified bacteria pathogenic for Dictyostelium. But it is only 10 years
ago that it started to be used as a host model for host-pathogen interaction studies. The establishment
of growth assays on different bacteria strains lawn (Klebsiella pneumoniae, Pseudomonas aeruginosa,
Bacillus subtilis, Staphylococcus aureus) has allowed the identification of proteins involved in the
phagocytosis or in the killing of those different bacteria, but also allowed to test the virulence of
numerous bacteria (Benghezal, Fauvarque et al. 2006, Froquet, Lelong et al. 2009, Lelong, Marchetti
et al. 2011). The use of this assay has allowed the isolation of P. aeruginosa avirulent strains and the
identification of two virulence pathways used by this bacterium: quorum-sensing mediated virulence
and type III secretion of cytotoxins (Pukatzki, Kessin et al. 2002). It has also allowed the identification
of a new secretion system important for Vibrio cholerae virulence (Pukatzki, Ma et al. 2006). It has
even been demonstrated that Dictyostelium can also be infected with human intracellular pathogens
such as Legionella pneumophila (Solomon, Rupper et al. 2000, Urwyler, Nyfeler et al. 2009, Finsel,
Ragaz et al. 2013), Salmonella typhimurium (Jia, Thomas et al. 2009, Sillo, Matthias et al. 2011) or
18 Mycobacteria (Solomon, Leung et al. 2003, Hagedorn and Soldati 2007, Hagedorn, Rohde et al.
2009). Its use has allowed the identification of both host and pathogen factors important for the
infection with these different intracellular pathogens (Steinert and Heuner 2005, Bozzaro, Bucci et al.
2008, Bozzaro and Eichinger 2011).
Mechanisms of phagocytosis and phagosome maturation in Dictyostelium will be described in
the following sections.
1.4-The endocytic pathway in Dictyostelium
As in mammalian phagocytic cells, the endocytic pathway of Dictyostelium is composed of different
populations of endosomes. The endosomes are segregated in early endosomes, late endosomes and
lysosomes according to their acidity and to the protein markers that they carry. In a very simplified
view, Rab5 is characteristic of the early endosomes, Rab7 of the late endosomes, and the lysosomal
enzymes of the lysosomes. The early endosomes are mildly acidic whereas late endosomes and
lysosomes are very acidic (Maniak 2002, Maniak 2003).
Dictyostelium cells can ingest both solid particles by phagocytosis but also fluids by
macropinocytosis. Both phagosomes and macropinosomes mature by interacting with the different
endosomes of the endocytic pathway. This allows them to acquire proteins necessary for the digestion
of the ingested food. They transiently fuse with endosomes in a process called “kiss and run”. This
transient fusion allows two compartments to exchange their content. But as described earlier, the
maturation is a well organized program. Indeed, endosomes fusions follow strict rules. Early
endosomes can fuse homotypicaly, with new-formed phagosomes and macropinosomes, and
sometimes with late endosomes. Late endosomes can fuse homotypicaly and with lysosomes.
Consequently, the maturation program can be seen as a linear process with first, fusion with early
endosomes, then late endosomes and finaly lysosomes (Maniak 2002, Maniak 2003). These fusion
events are also controlled by SNARE (soluble N-ethylmaleimide-sensitive fusion protein attachment
protein receptor) proteins present on the endosomes membrane. t-SNARE present on the target
compartment and v-SNARE present on the donor compartment form a complex. This permits to put in
close proximity the two compartments which have to fuse together (Bogdanovic, Bruckert et al. 2000,
Weidenhaupt, Bruckert et al. 2000, Bogdanovic, Bennett et al. 2002).
The endocytic pathway is also characterized by numerous fission events. As a professional
phagocyte, Dictyostelium continuously ingest particles and fluids. This implies the use of large amount
of plasma membrane to form phagosomes and macropinosomes membranes. To maintain its size and
keep internalizing food, Dictyostelium needs to recycle membrane from phagosomes and
macropinosomes back to the plasma membrane (Neuhaus and Soldati 2000, Neuhaus, Almers et al.
2002). These early recycling endosomes are p25-positive (Ravanel, de Chassey et al. 2001, Charette,
Mercanti et al. 2006). During phagosome maturation, different types of lysosomal enzymes are
delivered to the phagosome. However, these different enzymes do not colocalise and the first set of
enzymes is retrieved from the phagosome and recycled to lysosomes before adding the second set of
lysosomal enzymes (Buczynski, Bush et al. 1997, Souza, Mehta et al. 1997, Gotthardt, Warnatz et al.
19 2002). Unlike macrophages, the phagosome maturation in Dictyostelium has an additional late step,
which consists in the exocytosis of undigested remnants. Before exocytosis, Dictyostelium proceeds to
recycling of some phagosomal components back to endosomes (Neuhaus, Almers et al. 2002). The
H+-vATPase, responsible for phagosome acidification, is retrieved and recycled to late endosomes
(Clarke, Maddera et al. 2010). This process is dependent of the WASH complex, which will be further
described in the following section (Carnell, Zech et al. 2011).
Figure 5: Endosomal pathway in Dictyostelium (Gotthardt, Blancheteau et al. 2006). To mature, phagosomes
and macropinosomes interact with the endosomal pathway. Their maturation is characterized by sequential
fusion with the different populations of endosomes: early endosomes, late endosomes and lysosomes. During the
maturation process, some materials are sorted by fission events and recycled.
1.5-Phagocytosis in Dictyostelium
1.5.1-Phagosome formation: the ingestion step
1.5.1.1-The phagocytic receptors
The first step of phagocytosis consists in the recognition of the particle to ingest by receptors present
on the cell surface of Dictyostelium cells. These cells can engulf a large variety of particles and so, are
expected to have a large repertoire of receptors. However, only few receptors have been identified up
to now. In 1980, it has been shown that at least 3 types of receptors are present at the Dictyostelium
cell surface: a lectin, a receptor for hydrophobic surfaces and a receptor for hydrophilic surfaces.
However, the identity of those different receptors is not known (Vogel, Thilo et al. 1980).
More recently, the establishment of mutant libraries using REMI (restriction enzyme-mediated
insertion) has allowed the screening of thousands of mutants for their adhesion properties and their
phagocytic capacity. Those screens have permitted the identification of different proteins involved in
adhesion. Among them, the Sib family (Similar to Integrin Beta) has been identified. It is composed of
20 5 proteins, SibA to E, which display structural similarities to human β-integrins. They have a
cytoplasmic domain, also present in human β -integrins, which binds talin, connecting extracellular
space and cytosolic machinerie. Both sibA and sibC disruption lead to decreased phagocytosis of latex
beads but not of K. pneumoniae and decreased adhesion to generic surfaces. Those Sib proteins might
be the receptors for hydrophilic surface initially found by Vogel (Cornillon, Gebbie et al. 2006,
Cornillon, Froquet et al. 2008).
Phg1A was also considered as a potential receptor. It is a 9 transmembrane domains protein.
Its deletion leads to decreased phagocytosis of particles with hydrophilic surface (K. pneumoniae,
E.coli and latex beads). However, Phg1a does not seem to be a real receptor. Instead, it regulates the
cell surface composition. Indeed, it has been shown that Phg1a regulates the amount of SibA at the
cell surface by both controlling SibA production and targeting to the cell surface (Cornillon, Pech et
al. 2000, Benghezal, Cornillon et al. 2003, Froquet, le Coadic et al. 2012).
SadA (Substrate Adhesion Deficient mutant A) is also a 9 transmembrane domains protein. It
has 3 conserved EGF-like repeats. Its deletion leads to decreased phagocytosis of latex beads, K.
pneumoniae and E. coli. A recent study showed that SadA is also involved in the regulation of
Dictyostelium cell surface. As Phg1a deletion, the deletion of SadA leads to decreased amount of SibA
at the cell surface (Fey, Stephens et al. 2002, Froquet, le Coadic et al. 2012).
1.5.1.2-The role of actin
Actin is a 42 kDa protein existing in two different states. G-actin is the globular, monomeric form.
The polymerization of actin leads to the formation of F-actin or filaments.
During phagocytosis, the binding of a receptor to a particle activates a signaling cascade. This
leads to the recruitment at the binding site, below the membrane, of numerous proteins involved both
in actin polymerization and in the regulation of this process. The formation of actin filaments will
deform the membrane. The elongation of actin filaments then leads to the extension of pseudopods
around the particle to ingest. Actin clearly plays a crucial role during the ingestion phase. Indeed, the
use of actin depolymerising drugs abolishes uptake (Maniak, Rauchenberger et al. 1995).
Actin polymerization is induced by 2 major actin-nucleating families: the Arp2/3 (Actin
Related Protein 2/3) complex and the formins. Arp2/3 is involved in actin nucleation whereas formins
are more involved in actin filaments elongation. However, the Arp2/3 complex itself is inactive. It is
activated through binding to its regulators, the nucleation promoting factors: the Scar/WASP
(Suppressor of cAMP receptor/Wiskott-Aldrich syndrome protein) proteins. Both are recruited to the
phagocytic cup within seconds (Insall, Muller-Taubenberger et al. 2001, Seastone, Harris et al. 2001).
G proteins of the Ras and Rac families are also involved in the regulation of the actin
cytoskeleton and are recruited at the phagocytic cup. Among the eight Ras genes that Dictyostelium
possesses, only two, RasS and Rap1, are involved in phagocytosis. The RasS-null mutant and cells
expressing the dominant-negative form of Rap1 both show a phagocytosis defect (Seastone, Zhang et
21 al. 1999, Chubb, Wilkins et al. 2000). It has been shown in vitro that both of them interact with the
serine/threonine kinase Phg2 (Gebbie, Benghezal et al. 2004). Interestingly, the phg2-null mutant has
an actin polymerization defect and is defective for the phagocytosis of hydrophilic substrates. Among
the eighteen Rac genes present in the Dictyostelium genome, only Rac1, RacB, RacC, RacG and RacH
regulate phagocytosis. Rac1 overexpression induces the formation of extended filopodia. It leads to a
higher phagocytosis rate. On the contrary, cells expressing the dominant-negative form of Rac1 have a
phagocytosis defect (Dumontier, Hocht et al. 2000). The expression of both the constitutively active
form and the dominant-negative form of RacB lead to decreased phagocyosis (Lee, Seastone et al.
2003). RacC is a WASP activator, and is consequently involved in actin polymerization (Han, Leeper
et al. 2006). RacG is also involved in actin polymerization via Arp2/3, but in a WASP-independent
manner. Its overexpression or the expression of its constitutively active form increases the rate of
uptake. RacG disappears from the phagosome immediately after particle internalization (Somesh,
Vlahou et al. 2006). Finally, RacH overexpression leads to increased phagocytosis but its absence does
not affect uptake (Somesh, Neffgen et al. 2006).
The actin cytoskeleton formation is also regulated by phosphatidylinositol lipids or
phosphoinositides (PIPs) metabolism. PIPs are glycerolipids. Their D-myo-inositol head group can be
phosphorylated or dephosphorylated in different positions (3, 4 or 5) by different PI kinases and PI
phosphatases. This gives rise to different lipid species, which are segregated in the different
membranes of the cell. Several effector proteins with specific binding domains (PH, PX, FYVE,
ENTH/ANTH and FERM) can bind those lipids.
PI(4,5)P2 (phosphatidylinositol, 4-5, bisphosphate) is found in the plasma membrane. Actin
binding proteins and actin nucleating factors can bind PIP2 allowing the recruitment at the plasma
membrane of the protein machinery involved in actin polymerization and actin cytoskeleton
formation. PI(4,5)P2 is also the target of PLC, PI3K and Dd5P4. Dictyostelium PLC is similar to
mammalian PLC-δ. It cleaves PI(4,5)P2 into DAG (Diacylglycerol) and I(1,4,5)P3. DAG is a
stimulator of PKC whereas IP3 increases intracellular Ca2+. Both PLC and Ca2+ have been shown to be
important during the ingestion step (Peracino, Borleis et al. 1998, Seastone, Zhang et al. 1999, Yuan,
Siu et al. 2001). PI3K phosphorylates PI(4,5)P2 on the position 3 of its head group. This forms
PI(3,4,5)P3, a constituent of the membrane of the future closed phagosome. On the other hand,
Dd5P4, the OCRL-1 homologue, dephosphorylates PI(4,5)P2 to form PI(4)P. Dd5P4-null mutants
show a defect in yeast phagocytosis (Loovers, Kortholt et al. 2007).
22 Figure 6: Phosphoinositides metabolism during phagocytosis (Bozzaro, Bucci et al. 2008). Relative abundance
of different phosphoinositides and actin-binding proteins at the phagocytic cup.
Furthermore, not only actin nucleating proteins are recruited to the phagocytic cup. Numerous
other proteins with different roles in actin dynamics are observed. Some proteins anchor the actin
cytoskeleton into the membranes like talin, comitin or ponticulin. The role of some other proteins is to
maintain a sufficient amount of G-actin available for rapid actin polymerization. These proteins also
called severing proteins, induce F-actin fragmentation. Cofilin and its regulator Aip1 (actin interacting
protein 1) are involved in this process. They are both recruited to the phagocytic cup. Aip1 null
mutants have a phagocytosis defect (Aizawa, Katadae et al. 1999, Konzok, Weber et al. 1999). The Gactin generated by severing proteins can then bind G-actin binding proteins like profilins. These
proteins allow the recruitment of G-actin to elongating actin filaments. Coronin, an other actin
binding protein, is also present at the phagocytic cup. Its deletion leads to phagocytic defect (Maniak,
Rauchenberger et al. 1995).
The membrane deformation at the phagocytic cup is not only triggered by actin itself. This
process is driven by an other class of actin-binding proteins called myosins. MyoII, Myo1B, Myo1C
and Myo1K are present at the phagocytic cup (Dieckmann, von Heyden et al. 2010). Those proteins,
which are also called motor proteins, have a structure allowing them to bind both to actin via a linker
protein and also to the membrane. Myosins are composed of three domains: a N-terminal conserved
motor domain, a neck domain and a C-terminal tail. The motor domain and the neck domains are
involved in the movement along actin filaments. The motor domain binds F-actin in an ATP
dependant manner. The tail is the most variable part. In most Myosins I, it is composed of three subdomains: tail homology 1 (TH1), TH2 and Src homology 3 (SH3). TH1 is a lipid-binding domain,
allowing the binding to membranes. TH2 binds actin in an ATP-independent manner. Myo1B and C
are normal long-chain tail myosins (Soldati, Geissler et al. 1999). However, Myo1K is lacking the
neck and tail part. Instead, it has an oversized head with an inserted TH2-like domain and a short
farnesylated tail (Schwarz, Neuhaus et al. 2000, Dieckmann, von Heyden et al. 2010). Myo1B and
Myo1C bind indirectly to actin through a linker called CARMIL (Capping, Arp2/3, Myosin1 Linker).
23 This linker binds both to the SH3 domain of the myosins and to the actin-nucleating protein Arp2/3
(Jung, Remmert et al. 2001). Myo1K binds to actin via the Actin Binding Protein 1 (Abp1). Abp1
binds the inserted TH2-like domain of Myo1K head. Furthermore, MyoK is regulated by the kinase
PakB which is itself activated by the small GTPase Rac1 (de la Roche, Mahasneh et al. 2005,
Dieckmann, von Heyden et al. 2010). As Myosins bind both to the plasma membrane and to actin, by
walking along actin filaments via their motor activity, they push the membrane away from actin and
appose it to the particle. This leads to the deformation of the plasma membrane around the particle to
ingest (Clarke, Engel et al. 2010, Dieckmann, von Heyden et al. 2010).
Within a minute after closure of the phagosome membrane, the actin coat and the actinbinding proteins present around the phagosome are dissociated.
1.5.2-Phagosome maturation
The phagosome maturation in Dictyostelium can be divided into 2 phases: an early acidic phase and a
late non-acidic phase. The digestion is thought to occur during the acidic phase with the delivery of
lysosomal hydrolases. In Dictyostelium, at the end of the phagosome maturation, the content of the
phagosome is exocytosed. Prior to exocytosis, the phagosome is reneutralised.
1.5.2.1-Phagosome acidification
Almost immediately after closure, the actin coat is dissociated from the phagosome. The newly
formed phagosome is then available for fusion with incoming vesicles.
In mammalian cells, the NADPH oxidase complex, responsible for ROS production, is
assembled at the phagosome membrane during its formation. It is composed of 2 membrane subunits
and 3 cytosolic subunits (Bokoch and Zhao 2006, Bylund, Brown et al. 2010). Dictyostelium possesses
3 isoforms for one of the NOX membrane subunit (NoxA, NoxB, NoxC) and one isoform for the
p22phox membrane subunit (CybA). There is also a homolog of the cytosolic subunit p67phox (NcfA). It
has been recently shown that they are responsible for ROS production in Dictyostelium phagosomes
(Xuezhi Zhang, unpublished data).
Rapidly after closure, the newly formed phagosome fuses with acidic vesicles. The vacuolar
+
H -ATPase is delivered to the phagosome. This allows the acidification of the phagosome lumen
within a minute after its closure (Clarke and Maddera 2006). Concomitantly, Rab7 and proteins
involved in vesicle fusion like the SNARE components Vti1, syntaxin 7 and syntaxin 8 and the
lysosomal marker LmpB are also delivered (Gotthardt, Warnatz et al. 2002, Gotthardt, Blancheteau et
al. 2006). However, Rab7 and the vATPase are in different classes of vesicles and Rab7 is not
involved in vATPase delivery. Indeed, it has been shown that cells expressing a dominant negative
form of Rab7 are delayed for the lysosomal glycoprotein LmpA delivery but not for the vATPase
delivery (Rupper, Grove et al. 2001). Using FITC coated-beads, it has been shown that the
phagosomal pH can reach 4.5 at 30 minutes after phagosome formation (Gopaldass, Patel et al. 2012).
24 However, this is the lowest value measurable with FITC. In an other study, using Oregon green
coupled-dextran, an endosomal pH of 3 was measured (Marchetti, Lelong et al. 2009). Oregon green
has a pKa of 4.7 whereas it is 6.5 for FITC, thus allowing the measurement of lower pH values. So it
would not be surprising that the phagosomal pH reaches even more acidic value than the 4.5 measured
with FITC.
The protein Nramp1 (Natural Resistance Associated Membrane Protein 1) is also found in
early phagosomes. Dictyostelium encodes 2 Nramp proteins (Nramp1 and 2). Only Nramp1 is found
on phagosomes. Nramp2 is present on the contractile vacuole. They are metal transporters. Nramp1
confers resistance to different pathogenic bacteria by depleting the lumen of the phagosome in Fe2+. Its
activity is dependent on the v-ATPase. The Fe2+ export is favored by the H+ gradient established by
the v-ATPase (Peracino, Wagner et al. 2006, Peracino, Buracco et al. 2013).
1.5.2.2-Lysosomal enzymes trafficking
Following acidification, lysosomal enzymes are delivered to the phagosome and the phagosome
becomes a phagolysosome. An acidic pH is actually a prerequisite for an optimal digestive activity of
lysosomal enzymes. However, different classes of lysosomal enzymes exist. They are classified
according to the modification they bear. Those different classes are segregated into different
compartments and are delivered to the phagosome in a sequential manner. The first lysosomal
enzymes delivered to the phagosome are the cysteine proteases such as CprG (cysteine protease 34).
They carry a N-acetylglucosamine 1-phosphate (GlcNAC-1P) modification and are delivered to the
phagosome immediately after acidification (Souza, Mehta et al. 1997, Gotthardt, Warnatz et al. 2002).
Then, lysosomal enzymes bearing a Man-6-SO4 modification like Cathepsin D (CatD) or Man-6-PO4
modification, like α-mannosidase and β-glucosidase, are also delivered to the phagosome 15 minutes
after phagosome formation (Journet, Chapel et al. 1999, Gotthardt, Warnatz et al. 2002). However,
these different lysosomal enzymes appear not to temporally overlap in bacteria-containing
phagosomes (Souza, Mehta et al. 1997). Different waves of recycling likely allow the retrieval of the
first delivered enzymes before the addition of a new set of enzymes. Concomitantly with the addition
of lysosomal enzymes, the lysosomal marker LmpA accumulates at the phagosome (Gotthardt,
Warnatz et al. 2002).
1.5.2.3-Reneutralisation phase and exocytosis
Thirty minutes after phagosome formation, the H+-vATPase is retrieved from the phagosome allowing
its reneutralisation (Clarke, Kohler et al. 2002, Carnell, Zech et al. 2011). The WASH (Wasp And
Scar Homologue) complex, which is an actin nucleating promoter, is directly involved in the H+vATPase retrieval (Carnell, Zech et al. 2011). This complex will be further described in the following
section. The phagosome reaches a pH of 6 fifty to sixty minutes after closure. The phagosome is then
called a post-lysosome. The Scar (Seastone, Harris et al. 2001) and Arp2/3 (Insall, MullerTaubenberger et al. 2001) proteins are recruited to the phagosome to induce the formation of an actin
25 coat. The actin-binding protein Coronin also accumulates on the phagosome at thirty to forty minutes
post phagosome closure. It is progressively replaced by Vacuolin A and B sixty to ninety minutes after
particle ingestion (Rauchenberger, Hacker et al. 1997). In the meantime, once the phagosome is
reneutralised, it undergoes homotypic fusion to form a big post-lysosome compartment. This
homotypic fusion process is dependant of the PI 3-kinase and of PKB (Rupper, Lee et al. 2001).
Rab14 is also a regulator of this process. Indeed, cells expressing constitutively active Rab14 show an
increased rate of phagosome fusion, whereas cells expressing the dominant-negative form of Rab14
show decreased phagosome fusion (Harris and Cardelli 2002). The LvsB protein also accumulates on
large post-lysosome. LvsB is similar to LYST/Beige, the mutated gene in the Chediak-Higashi
Syndrome. It has been shown that this protein is a negative regulator of heterotypic fusion. Indeed, its
deletion leads to the formation of acidified post-lysosomes probably resulting from inappropriate
fusion with early endosomes (Cornillon, Dubois et al. 2002, Kypri, Schmauch et al. 2007). Then, the
post-lysosome releases its content by exocytose. It releases undigested remains and some lysosomal
enzymes bearing Man-6PO4 modification. This event takes only few seconds (Clarke, Kohler et al.
2002, Neuhaus, Almers et al. 2002). Then, a patch of actin and vacuolin remains at the site of
exocytosis and is finally dissociated from the membrane.
1.5.3-Membrane trafficking: the fusion and fission machineries
1.5.3.1-Fusion machinery
Phagosomal and endosomal membrane fusion steps require tethering complexes. Their role is to bring
the membranes of the compartments to fuse in close proximity. From studies done mainly in yeast but
also in mammals, two major tethering complexes have been found in the phago-endosomal pathway:
CORVET (Class C core vacuole/endosome/tethering) and HOPS (homotypic fusion and protein
sorting). These tethering complexes are recruited to endosomal membranes through binding to Rab5
or Rab7 GTPases. Both complexes are composed of four class C subunits called the Class C core
complex: Vps11, Vps 18, Vps 16 and Vps33. Vps33 binds to SNARE proteins (Subramanian,
Woolford et al. 2004). Two additional subunits differ between the two complexes and selectively bind
to Rab5 or Rab7. The HOPS complex has been more intensely studied. However, both CORVET and
HOPS are supposed to function similarly. A working model of their action is described in figure 7.
The additional subunits bind to Rab proteins on both compartments. Then, the Vps33 subunit of the
Class C core complex binds to SNARE proteins on both compartments. This achieves tight tethering
of the two compartments and is supposed to promote SNAREs assembly and fusion (Nickerson, Brett
et al. 2009, Balderhaar and Ungermann 2013).
26 Figure 7: Function of the HOPS complex (Balderhaar and Ungermann 2013). The HOPS complex binds Rab7
present on the membrane of the two compartments to fuse together. Then, the Vps33 subunit of the complex
binds the SNARE proteins present on both compartments to favor their assembly and achieve tethering.
1.5.3.1.1-CORVET
CORVET selectively binds to Rab5. Rab5 is present on early endosomes and to a lesser extent on late
endosomes. CORVET then promotes homotypic fusions between early endosomes, as well as fusions
between early and late endosomes. It is composed of the Class C core complex and the two subunits
Vps3 and Vps8.
1.5.3.1.2-HOPS
HOPS selectively binds to Rab7. Rab7 is present on late endosomes and lysosomes. HOPS then
promotes homotypic fusions between late endosomes, and fusions between late endosomes and
lysosomes. HOPS is composed of the Class C core complex and the two subunits Vps39 and Vps41.
Vps39 acts as a GEF to activate Rab7 (Wurmser, Sato et al. 2000).
1.5.3.2-Fission machinery
Together with fusion steps, phagosomes and endosomes maturation also relies on fission events. This
allows the retrieval and recycling of “escaped” and unnecessary material as well as of factors that have
already performed their function and need to be returned to their compartment of origin.
1.5.3.2.1-The retromer
The retromer complex was first discovered in yeast. It is involved in retrograde transport of
transmembrane proteins from endosomes to the Trans-Golgi Network (TGN) and is notably
responsible for the sorting of the mannose-6-phosphate receptor. It is composed of a trimer of Vps26Vps29-Vps35 and a dimer of sorting nexins. In yeast, these two sorting nexins are Vps5 and Vps17
(Bonifacino and Hurley 2008, Seaman, Gautreau et al. 2013). However, in mammals, there are two
orthologues of Vps5, SNX1 and SNX2, which dimerise with a Vps17 orthologue, SNX5 or SNX6
(Wassmer, Attar et al. 2007). The SNX proteins contain a BAR-domain and a Phox homology (PH)
27 domain. The BAR domain can induce curvature of membranes and are responsible for the formation
of endosomal membrane tubules into which cargo proteins are sorted. The PH domain allows the
binding of the SNX proteins to PI3P present in endosomal membranes. Via binding to membranes,
SNX1 and SNX2 proteins recruit the trimer Vps26-Vps29-Vps35 (Rojas, Kametaka et al. 2007). The
trimer is also called “cargo recognition complex”. Indeed, the subunit Vps35 binds to cargos. It has
been shown to recognize the cytoplasmic tail of the mannose-6-phosphate receptor (Arighi, Hartnell et
al. 2004). Vps35 is the central subunit of the “cargo recognition complex”, and binds both to Vps26
and Vps29. Two other proteins might be required for the recruitment of the cargo-selective trimer.
Both Rab7a and SNX3 are involved in this process (Seaman, Harbour et al. 2009, Harterink, Port et al.
2011, Vardarajan, Bruesegem et al. 2012).
The retromer has been shown to interact with several proteins. SNX5 and SNX6 bind to the
dynein complex (Wassmer, Attar et al. 2009). By linking the retromer to microtules, this probably
allows the elongation of nascent SNX-generated tubules along microtubules toward the Golgi. The
retromer also interacts with EHD proteins (Gokool, Tattersall et al. 2007). These proteins are thought
to help stabilize the formed tubules. EHD proteins are required for the recycling of different cargo
proteins such as the transferrin receptor or the MHC class I (Caplan, Naslavsky et al. 2002, Rapaport,
Auerbach et al. 2006). Several interactions have been described between the retromer complex and the
WASH complex and will be described in the following section.
The subunits Vps26-Vps29-Vps35 of the retromer are also found in Dictyostelium, as well as
the sorting nexin Vps5.
28 Figure 8: The retromer complex and its interacting partners (Seaman 2012). The retromer complex is recruited
to endosomes. By interacting with the dynein complex, it is involved in the retrograde transport of the mannose6-phosphate receptor. The retromer complex can also recruit the WASH complex to endosomes to allow the
recycling of the transferrin receptor to the plasma membrane.
1.5.3.2.2-The WASH complex
The WASH complex is involved in different sorting pathways from endosomes. In mammals, it is
notably required for the retrograde transport of cargo proteins, such as the mannose-6-phosphate
receptor, from endosomes to the Golgi (Gomez and Billadeau 2009). It is also involved in the
recycling of proteins, such as the transferrin receptor, from endosomes to the plasma membrane
(Derivery, Sousa et al. 2009). The WASH complex is an actin nucleation-promoting factor recruited to
endosomes, where it induces the formation of actin filaments. The actin is hypothesised to help
maintain specific microdomains in the endosomal membrane into which sorting proteins would be
directed (Puthenveedu, Lauffer et al. 2010). This complex is composed of five subunits: KIAA1033
(or SWIP), Strumpellin, Fam21, CCDC53 and Wash1 (Wiskott-Aldrich syndrome homologue 1)
(Seaman, Gautreau et al. 2013). It is recruited to endosomes via interaction with the retromer complex
(Harbour, Breusegem et al. 2010). Indeed, it has been shown that the Vps35 subunit of the retromer
can bind to Fam21 and Wash1. However, only the Vps35-Fam21 interaction is necessary for the
recruitment of WASH. An interaction between SNX1 and SNX2 of the retromer complex and the
WASH complex has also been reported (Gomez and Billadeau 2009, Harbour, Breusegem et al. 2012).
The WASH complex is also found in Dictyostelium where it is required for the retrieval of the
+
H -vATPase from post-lysosomes (Carnell, Zech et al. 2011). The deletion of Wash1 leads to a
29 phagosome reneutralisation defect. Furthermore, in Dictyostelium, Fam21 is likely dispensable for the
recruitment of WASH to endosomes. Instead, it would recycle the WASH complex from postlysosomes to compartments earlier in the phago-lysosomal pathway (Park, Thomason et al. 2013).
1.6-Intracellular pathogens
Pathogens have evolved strategies to avoid their degradation by the innate immune system and by
professional phagocytes in general. The simplest one is to avoid ingestion by these cells. To prevent
their recognition by receptors, some bacteria express a polysaccharide capsule around their cell wall,
such as Neisseria meningitidis, P. aeruginosa or Streptococcus spp. Some others such as S.
typhimurium, Helicobacter pylori or Yersinia spp chemically modify their LPS. Some bacteria can
alter the signaling cascade normally activated by the binding to a receptor. For example, both Yersinia
and S. typhimurium can inject effectors into the host cell via a type III secretion system. Those
effectors target host proteins involved in the signaling cascade (Sarantis and Grinstein 2012).
However, an other interesting category of pathogens has established complex strategies in
order to infect innate immune cells and to replicate intracellularly or even intraphagosomally. These
intracellular bacteria usually interfere with the phagosomal maturation pathway. For example, L.
pneumophila, the causative agent of “legionnaires’ disease”, is taken up by phagocytosis. Those
bacteria possess a Type IV secretion system, which injects numerous effectors in the cytosol of the
infected cell. By binding to host proteins involved in the endocytic maturation pathway, those
effectors allow the modification of the maturation of the Legionella-containing-phagosome
(Flannagan, Cosio et al. 2009, Bozzaro and Eichinger 2011). Listeria monocytogenes is able to evade
from its containing phagosome into the cytosol by secreting a pore-forming toxin called LLO
(Listeriolysin O) (Flannagan, Cosio et al. 2009).
Mycobacteria are also intracellular pathogens. Their pathogenicity will be further described in
the following sections.
1.6.1-Mycobacterium tuberculosis
M. tuberculosis is the causative agent of tuberculosis. The latest “global tuberculosis report” in 2012
from the WHO reported 8.7 million new tuberculosis cases in 2011 and 1.4 million deaths caused by
this infectious disease. It is estimated that one third of the world population is infected. However, only
5-10% of the infected individuals develop active tuberculosis. Immuno-compromising conditions,
such as AIDS, poor nutrition, old age and stress trigger the activation of the disease. Tuberculosis is
actually the most common cause of death of HIV infected people. Furthermore, active cases are
disproportionally distributed in the world, in regions which correlate with reduced socio-economic
status. Only 22 countries, mainly in sub-sahara Africa and Asia, bear 80% of the world total number
of active tuberculosis (Russell 2007, Russell 2011) (Global tuberculosis report, 2012).
30 1.6.1.1-Pathogenesis
M. tuberculosis is transmitted from human to human by coughing and inhalation of the contaminated
aerosols. Then, the bacterium enters the new host via the lungs, where it is phagocytosed by alveolar
macrophages. By producing TNF-α and cytokines, this infected macrophage induces the recruitment
of various immune cells: neutrophils, natural killer, CD4+ T cells and CD8+ T cells. However, those
cells are recruited in successive waves. This leads to the formation of a stratified immune cells
structure around the infected macrophage, called granuloma. In its mature form, the granuloma
develops a fibrous cuff and is vascularized. Mycobacteria proliferate inside the macrophages of the
granuloma.
The granuloma usually contains the infection. But, when the immune status of the host
changes, the granuloma further evolved to a late stage. It looses its vascularization leading to hypoxia
and caseation. The granuloma then looses its structure. It ruptures and releases large amount of
infectious mycobacteria (Russell 2007, Ramakrishnan 2012).
Figure 9: M. tuberculosis pathology (Russell 2007). In the lungs, alveolar macrophages get infected by ingesting
aerosols contaminated with M. tuberculosis. The infected macrophage induces the aggregation of immune cells
to form a granuloma in which mycobacteria proliferate. The caseation of the granuloma leads to its rupture and
the release of numerous mycobacteria in the airways.
31 1.6.1.2-Manipulation of the macrophage phagosomal pathway
M. tuberculosis avoids its destruction by macrophages by manipulating the maturation of its
containing-phagosome. Indeed, it has been shown that a mycobacteria-containing compartment is
arrested in an early phagosomal stage. First, it retains the actin-binding protein Coronin, which is
normally dissociated within a minute after phagosome formation (Ferrari, Langen et al. 1999, Fratti,
Vergne et al. 2000, Deghmane, Soualhine et al. 2007). It also retains the early endosomal marker Rab5
(Via 1997, Clemens, Lee et al. 2000, Fratti, Backer et al. 2001, Kelley and Schorey 2003, Rohde,
Yates et al. 2007). However, Rab5 effectors PI3K and EEA1 (Early Endosomal Antigen 1) are
excluded from the mycobacteria-containing compartment (Fratti, Backer et al. 2001, Chua and Deretic
2004, Hestvik, Hmama et al. 2005). As these effectors are necessary for the phagosome maturation,
their absence blocks further maturation. The mycobacteria-containing compartment is still accessible
to the early recycling pathway. Indeed, the transferrin receptor and the GM1 ganglioside are still
trafficking through the mycobacteria-containing compartment (Clemens and Horwitz 1996, Russell,
Dant et al. 1996, Sturgill-Koszycki, Schaible et al. 1996, Rohde, Yates et al. 2007). Furthermore, the
mycobacteria-containing compartment fails to acidify (Sturgill-Koszycki, Schlesinger et al. 1994,
Mwandumba, Russell et al. 2004, Pethe, Swenson et al. 2004). It also fails to accumulate lysosomal
markers like the lysosomal enzyme Cathepsin D (Sturgill-Koszycki, Schaible et al. 1996, Malik, Iyer
et al. 2001) or the mannose 6-phosphate receptor (Xu, Cooper et al. 1994). It has also been shown that
M. tuberculosis is able to prevent the increase of intracellular calcium concentration in the infected
macrophage (Malik, Iyer et al. 2001, Vergne, Chua et al. 2003). Calcium concentration is normally
increased during uptake of inert particle and is involved in phagosome-lysosome fusion (Colombo,
Beron et al. 1997, Peters and Mayer 1998, Vergne, Chua et al. 2003). Indeed, Ca2+ activates the
Ca2+/calmodulin dependent PI3K, which is involved in the recruitment of EEA1 to the phagosome. By
preventing the calcium concentration increase, M. tuberculosis again blocks phagosome maturation
(Vergne, Chua et al. 2003).
1.6.1.3-Virulence factors
To establish a successful infection, M. tuberculosis possesses a battery of virulence factors. Some are
lipids and are shed from the lipidic cell wall of the bacteria, and others are proteins. They can be
involved in different steps of the infection: the manipulation of the phagosome maturation, the
intracellular growth or the escape from their containing-compartment and the dissemination to other
cells.
1.6.1.3.1-Virulence factors involved in macrophage manipulation
M. tuberculosis cell wall lipids can block phagosome and lysosome fusion. ManLAM (MannoseLipoarabinomannan) is the major lipoglycan of M. tuberculosis cell wall. ManLAM coated-beads are
able to decrease the calcium flux usually induced during phagocytosis. This leads to a decreased
activation of the Ca2+/Calmodulin dependent PI3K and prevents the recruitment of EEA1 to the M.
32 tuberculosis-containing compartment (Fratti, Chua et al. 2003, Vergne, Chua et al. 2003). Latex-beads
coated with an other cell wall lipid, the trehalose dimycolate (TDM), are also able to delay phagosome
maturation (Indrigo 2003, Axelrod, Oschkinat et al. 2008). M. tuberculosis strain lacking fbpA is
defective in transferring mycolic acids to trehalose to form TDM. This strain lacking TDM allows a
partial phagosome maturation (Katti, Dai et al. 2008). By screening a transposon mutant library of M.
marinum, it has been shown that phenolic glycolipid phenolphtiocerol diester (PGL-1), an other
mycobacterial cell wall lipid, is also involved in the phagosome maturation arrest (Robinson, Wolke et
al. 2007, Robinson, Kolter et al. 2008). A similar screen performed on a M. tuberculosis mutant
library allowed the isolation of mutant defective for phagosome maturation arrest. Several mutants
have transposon insertion in a same operon, which encodes for enzymes involved in isoprenol
compounds synthesis (Pethe, Swenson et al. 2004). One of these enzymes synthesizes a cell wall lipid,
the isoprenoid edaxadiene. Latex-beads coated with edaxadiene block phagosome maturation (Mann,
Prisic et al. 2009, Mann, Xu et al. 2009).
Secreted proteins are also able to manipulate the phagosome maturation. However, these
proteins lack signal sequences and it is still unclear how they can exit the mycobacteria-containing
compartment and access the cytosol.
The secreted acid phosphatase M (SapM) dephosphorylates the PI3P present in the
phagosomal membrane. This prevents the binding of PI3P binding proteins like EEA1, which is
necessary for the phagosome maturation (Vergne, Chua et al. 2005). The Serine/Threonine kinase
PknG has also been shown to arrest phagosome maturation. Indeed, its expression prevents the
targeting of the non-pathogenic strain M. smegmatis to lysosomes. However, its mechanism of action
is still unknown (Cowley, Ko et al. 2004, Walburger, Koul et al. 2004). The lipoamide dehydrogenase
(LpdC) is able to interact with coronin in a cholesterol dependant manner. This retains coronin around
the mycobacteria-containing compartment and prevents its maturation (Deghmane, Soualhine et al.
2007). PtpA is a low-molecular-weight tyrosine phosphatase also arresting the phagosome maturation.
Its substrate is Vps33b, a protein involved in HOPS complex assembly to allow vesicles fusion.
Vps33b also binds to the H+-vATPase. It has been shown that PtpA binds to the subunit H of the H+vATPase to destabilize the binding between Vps33B and the H+-vATPase. Then, it dephosphorylates
Vps33b to dissociate the HOPS complex and prevent the H+-vATPase delivery to the mycobacteriaconaining compartment (Bach, Papavinasasundaram et al. 2008, Wong, Bach et al. 2011).
PE_PGRS30 is the homolog of the M. marinum Mag24 gene. It has been shown that Mag24 is
upregulated in the macrophage immediately after phagocytosis of the bacteria. M. marinum strain with
a transposon insertion in the Mag24 gene (L1D) is not able to stop phagosome maturation. Notably, it
cannot prevent the delivery of the H+-vATPase (Ramakrishnan 2000, Hagedorn and Soldati 2007,
Hagedorn, Rohde et al. 2009). Recently, the role of PE_PGRS30 in macrophage manipulation has
been confirmed using M. tuberculosis strain lacking the gene encoding PE_PGRS30 (Iantomasi, Sali
et al. 2012).
33 1.6.1.3.2-Virulence factors involved in intracellular growth
M. tuberculosis encodes a PhoP protein. It has been shown that the PhoP homolog in Salmonella is the
major pH sensor (Martin-Orozco, Touret et al. 2006). M. tuberculosis strain lacking PhoP is not able
to multiply intracellularly (Perez, Samper et al. 2001). PhoP is notably regulating the transcription of
proteins involved in cell wall complex lipids biosynthesis (Gonzalo Asensio, Maia et al. 2006,
Walters, Dubnau et al. 2006). The PhoP mutant is actually more attenuated than the BCG (Bacillus
Calmette-Guerin) used for vaccination. It has been shown that it can be used as a vaccine to confer
protective immunity in guinea pigs and mice (Aguilar et al., 2006; Martin et al., 2006).
It has been shown that the lipid metabolism is particularly important for the intraphagosomal
life of M. tuberculosis. Indeed, inside phagosomes, M. tuberculosis uses fatty acids as primary carbon
source. In this condition, the carbon flux is diverted from the Kreb’s cycle into the glyoxylate cycle.
The two isocitrate lyases (ICL1 and ICL2) encoded by M. tuberculosis allow the diversion of
isocitrate into the glyoxylate cycle to produce pyruvate (McKinney, Honer zu Bentrup et al. 2000,
Munoz-Elias and McKinney 2005). However, the metabolism of cholesterol leads to the formation of
propionyl-CoA, which is toxic for M. tuberculosis if is not metabolized. The 2-methyltcitrate cycle
allows its metabolism by condensing it with oxaloacetate to produce pyruvate. In this cycle, the ICL
enzymes operate as 2-methylisocitrate lyases (Gould, van de Langemheen et al. 2006). These ICL
enzymes play two important functions for intracellular growth of the bacteria. It is not surprising that
M. tuberculosis strain lacking these two ICL is not able to grow intracellularly (Munoz-Elias and
McKinney 2005).
Figure 10: ICL1/ICL2 in the glyoxylate cycle and the methylcitrate cycle in M. tuberculosis (Munoz-Elias and
McKinney 2005). During infection, the M. tuberculosis ICL enzymes divert the isocitrate from the TCA cycle
into the glyoxylate cycle. They also act as 2-methylisocitrate lyases in the methylcitrate cycle to detoxify the
propionyl-CoA produced by the metabolism of fatty acids.
34 1.6.1.3.3-Virulence
factors
involved
in
phagosome
escape
and
dissemination
M. tuberculosis possesses five type VII secretion systems: ESX1-ESX5 (Abdallah, Gey van Pittius et
al. 2007, Stoop, Bitter et al. 2012). Among them, the ESX-1 secretion system is the most studied. The
different components of ESX-1 are encoded in the RD1 (region of difference 1) locus. This locus,
absent from the vaccine strain BCG, has been shown to be important for M. tuberculosis
pathogenicity. Indeed, its deletion from M. tuberculosis leads to attenuated virulence, whereas its
introduction in the BCG strain enhances the virulence (Pym, Brodin et al. 2002, Lewis, Liao et al.
2003). ESX1 notably secretes two small proteins also encoded in the RD1 locus: ESAT-6 (6 kDa
Early Secreted Antigenic Target) and CFP-10 (10 kDa Culture Filtrate Protein) (Pym, Brodin et al.
2003, Gao, Guo et al. 2004, Guinn, Hickey et al. 2004). ESAT-6 and CFP-10 form a dimer (Brodin, de
Jonge et al. 2005, Renshaw, Lightbody et al. 2005). The C-terminal region of CFP-10 is crucial to
allow the secretion of the ESAT-6/CFP-10 complex (Champion, Stanley et al. 2006). It has been
shown that ESAT-6 has a pore-forming activity (de Jonge, Pehau-Arnaudet et al. 2007, Smith,
Manoranjan et al. 2008) and is involved in the escape of M. tuberculosis from its phagosome into the
cytosol of the infected cell (van der Wel, Hava et al. 2007, Hagedorn, Rohde et al. 2009, Houben,
Demangel et al. 2012). This phagosome escape is necessary to then allow the spreading of the
infection. Indeed, deletions of RD1 or of ESAT-6 lead to a decreased spreading of M. tuberculosis to
uninfected phagocytes (Guinn, Hickey et al. 2004, Hagedorn, Rohde et al. 2009) and reduced tissue
invasiveness (Hsu et al., 2003).
1.6.2-Mycobacterium marinum
M. marinum is a close cousin of M. tuberculosis. It is a natural pathogen of fish, frogs and coldblooded animals. The mechanisms of virulence are well conserved between M. marinum and M.
tuberculosis. Indeed, M. marinum also infects macrophages and induces the formation of granuloma.
As M. tuberculosis, it possesses a homolog of the PE_PGRS30 cell surface protein, Map24-1 and the
RD1 locus. However, M. marinum grows faster than M. tuberculosis and has an optimal growth
temperature of 32ºC. This low growth temperature makes of M. marinum a safer mycobacteria to work
with. Indeed, it can hardly cause severe infection in humans. However, it can still induce the formation
of superficial skin lesions on the extremities. These characteristics make M. marinum a good model to
mimic and study M. tuberculosis infections.
M. marinum infection has been studied in zebrafish and flies (Dionne, Ghori et al. 2003, Pozos
and Ramakrishnan 2004) (Pozos and Ramakrishnan, 2004; Dionne et al., 2003). Ten years ago, it has
been shown that the amoeba Dictyostelium can also be infected with M. marinum (Solomon, Leung et
al. 2003).
35 1.6.2.1-Infection of Dictyostelium
A previous study described three phases in the infection course of Dictyostelium with M. marinum
(Hagedorn and Soldati 2007). During the first twelve hours, the bacteria do not replicate. This phase
has been called the manipulation phase. Indeed, after ingestion by Dictyostelium, M. marinum resides
in a phagosome. As M. tuberculosis, it has to manipulate its containing compartment in order to
prevent further maturation and avoid its killing. During this phase, the M. marinum-containing
compartment transiently acquires the H+-vATPase. However, it is rapidly retrieved from the
compartment and then, the M. marinum-containing-compartment does not acquire the lysosomal
enzyme Cathepsin D and slowly acquires the late phagosomal markers p80 and vacuolin. The second
phase of the infection is a proliferation phase. During this phase, M. marinum strongly proliferates
inside its manipulated compartment. In the meantime, the M. marinum-containing compartment
become spacious and strongly accumulates the late phagosomal markers p80 and vacuolin. Finally, the
last phase is a phase of bacteria release. During this phase, M. marinum is able to break the membrane
of its containing-compartment. As for M. tuberculosis, this process is dependent of the secretion of the
proteins ESAT6 and CFP10 encoded in the RD1 locus of M. marinum. Once in the cytosol, the
bacteria can escape the cell using a nonlytic ejection system called ejectosome and disseminate to
other Dictyostelium cells (Hagedorn, Rohde et al. 2009).
36 1.7-Aim of the thesis
Like M. tuberculosis, M. marinum diverts the maturation of its containing-phagosome to the
establishment of a compartment allowing the replication of the bacteria. This compartment is known
to be depleted for H+-vATPase, late phagosomal markers and lysosomal enzymes. A number of
studies have identified a few host proteins targeted by mycobacteria and involved in phagosome
maturation arrest. However, this list of targeted proteins is far from being complete and the events
happening during the first hours of infection are still poorly understood. The aim of this thesis is to
understand how M. marinum manipulates the phagosomal pathway. To achieve this aim, we will
dissect the temporal modification of the composition of the M. marinum-containing compartment in
the professional phagocyte Dictyostelium, during the first hours of infection to understand how this
compartment is diverted from the normal phagosomal pathway. My main experimental strategy will
be to establish a method to purify the bacterium-containing compartment and exploit it to obtain an
exhaustive, quantitative and comparative, time-resolved proteomic composition as a function of the
contained virulent or attenuated bacterium. We also plan to identify and characterise host pathways
targeted by M. marinum to establish a successful infection.
A previous study in the lab showed that the H+-vATPase is transiently delivered to the early
M. marinum-containing compartment. However, it was demonstrated that the M. tuberculosiscontaining compartment fails to significantly acidify. We will further characterise the pH changes in
the M. marinum-containing compartment. In Dictyostelium, the WASH complex was shown to be
responsible for the H+-vATPase retrieval from phagosomes and consequently for the reneutralisation
of these compartments. The potential role of this complex will be further investigated in the context of
mycobacterial infections in Dictyostelium.
37 38 2-Material and methods
39 2.1-Material
2.1.1-Media
HL5c medium
5 g/L Yeast extract
(Formedium, UK)
5 g/L Proteose tryptone
5 g/L Proteose peptone
pH 6.2
1.2 g/L (8.8 mM) KH2PO4
0.35 g/L (2.5 mM) Na2HPO4
10 g/L (56 mM) Glucose
Low Fluorescence medium
11 g/L Glucose
(LoFlow, Formedium, UK)
0.68 g/L KH2PO4
5 g/L casein peptone
26.8 mg/L NH4Cl
37.1 mg/L MgCl2
1.1 mg/L CaCl2
8.11 mg/L FeCl3
4.84 mg/L Na2-EDTA
2.30 mg/L ZnSO4
1.11 mg/L H3BO3
0.51 mg/L MnCl2.4H2O
0.17 mg/L CoCl2
0.15 mg/L CuSO5.5H2O
0.1 mg/L (NH4)6Mo7O24.4H2O
Middlebrook 7H9
4.7 g/ 900 mL
2% glycerol
0.05% Tween 80
After autoclaving and cooling to 50ºC, 10% OADC
2.1.2-Buffers and solutions
Phosphate Buffer Saline (PBS)
140 mM NaCl
pH 7.4
3.4 mM KCl
1.8 mM KH2PO4
10 mM Na2HPO4
Tris Buffer Saline (TBS)
10 mM Tris
pH 7.4
200 mM NaCl
Sorensen Buffer
15 mM KH2PO4
pH 6
2 mM Na2HPO4
40 Sorensen-Sorbitol Buffer
15 mM KH2PO4
2 mM Na2HPO4
120 mM sorbitol
SDS PAGE sample buffer (2X):
125 mM Tris pH 6.8
4% SDS
20% glycerol
0.02% Bromophenol Blue
10% β-mercaptoethanol (add freshly)
SDS PAGE running buffer
0.1% SDS
pH 8.0
25 mM Tris
192 mM Glycine
Towbin transfer buffer
25 mM Tris
pH 8.6
192 mM Glycine
20% Methanol
0.02% SDS
2.1.3-Antibodies
Primary antibodies
Antigen
Antibody type
Method, dilution
Gift from, Reference
p80
M mAb, H161
WB, IF 1/10
Dr. Pierre Cosson (Ravanel, de Chassey et
al. 2001)
Vacuolin
M mAb, 221-1-1
WB, IF 1/10
Dr. M. Maniak (Rauchenberger, Hacker et
al. 1997)
VatA
M mAb 221-35-2
WB, IF 1/10
Dr. M. Maniak (Jenne, Rauchenberger et
al. 1998)
GFP
R pAb
WB, IF 1/1000
p80-Cy5
M mAb H161
IF 1/500
MBL
labelled
Ab, antibody; M, mouse; R, rabbit; mAb, monoclonal antibody; pAb, polyclonal antibody; WB,
western blot; IF, immunofluorescence
Secondary antibodies
Antibody
Method, Dilution
Supplier
Goat anti-mouse/rabbit HRP
WB 1/5000-1/10000
Biorad
IF 1/500-1/1000
Molecular probes
coupled
Goat anti-mouse/rabbit Ig
Alexa 488, 594, 633
41 2.1.4-Antibiotics
Antibiotic
Stock concentration
Working concentration
Penicilin
10000 U/mL
10 U/mL
Streptomycin
10 mg/mL
100 µg/mL
Kanamycin
100 mg/mL
50 µg/mL
Hygromycin
100 mg/mL
50 µg/mL
Apramycin
50 mg/mL
50 µg/mL
G418
10 mg/mL
10 µg/mL
2.1.5-Kits
The Tandem Mass Tag (TMT) kit is from Thermo Scientific (TMT sixplex Label Reagent Set, 5 x 0.8
mg cout., no. art 90066).
2.1.6-D. discoideum cell lines
Cell line
Received from
Reference
AX2
Dr. G. Gerisch
AX2ΔWash
Dr. R. Insall
(Carnell, Zech et al. 2011)
AX2ΔFam21
Dr. R. Insall
(Park, Thomason et al. 2013)
AX2ΔWash- GFP-Wash
Dr. R. Insall
(Carnell, Zech et al. 2011)
2.1.7-Mycobacteria strains
Strain
Received from
Reference
M. marinum
Dr. L. Ramakrishnan
(Cosma, Sherman et al. 2003)
M. marinum-L1D
Dr. L. Ramakrishnan
M. marinum GFP
TS lab
(Hagedorn and Soldati 2007)
M. smegmatis GFP
TS lab
(Hagedorn and Soldati 2007)
M. marinum-L1D GFP
Dr. L. Ramakrishnan
(Ramakrishnan 2000)
M. marinum ΔRD1 GFP
Dr. L. Ramakrishnan
M. marinum mCherry10
TS lab
M. smegmatis DsRed
TS lab
M. marinum LuxABCDE
TS lab
(Arafah, Kicka et al. 2013)
M. marinum ΔRD1 Lux ABCDE
TS lab
(Arafah, Kicka et al. 2013)
42 2.2-Methods
2.2.1-Cell culture
2.2.1.1-D. discoideum cell culture
Cells are cultivated at 22ºC in HL5c (Formedium) supplemented with 100 U/mL Penicillin and 100
µg/mL Streptomicin (Gibco). Under adherent conditions, the cells are passaged every 2 to 3 days
according to confluency. Under shaking conditions, cells are cultivated at densities ranging between
5.104 to 5.106 at 180 rpm.
Cell stocks
Cells from one confluent dish are collected and counted using a Neubauer counting chamber. They are
resuspended at a density of 5.106 cells/mL in HL5c 10% DMSO. Aliquots of 1 mL are prepared and
immediately placed in ice-cold Nalgene MisterFreeze boxes filled with isopropanol. They are then
transferred at -80ºC to be slowly frozen. After 24 h, they are transferred in liquid nitrogen for longterm storage.
Spores
Cells are collected from subconfluent dishes. They are washed in Sorensen Buffer and resuspended at
5.107 cells/mL in Sorensen Buffer. 2 mL of the cell suspension are deposited on starvation agar plates
(Sorensen buffer with 2% bacto-agar) and spread by swirling. The plates are incubated 30 minutes
with an open lid until they are almost dry but still humid. They are then incubated upside down at
22ºC for at least 24h in a humid atmosphere.
The sori are collected in the lid of the plates by repeated tapping of the plate against the lid. They are
vigorously resuspended in Sorensen buffer. After counting, they are washed in Sorensen Buffer and
pelleted (2400 rpm, 5 min). They are then resuspended in Sorensen buffer + 10% glycerol at a density
of 107 spores/mL. 1mL aliquots are prepared and frozen in Nalgene MisterFreeze boxes. After 24h,
they are transferred in liquid nitrogen for long-term storage.
2.2.1.2-Mycobacteria culture
Mycobacteria are cultivated in presence of 5 mm glass beads (Sigma) in 7H9 medium, 0.2% glycerol,
0.05% Tween 80 supplemented with 10% OADC in shaking condition at 32˚C.
Mycobacteria can also be cultivated on 7H11 agar plates. 7H11 is complemented with 5% glycerol
and autoclaved. After cooling to 50ºC, 10% OADC and the appropriate antibiotics are added and the
plates are poured.
43 M. marinum-L1D, M. marinum-GFP, M. marinum ΔRD1-GFP, M. smegmatis-GFP
and M.
smegmatis-DsRed are cultivated in presence of 50 µg/mL kanamycin. M. marinum L1D-GFP is
cultivated in presence of 50 µg/mL apramycin. M. marinum-mCherry10, M. marinum-LuxABCDE
and M. marinum ΔRD1-LuxABCDE are cultivated in presence of 50 µg/mL hygromycin.
Glycerol stocks
500 µL of an overnight culture are mixed with 500 µL of 7H9 + 50% glycerol and deposited in a
cryogenic vial. After flash freezing in liquid nitrogen, it is conserved at -80ºC.
2.2.2-Samples preparation
2.2.2.1-Latex-bead phagosomes isolation
The latex-bead phagosomes isolation is performed as described in (Dieckmann, Gopaldass et al.
2008). The cells are fed with latex beads. At corresponding time points, the cells are homogenised and
the latex-bead containing phagosomes are recovered by flotation on a sucrose gradient.
Materials:
- 1.2 L HL5c medium at room temperature
- 50 mL HL5c medium ice cold
- 100 mL Sorensen/120 mM Sorbitol pH8
- 4 L Sorensen/120 mM Sorbitol
- 200 mL sucrose, 2.5 M
- 100 mL HEPES, 1 M, pH7.2
- 2 L of HESES:
40 mL HEPES, 1 M
200 mL sucrose, 2.5 M
1660 mL sterile ddH2O
- Homogenisation buffer: to 13.5 mL HESES buffer, add 500 µL Complete EDTA-free protease
cocktail inhibitor (Roche, Cat no. 1 873 580) - 1 tablet to be dissolved in 1 mL sterile water as stock
and stored at -800C.
44 - 10 mL ATP, 100 mM (Adenosine 5’ triphosphate, 127531, Roche):
200 µL HEPES, 1 M
4.675 mL sucrose, 2.5 M
5.125 mL sterile water
Adjust pH to 7 with 5 M KOH.
-Sucrose solutions:
HEPES 1 M, pH
7.2
Sucrose 2.5 M
Sterile water
Final volume
10% sucrose
1.0 mL
5.84 mL
43.16 mL
50 mL
25% sucrose
2 mL
29.21 mL
68.79 mL
100 mL
35% sucrose
2 mL
40.90 mL
57.10 mL
100 mL
60% sucrose
1 mL
35.06 mL
13.94 mL
50 mL
71.4% sucrose
0.5 mL
20.86 mL
3.64 mL
25 mL
- 500 mL Membrane Buffer:
10 mL HEPES, 1 M
(20 mM final conc.)
10 mL KCl, 1 M
(20 mM final conc.)
1.25 mL MgCl2, 1M
(2.5 mM final conc.)
2 mL NaCl, 5 M
(20 mM final conc.)
Material preparation:
- Prepare 10 x 500 mL Beckmann centrifuge tubes with 330 mL ice-cold Sorensen/Sorbitol buffer on
an ice/water slush bath. Label centrifuge tubes P1 to P6 for the pulse time points, and C1 to C4 for the
chase time points.
- Prepare 10 x 250 mL conical flasks with 100 mL filtered HL5c media at room temperature.
- Prepare a second ice/water slush bath to chill the ATP and the samples before and after
homogenisation.
Preparation of latex beads:
- Spin down 2 x 2 mL of latex bead suspension with a particle diameter of 0.807 µm (Sigma, Cat LB8) in 2 x 2 mL eppendorf tubes in a microcentrifuge for 5 minutes, maximum speed, at room
temperature. For 109 cells, prepare 0.5 mL beads. For 1.33.109 cells, prepare 0.66 mL beads.
45 - Remove supernatant and add 1.5 mL Sorensen/Sorbitol pH 8.0 to each eppendorf tube to resuspend
beads. (1 mL Sorensen/Sorbitol pH 8.0 for 0.5 mL beads aliquot).
- Spin down again as before to wash beads.
- Repeat the washing step and place the resuspended beads in a sonicator water bath for 5 minutes.
- Place on ice.
Preparation of cells:
Phagocytosis will be studied at 6 time points (use 109 or 1.33.109 cells per time point according to
availability):
P1 – 5 minutes pulse
P2 – 15 minutes pulse
P3 – 15 minutes pulse/15 minutes chase
P4 – 15 minutes pulse/45 minutes chase
P5 – 15 minutes pulse/1 hr 45 minutes chase
P6 – 15 minutes pulse/2 hr 45 minutes chase
- Spin down 8.109 cells for 8 minutes at 2,000 rpm (Rotor JLA10.500; 740xg), at 4ºC.
- Resuspend cells in 50 mL Sorensen/Sorbitol buffer pH 8 and pool into a 50 mL Falcon tube.
- Wash cells by spinning down for 5 minutes, 1,600 rpm (Beckmann Allegra 6R –Rotor GH3.8A–
clinical table top centrifuge) at 4ºC.
- Resuspend cells in 20 mL Sorensen/Sorbitol buffer pH 8.
- Add sonicated latex bead suspension to cells.
- Top up volume of cells/beads suspension to 33 mL with Sorensen/Sorbitol buffer pH 8.
- Pre-incubate cells/beads on ice for 15 minutes to allow binding of beads to the cell surface.
Phagocytosis:
- Starting with P6, pipette 5 mL of the cells/beads suspension in P6 to P3, and then 6 mL in P2 and 7
mL (+ the rest) in P1. Swirl the flasks to mix and place on orbital shaker at 120 rpm, 22ºC.
46 - Label centrifuge tubes P1 to P6 for the pulse time points, and C1 to C4 for the chase time points.
- Stop phagocytosis after appropriate pulse time by plunging cells/beads into the prepared 330 mL icecold Sorensen/Sorbitol buffer.
- Spin cells for 8 min at 2,000 rpm (740g) at 4ºC to remove medium and excess beads.
- Wash cells further by resuspension in 50 mL ice-cold HESES buffer and centrifuge for 4 minutes at
1,600 rpm (Beckmann Allegra 6R), 4ºC and remove supernatant.
- For samples P3 to P6, resuspend cells in 5 mL ice cold HL5c (working from the latest time point, P6
to the earliest, P3) and add into new 250 mL conical flasks containing 100 mL HL5c at room
temperature for the chase time points. Swirl the flasks to mix suspension.
- Place flasks C1 to C4 onto orbital shaker at 120 rpm, 22ºC.
- For P1 and P2, repeat twice the washing step of the cells in 50 mL ice-cold HESES buffer and
centrifuge for 4 minutes at 1600 rpm (Beckmann Allegra 6R), 4ºC.
- Keep pelleted cells on ice/water until required for homogenisation.
- Stop chased cells (C1 to C4) at the appropriate times as before by plunging into 330 mL ice-cold
Sorensen/Sorbitol buffer, wash in HESES buffer twice only and keep the pellets on ice-water until
homogenisation.
Chase Time Points:
C1 is stopped after 15 minutes chase (30 minutes total, including 15 minutes pulse)
C2 is stopped after 45 minutes chase (1 hour total, including 15 minutes pulse)
C3 is stopped after 1 hour 45 minutes chase (2 hours total, including 15 minutes pulse)
C4 is stopped after 2 hours 45 minutes chase (3 hours total, including 15 minutes pulse)
Homogenisation:
- The homogenisation part is performed at 4ºC.
- Resuspend each cell pellet in 2 mL of homogenisation buffer.
- Homogenise cells by passing them eight times (single passages) through a ball homogeniser (HGM,
Germany) with a barrel diameter of 8.000 mm and a ball diameter of 7.990 mm (clearance 10 µm).
The ball homogeniser is placed on a metal bloc on ice.
47 - Between each sample, flush the homogeniser once with 10 mL HESES buffer, in one passage, not
back and forth!
- Keep the samples on ice-water in 15 mL Falcon tubes.
- To each sample add 70 µL of 1 M MgCl2, 700 µL of ATP/sucrose and 3.5 mL of 71.4% sucrose. The
total volume is about 7 mL and the end concentration of sucrose is about 40%.
- Mix samples slowly on wheel for 15 minutes at 4ºC.
Sucrose gradients:
- The sucrose gradients can be prepared one day in advance (in a cold room) in open top polyallomer
centrifuge tubes (Beckmann, Cat no. 326823) for SW28 rotor.
Layer in each tube:
4 mL of 60% sucrose
12 mL of 35% sucrose
12 mL of 25% sucrose
- Using a 1.4x100 (17G) mm needle connected to a 10 mL syringe, add samples to the 35%-60%
sucrose gradient interface.
- Finally overlay with 4 mL of 10% sucrose and balance carefully for ultracentrifugation.
- Balance carefully the tubes with 10% sucrose (less than 5 mg difference, Beckmann dixit)
- Ultracentrifuge samples at 28,000 rpm (SW28-Rotor) for 3.0 hours OR overnight, deceleration
without brake or “to 800 rpm” only, at 4ºC.
- Following centrifugation, collect material (phagosomes) from between the 10%-25% sucrose
interface into 15 mL Falcon tubes. Total volume between 4 and 7 mL.
- Dilute phagosomes to 14 mL with HESES and take 50 µL for light scattering measurements at 600
nm.
Light scattering measurements of phagosomes:
- Take 50 µL of phagosome sample and dilute into 950 µL water.
- Take light scattering measurements at 600 nm using water as background reference.
- Multiply the reading by 1,000 and then divide by 1.5 to determine the protein quantity.
48 Collection and storage of phagosomes:
- Transfer phagosomes to ultraclear centrifuge tubes (344058) for SW28 rotor.
- Add 23 mL Membrane buffer to the phagosomes to make a total volume of 37 mL.
- Balance tubes carefully using Membrane buffer for ultracentrifugation.
- Pellet phagosomes by ultracentrifugation at 28,000 rpm (100,000g), brake on, for 45min., 4ºC.
- Following ultracentrifugation, remove supernatant.
- Phagosomes pellets can be flash frozen in liquid nitrogen. The tubes are covered with Parafilm® and
the pellets are conserved at -80ºC.
- According to light scattering measurement, pellets can also be resuspended in the appropriate volume
of SDS-PAGE sample buffer at a concentration of 1 µ g/µL. After flash freezing in liquid nitrogen,
they are stored at -80ºC.
2.2.2.2-Mycobacteria-containing compartments isolation
This protocol has been established during the thesis. It is adapted from the latex-bead phagosomes
isolation protocol. Its optimisation will be described in the results part.
Materials:
- Borate 0.1 M pH 8.5
- HL5c medium No PS
- 0.2 µm beads (Estapor) : 109 µL for 5.109 bacteria (Beads:bacteria ratio 500:1)
- Sorensen- azide 5 mM
- HEPES, 1 M, pH 7.2
- HESES-azide 5 mM
- Homogenisation buffer: to 6.75 mL HESES-azide 5 mM, add 250 µL EDTA-free protease cocktail
inhibitor (Roche)
49 - ATP 100 mM (Adenosine 5’ triphosphate, 127531, Roche):
200 µL HEPES, 1 M
4.675 mL sucrose, 2.5 M
5.125 mL sterile water
Adjust pH to 7 with KOH
- Sucrose 2.5 M
- Sucrose solutions:
HEPES 1 M
Sucrose 2.5 M
Sterile water
Total volume
10% sucrose
800 µL
4.68 mL
34.52 mL
40 mL
20% sucrose
800 µL
9.2 mL
30 mL
40 mL
30% sucrose
800 µL
14 mL
25.32 mL
40 mL
40% sucrose
800 µL
18.68 mL
20.52 mL
40 mL
60% sucrose
800 µL
28.04 mL
11.16 mL
40 mL
71.4% sucrose
800 µL
33.36 mL
5.84 mL
40 mL
- Membrane buffer (500 mL)
10 mL HEPES, 1 M
(20 mM final conc.)
10 mL KCl, 1 M
(20 mM final conc.)
1.25 mL MgCl2, 1M
(2.5 mM final conc.)
2 mL NaCl, 5 M
(20 mM final conc.)
Preparation of the beads
- Prepare 4 eppendorf tubes with 109 µL of 0.2 µm latex beads (K020, Estapor) and complete with
890 µL of Borate 0.1 M (Use Low Binding Eppendorf tubes): 109 µL for 5.109 bacteria
(Beads:Bacteria ratio 500:1)
- Spin down in a microcentrifuge for 10 minutes, 12,000 rpm
- Resuspend the pelleted beads in 1 mL Borate 0.1 M and spin down again
- Repeat this step
- Resuspend the beads in 500 µL Borate 0.1 M
- Sonicate the resuspended beads 5 minutes in a sonicator bath
50 Preparation of the bacteria
- Spin down 4 x 5.109 bacteria in 50 mL Falcon tubes for 10 minutes at 2,000 rpm
- Resuspend the bacteria in 1 mL of Borate 0.1 M and spin down again (Use Low Binding Eppendorf
tubes)
- Repeat this step twice
- Resuspend the pelleted bacteria in 500 µL of Borate 0.1 M and mix with the sonicated beads
Adsorption of the beads on the bacteria
- Incubate the beads and the bacteria on a wheel during 2 hours
- Pellet the beads and bacteria 10 minutes at 12,000 rpm
- Resuspend the pellet in 1 mL of HL5c without PS and spin down again
- Resuspend the pellet in 1 mL HL5c without PS
Isolation of the bacteria+beads complexes
- Prepare a sucrose gradient in 4 small centrifuge tubes (thin wall, 11x34mm) for TLS55 rotor:
300
µL 60% sucrose
1 mL 20% sucrose
- Add the resuspended beads and bacteria at the top
- Ultracentrifuge 30 minutes at 55,000 rpm
- Recover the 20-60% interphases in Low binding Eppendorf tubes and add 500 µL HL5c without PS
- Centrifuge 10 minutes at 12,000 rpm
- Resuspend the pellets in 1 mL of HL5c without PS
Preparation of the host cells
- At least 30 minutes before infection, prepare 8 dishes of cells. Plate 5.107 cells per 10 cm culture dish
- Let the cells adhere 20-30 minutes
- Remove the medium and overlay the attached cells with 5 mL of fresh HL5c without PS
51 Infection
- Add 500 µL of bacteria per plate of host cells (Start with only 4 plates, and infect the 4 last ones
once the infection of the first ones is done)
- Seal the 10 cm dishes with strips of parafilm
- Centrifuge at 500 g (1,500 rpm) at RT for 2 x 12 min (in between rounds, gently “move” the dish to
redistribute bacteria and turn the dishes 180º)
- Let cells phagocytose for an additional 10-20 minutes
- Wash off extracellular bacteria with repeated rinses using HL5c without PS (2-5 washes, 10 mL
pipettes)
For isolation of 1 hpi mycobacteria-containing compartments:
- At 1 hpi, bang the dishes and take up the cells in 5 mL of HL5c without PS
- Pellet the cells 8 minutes at 2,000 rpm at 4ºC
For longer infections:
- Bang the dishes and take up the cells in 5 mL HL5c + PS (final 5 µg/mL streptomycin)
- Put the cells in an erlenmeyer and complete to 150 mL with HL5c + PS (final 5 µg/mL streptomycin)
- Incubate at shaking (130 rpm) at 25°C
- At the appropriate time, stop infection by pelletting 8 minutes at 2,000 rpm at 4°C
Homogenisation
- The homogenisation part is performed in the cold room
- To 6.75 mL HESES-azide buffer, add 250 µL Complete EDTA-free protease cocktail inhibitor
(Roche)
- Resuspend the pelleted cells in 2 mL homogenisation buffer
- Homogenise cells by passing them 8 times through the ball homogeniser (HGM, Germany) with a
barrel diameter of 8.000 mm and a ball diameter of 7.990 mm (clearance 10 µm)
- Add 3 mL of sucrose 71.4%, 500 µL of ATP 100 mM/sucrose and 50 µL of MgCl2 1M
52 - Mix 15 minutes on wheel
Sucrose gradients
- Prepare sucrose gradient in Ultra-clear centrifuge tubes (344060) for SW40 rotor
- Layer in each tube:
1 mL of 60% sucrose
5 mL of sample
3 mL of 40% sucrose
3 mL of 30% sucrose
1 mL of 10% sucrose
- Balance tubes with 10% sucrose for ultracentrifugation
- Ultracentrifuge over-night at 28,000 rpm, deceleration without break or “to 800 rpm”, at 4°C
Collection and storage of the mycobacteria-containing compartments
- Following centrifugation, collect material from the 10-30% interphase into a 15 mL Falcon tube
using a pipette Pasteur
- Dilute to 13 mL with membrane buffer
- Transfer the mycobacteria-containing compartments to Ultra-clear centrifuge tubes (344060) for
SW40 rotor
- Balance tubes for ultracentrifugation with membrane buffer
- Pellet the mycobacteria-containing compartments by ultracentrifugation at 35,000 rpm, brake on, for
1h30, at 4°C
- Snap freeze the pelleted mycobacteria-containing compartments in liquid nitrogen and keep the
pellets at -80ºC
53 2.2.3-Cell biology
2.2.3.1-Infections
Infections of Dictyostelium with Mycobacteria are performed as described in (Hagedorn and Soldati
2007) and in (Arafah, Kicka et al. 2013).
Preparation of the host cells
Dictyostelium cells are cultivated at least overnight in medium without antibiotics, in adherent or in
shaking conditions.
Cells grown in shaking condition are plated in 10 cm dish (5.107 cells/dish) at least 30 min before
infection.
After adhesion, replace medium with 5 mL of fresh HL5c without antibiotics.
Preparation of the mycobacteria
Mycobacteria are grown at OD600=0.8-1 in 7H9 (OD600=1 corresponds to 5.108 bacteria/mL).
For one infection, 5.108 mycobacteria (MOI=10) are washed twice in HL5c and passaged 5-10 times
through a 26-gauge needle to declump aggregates.
Infection
- Add 5.108 mycobacteria per plate of host cells
- Centrifuge at 500 g (1,500 rpm) at RT for 2 x 12 min (in between rounds, gently “move” the dish to
redistribute bacteria and turn the dish 180º)
- Let cells phagocytose for an additional 10-20 minutes
- Wash off extracellular bacteria with repeated rinses using HL5c without PS (2-5 washes)
-Bang the dishes and take up the cells in 5 mL HL5c + PS (final 5µg/mL streptomycin)
- Fill up to 37 mL with HL5c + PS (final 5 µg/mL streptomycin)
- Distribute in a 6-well dish (5 mL per well) and incubate for 2 days with shaking (130 rpm) at 25°C.
- The infection status is monitored at the following time points:
0.5, 12, 21, 37, 43 hpi or 0.5, 1, 2, 4, 6 hpi
54 Monitoring the infection
- Flow cytometry assay
Infections with GFP-expressing bacteria are monitored by FACS. Increased green fluorescence of the
Dictyostelium cells corresponds to intracellular mycobacterial growth in Dictyostelium cells.
A 500 µL aliquot of the infection is mixed with 500 µL of Sorensen-5 mM Azide and is pelleted 4
min. at 12,000 rpm, RT. The pellet is resuspended in 500 µL of Sorensen-Sorbitol. Prior to FACS
measurement, 4.5 µm fluorescent beads (100 beads/µL, YG-beads, PolySciences) are added to the
sample as an internal particle concentration standard.
FACS measurement is performed on a FACScalibur (Beckton Dickinson) and the data are analysed
with FlowJo (TreeStar, USA).
- Plate reader assay
Infections with bioluminescent Mycobacteria are monitored with a plate reader (Synergy Mx, Biotek)
measuring luminescence. Increased luminescent signal monitors growth of the mycobacteria inside
Dictyostelium cells.
The plate reader temperature is pre-set at 25ºC. 150 µL aliquots of the infection are deposited in a
white 96-well plate (F96 MicroWell™ Plates, non-treated from Nunc). 150 µL of HL5c are also
deposited to measure background signal. The luminescent signal is then measured with the help of the
plate reader.
- Microscopy
Dictyostelium cells are pelleted on Polylysine coated-coverslips. They are fixed by rapid freezing as
described in section 2.2.5.2. Immunostaining is performed as described in 2.2.5.2. Microscopy
pictures are acquired using a Leica SP2 or SP5 confocal using 100x oil immersion-objective.
2.2.3.2-Acidification and proteolysis assays
The acidification and proteolysis assays are performed as described in (Sattler, Monroy et al. 2013).
Briefly, for the acidification assay, beads or mycobacteria are labelled with 2 fluorophores: one pH
insensitive fluorophore (Alexa 594, TRITC), used as a reference, and one pH sensitive fluorophore
(FITC). Dictyostelium cells are fed with those beads or mycobacteria and the signal of both
fluorophores are measured with a fluorescent plate reader during several hours. The fluorescence
signal ratio of the pH sensitive versus the pH insentive fluorophores shows the evolution of the
intraphagosomal pH. For the proteolysis assay, beads are coupled via BSA to 2 fluorophores: one pH
insensitive fluorophore (Alexa 594, TRITC), used as a reference, and a self-quenching fluorophore
55 (DQgreen). Again, Dictyostelium cells are fed with those beads and the signal of both fluorophores are
measured with a fluorescent plate reader during several hours. During proteolysis, the fluorophores are
released from the beads and the self-quenching fluorophore starts to emit fluorescence. The
fluorescence signal ratio of the self-quenching fluorophore versus the pH insensitive fluorophore
shows the evolution of the proteolytic activity in phagosomes.
Mycobacteria labelling
- Dissolve FITC and TRITC in DMSO (5 mg/mL stock solutions).
- Wash 5.108 of overnight grown Mycobacteria (OD600=0.8-1) in PBS + 0.05% Tween 80, 4 min. at
12,000 rpm, RT.
- Resuspend the mycobacteria in 1 mL PBS pH 7.5 + 20 µL of FITC stock solution and 20 µL of
TRITC stock solution.
- Incubate for 2 h at 4ºC on wheel (Cover in foil).
- Wash at least 4 X with PBS + 0.05% Tween 80, 4 min. at 12 000 rpm, RT, until the supernatant is
clear.
- Resuspend the mycobacteria in 1 mL of LoFlow.
Beads preparation
- Wash 50 mg of 3 µm carboxylated silica particles (Kisker Biotech) three times with 1 mL of PBS by
brief vortexing and spin at 2,000 g for 60 s in a tabletop centrifuge.
- Resuspend beads in 700 µL of PBS (pH 7.2) containing 17.5 mg of cyanamide (concentration 25
mg/mL). The solution has to be freshly made!
- Incubate at room temperature with shaking for 15 min.
- Remove the cyanamide by washing twice with coupling buffer (0.1 M Sodium Borate, pH 8) at
2,000 g for 60 s in a tabletop centrifuge.
- Incubate overnight with agitation at 4°C in 1 mL of coupling buffer containing 5 mg of defatted BSA
for the pH-sensitive reporter beads or 1 mg of DQgreen-labeled BSA and 250 µg of defatted BSA for
the proteolysis-reporter beads.
- Wash beads twice with quenching buffer (250 mM glycine in PBS pH 7.2) to quench unreacted
cyanamide.
- Wash beads twice with coupling buffer to remove soluble amine groups.
56 - Resuspend beads in 700 µL of coupling buffer and add 20 µL of FITC and 20 µL of Alexa 594
succinimidyl ester stocks (for each total amount 0.25 mg) for the pH-sensitive reporter beads or 20 µL
Alexa 594 succinimidyl ester (total amount 0.25 mg) for proteolysis-reporter beads.
- Incubate for 1 h at room temperature with shaking to allow labeling of BSA with fluorophores. Wash
particles once with quenching buffer and twice with PBS.
- Resuspend in 1 mL of PBS with 0.01% w/v sodium azide as a preservative.
- Store at 4°C in the dark and avoid drying of the particles.
- Before adding beads to cells wash with PBS to remove sodium azide and prepare a dilution (final
concentration 1. 25.1010 beads/mL) in PBS.
Assay
Wavelength
FITC
TRITC
DQgreen
Alexa 594
Excitation
495
547
500
594
Emission
520
572
520
618
- Measurements are performed in clear bottom black side polystyrene 96-well plates (Cell Carrier or
Costar).
- 3.106 Dictyostelium cells/mL are washed twice with LoFlow medium (1,600 rpm for 4 min).
- To achieve a monolayer of cells, 100 µL of the suspension is added to each well of a 96-well plate
(equals 3.105 cells per well).
- After 20-30 min the cells should be attached, which should be checked at the microscope.
- Read background fluorescence with plate reader (Synergy Mx, Biotek).
- Addition of 10 µL of FITC/Alexa 594 beads (at a bead: cell ratio of 1:2 = 1.5.107 beads/1mL) or of
12 µL (MOI 20) or 6 µL (MOI10) of FITC-TRITC labelled mycobacteria.
- To synchronise phagocytosis, quick spin at 1,200 rpm until speed is up and then immediately stop
- Wash quickly 2-3x with 100 µL of LoFlow
- Measure emission fluorescence for the next 2-3 hours (measurements every 1 min) at 495 nm and
594 nm (pH-reporter beads), 495 nm and 547 nm (mycobacteria) or 520 nm and 618 nm (proteolysisreporter beads).
57 - The ratio 495/594 nm reflects the change of pH in the bead-containing phagosomes, the ratio
495/547 nm reflects the change of pH in the mycobacteria-containing phagosomes and the ratio
520/618 nm reflects the proteolytic activity in the bead-containing phagosomes.
- To calculate the pH of the phagosome a calibration curve has to be prepared
Calibration curve
- Reference pH buffers are prepared and distributed in a 96-well plate:
0.1 M piperazine- N,N’-bis(2-ethanesulfonate) (PIPES), 0.1 M KCl adjusted to pH 6, 6.5, 5,
7, 7.5, and 8 with 10 M NaOH.
0.15 M potassium acetate adjusted to pH 4, 4.5, 5, 5.5 with 5 M HCl.
- 10 µL of beads or of mycobacteria are added in the different reference pH buffers
- Fluorescence is measured at appropriate wavelengths.
- A calibration curve is obtained from the data and allows the calculation of the pH values
corresponding to the measured ratios.
2.2.3.3-Phagocytosis assay
The phagocytosis assay is performed as described in (Sattler, Monroy et al. 2013). Briefly,
Dictyostelium cells are incubated with fluorescent beads or GFP-expressing mycobacteria. Aliquots
are taken at different times and the fluorescence of the cells is measured by flow cytometry. The
measured fluorescence reflects the amount of ingested beads or mycobacteria, and consequently the
phagocytic efficiency of the cells.
Preparation of the cells
- Cells from a confluent dish are collected and counted using a Neubauer counting chamber.
- Cells are centrifuged at 500 g for 5 min.
- 107 cells are resuspended in 5 mL of fresh HL5c (HL5c without antibiotic if working with
mycobacteria) to have a cell density of 2.106 cells/mL and are deposited in a 6-wells plate.
- Cells are agitated for 2 h at 150 rpm on a horizontal shaker at 22ºC before starting the assay.
58 Preparation of the beads
-YG-carboxylated polysyrene beads (Polysciences) are used. The beads:cells ratio is adapted
according to the size of the beads used.
Bead diameter
(µm)
Bead stock
Final number of
Cell density
beads
(cells/mL)
3.64 x 1011
8 x 109
2 x 106
800:1
10
2 x 10
9
2 x 10
6
200:1
1 x 10
8
2 x 10
6
10:1
concentration
(beads/mL)
0.5
1
4.5
4.55 x 10
4.99 x 10
8
Bead to cell ratio
- Beads are washed in HL5c, 10 min., at 12,000 rpm, RT.
- Beads are washed a second time.
- Beads are resuspended in 800 µL of HL5c and sonicated 5 min. in a bath sonicator.
Material preparation
- Pre-cool the centrifuge at 4ºC.
- Prepare FACS tubes: 1/time point, 1 for the zero time point, 1 for beads only.
- Prepare 15 mL Falcon tubes filled with 3 mL Sorensen-Sorbitol-Azide: 1/time point, 1 for the zero
time point. Keep the tubes on ice.
- Prepare cut P1000 tips.
Assay
- Place 500 µL of cells only in the Falcon tube “zero” containing Sorensen-Sorbitol-Azide
- Centrifuge 10 min. at 1,200 rpm at 4ºC.
- Resuspend the pellet in 500 µL of Sorensen-Sorbitol using a cut P1000 tip and transfer into a FACS
tube. Keep on ice. This control cell sample is used to measure the cells’ autofluorescence.
- Start the assay by adding the beads to the cell suspension.
- 500 µL aliquots are taken at the following time points: 10, 20, 30, 40, 60 and 90 min.
- Each aliquot is processed as follows: deposit the aliquot into the corresponding Falcon tube
containing Sorensen-Sorbitol-Azide. Centrifuge 10 min. at 1,200 rpm at 4ºC. Resuspend the pellet in
500 µL of Sorensen-Sorbitol using a cut P1000 tip and transfer into a FACS tube.
59 - Keep samples on ice until the end of the assay.
- Samples are analysed using a FACS Calibur flow cytometer (Beckton Dickinson).
- Data are analysed using FlowJo (TreeStar, USA).
2.2.4-Biochemistry
2.2.4.1-Quantitative mass spectrometry
2.2.4.1.1-TMT labelling
The reduction, alkylation, digestion and TMT labelling is mainly performed as described by (Dayon,
Hainard et al. 2008). Briefly, the phagosomes or the mycobacteria-containing compartments pellets
are resuspended in 50 µL of TEAB (Triethylammonium hydrogen carbonate buffer) 0.1 M pH 8.5, 6
M urea. After addition of 50 mM of TCEP (tris-(2-carboxyethyl) phosphine hydrochloride), reduction
is performed during 1 h at 37˚C. The samples are then centrifuged 10 min, at 12,000 rpm to remove
the latex beads and the supernatants are collected. Supernatants are centrifuged and collected several
times until they are completely free of beads. 2 µL of each supernatant are taken to evaluate the
protein concentration in each sample with the help of a nanodrop. 100 µg of proteins are taken from
each sample and are alkylated at RT in the dark for 30 min. after addition of 1 µL of IAA (Iodoacetamide) 400 mM. Then, the volume of the samples is adjuted to 100 µL with TEAB 0.1 M pH 8.5.
2 µg of trypsin are added and the digestion is performed overnight at 37˚C. Each sample is labelled
with one TMT reagent according to manufacturer’s instructions and then, 50 µL of each labelled
sample are pooled and evaporated under speed-vacuum.
2.2.4.1.2-OGE
Off-gel electrophoresis is performed according to manufacturer’s instructions (Agilent). After
desalting, the mix containing the pooled labelled samples is reconstituted in OFFGEL solution. A 24wells frame is set-up on an Immobiline DryStrip pH 3-10, 24 cm and isoelectric focusing is performed
with those settings: 8,000 V, 50 µA, 200 mW until 20 kVh is reached. The 24 fractions are then
recovered and desalted using C18 MicroSpin columns.
2.2.4.1.3-Mass Spectrometry
ESI LTQ-OT MS is performed on an LTQ Orbitrap Velos from Thermo Electron (San Jose, CA,
USA) equipped with a NanoAcquity system from Waters. Peptides are trapped on a home-made 5 µm
200 Å Magic C18 AQ (Michrom) 0.1 × 20 mm pre-column and separated on a home-made 5 µm 100
Å Magic C18 AQ (Michrom) 0.75 × 150 mm column with a gravity-pulled emitter. The analytical
separation is run for 65 min using a gradient of H2O/FA 99.9%/0.1% (solvent A) and CH3CN/FA
99.9%/0.1% (solvent B). The gradient runs as follows: 0–1 min 95% A and 5% B, then to 65% A and
60 35% B at 55 min, and 20% A and 80% B at 65 min at a flow rate of 220 nL/min. For MS survey scans,
the OT resolution is set to 60000 and the ion population is set to 5 × 105 with an m/z window from 400
to 2000. A maximum of 3 precursors are selected for both collision-induced dissociation (CID) in the
LTQ and high-energy C-trap dissociation (HCD) with analysis in the OT. For MS/MS in the LTQ, the
ion population is set to 7000 (isolation width of 2 m/z) while for MS/MS detection in the OT, it is set
to 2 × 10E5 (isolation width of 2.5 m/z), with resolution of 7500, first mass at m/z = 100, and
maximum injection time of 750 ms. The normalized collision energies are set to 35% for CID and
60% for HCD.
2.2.4.1.4-Protein identification
Protein identification is perform with the help of the Easyprot platform (Gluck, Hoogland et al. 2013).
The Easyprot platform proceeds as follows: peak lists are generated from raw data using (ReadW).
After peaklist generation, the CID and HCD spectra are merged for simultaneous identification and
quantification
(Dayon,
Pasquarello
et
al.
2010)
and
http://www.expasy.org/tools/HCD_CID_merger.html). The peaklist files are searched against the
uniprot_sprot database (2011_02 of 08-Feb-2011). Dictyostelium discoideum, Mycobacterium
marinum and Mycobacterium smegmatis taxonomies are specified for database searching. The parent
ion tolerance is set to 10 ppm. TMT-sixplex amino terminus and TMT-sixplex lysine (229.1629 Da),
carbamidomethylation of cysteines are set as fixed modifications. Variable amino acid modifications
are oxidized methionine. Trypsin is selected as the enzyme, with one potential missed cleavage, and
the normal cleavage mode is used. All datasets are searched once in the forward and once in the
reverse database. Separate searches are used to keep the database size constant. Protein and peptide
scores are then set up to maintain the false positive peptide ratio below 1%. This results in a slight
overestimation of the false-positive ratio (Elias et al., 2007). For identification, only proteins matching
two different peptide sequences are kept.
2.2.4.1.5-Protein quantification
Isobaric quantification is performed using the IsoQuant module of Easyprot’s protein export as
described previously (Gluck, Hoogland et al. 2013). Briefly, a false discovery rate of 1% and a
minimum of 2 peptides per protein are selected. TMT sixplex is selected as reporter and a mass
tolerance of 0.05 m/z is used. The different protein ratios are calculated. A global normalisation of
“median log peptide ratio=0” and a confidence threshold of 95% are selected. EasyProt's Mascat
statistical
method,
inspired
by
Mascot's
quantification
module
(http://www.matrixscience.com/help/quant_statistics_help.html), and Libra, inspired by the TransProteomic
Pipeline's
isobaric
quantification
(http://tools.proteomecenter.org/wiki/index.php?title=Software:Libra),
are
chosen,
module
along
the
generation of the list of proteins featuring a ratio fold over 1.5.
61 2.2.4.2-One-dimensional SDS polyacrylamide gel electrophoresis (1D
SDS-PAGE)
SDS-PAGE allows the separation of protein samples on polyacrylamide gels. After reduction, proteins
migrate in the polyacrylamide gel according to their molecular weight. The gels can then be stained or
transferred to nitrocellulose membrane for immnunodetection of proteins of interest.
2.2.4.2.1-1D SDS-PAGE
Stock solutions
Separation gel
Stacking gel
8%
10%
12%
(6%)
30% acrylamide-bisacrylamide
2.7 mL
3.3 mL
4.0 mL
1 mL
1.5 M Tris pH 8.8
2.5 mL
2.5 mL
2.5 mL
-
1 M Tris pH 6.8
-
-
-
750 µL
ddH2O
4.6 mL
4.0 mL
3.3 mL
4.1 mL
SDS 10%
100 µL
100 µL
100 µL
80 µL
Bromophenol Blue 0.5%
-
-
-
10 µL
100 µL
100 µL
100 µL
80 µL
10 µL
10 µL
10 µL
8 µL
Ammonium persulfate (APS)
10%
TEMED
Polyacrylamide gels are poured in a BioRad III system and 10 wells-combs of 1 mm thickness are
used. Gels are run at 50 V in SDS-Tris-Glycine Buffer until the samples enter into the stacking gel.
Then, they are run at 150 V until the Bromophenol-Blue goes out of the gel.
2.2.4.2.2-Western blotting
Samples are transferred to a nitrocellulose membrane (Protran S, Schleicher & Schuell) using the
BioRad II wet transfer chamber with the Towbin buffer, at 30 V, O/N, at 4°C. Membranes are stained
with Ponceau S (0.2% Ponceau S, 3% tetrachloracetic acid (TCA), 3% sulfosalicylic acid). They are
then destained with ionised water before immunodetection.
2.2.4.2.3-Immunodetection
Membranes are blocked in PBS 5% milk for 1 h, RT or 3% milk, O/N, at 4°C. Then, they are
incubated with the first antibody diluted in PBS 3% milk for 1h or O/N. After 3 x 5 min washes in
PBS, the membranes are incubated with the secondary antibodies diluted 1/10000 in PBS 3% milk for
1 h. The signal of the secondary HRP linked antibody is detected with ECL reaction in a UVP
EpiChem II Darkroom equipped with a digital camera. ECL+ (the less sensitive), ECL Advance and
62 ECL prime (the more sensitive) (GE Healthcare) are available in the lab. The sensitivity of the used
ECL has to be adapted to the abundance of the immunodetected protein to avoid too low but also too
strong signal.
2.2.5-Microscopy
2.2.5.1-Live imaging
- 3 drops of a confluent dish are deposited in a 3.5 cm iBidi dish. 2 mL of medium (without PS if
working with mycobacteria) are added and the cells are grown overnight at 22ºC.
- 30 min. before the experiment, HL5c is removed and 2 mL of LoFlow are added.
- Prepare 1% agar in LoFlow in a beaker. Heat using microwave and then pour onto a glass plate to
obtain a 1 mm agar layer. Cut 1.5 x 1.5 cm squares.
Live acidification assay
- Remove LoFlow and add 10 µL of beads to the cells.
- Overlay the cells with a square of 1 mm thick agar.
- Absorb excess medium to flatten cells.
- Start imaging. Acquire pictures every 30 s to 1 min during 3-4 h using a spinning disc confocal
system (Intelligent Imaging Innovations Marianas SDC) mounted on an inverted microscope (Leica
DMIRE2). Imaging is performed with 63x glycerin or 100x oil immersions objectives.
Live infection
- Pre-cool the centrifuge at 4ºC.
- Prepare 4.107 mycobacteria as described in 2.2.3.1.
- Add the mycobacteria to the cells.
- Quick spin at 4ºC at 1,200 rpm until full speed is reached and then immediately stop.
- Remove LoFlow and overlay the cells with a square of 1 mm thick agar.
- Absorb excess medium to flatten cells.
- Start imaging. Acquire pictures every 30 s to 1 min during 3-4 h using a spinning disc confocal
system (Intelligent Imaging Innovations Marianas SDC) mounted on an inverted microscope (Leica
DMIRE2). Imaging is performed with 63x glycerin or 100x oil immersions objectives.
63 2.2.5.2-Immunofluorescence
Two cell fixation techniques are used in the lab: PFA (Paraformaldehyde) fixation or shock-freezing
fixation in -85ºC methanol.
Coverslips preparation
- Coverslips are placed in HNO3 for 1 h in shaking condition
- Rinse 10 x with ddH2O
- Wash 3 x in 100% ethanol
- Dry converslips in microwave
- Store between Kimwipes or distribute them in 24-well-plates
Fixation
- For methanol fixation:
Remove excess liquid with a Kleenex
Plunge rapidly the coverslips in MeOH -85ºC with a 15º angle
Transfer quickly to the rack
Rewarm slowly to -35ºC
- For PFA fixation
Prepare freshly 4% PFA in PBS on a hot plate
Distribute in 24-well plates and add coverslips to the wells
Fixe for 30 to 45 min.
Stop fixation by incubating the coverslips in PBS 100 mM glycine for 15 min.
Rinse coverslips in PBS and transfer them in 24-well plates containing fresh PBS
Coverslips can be stored at 4ºC for several weeks
64 Immunodetection
- Coverslips are blocked in PBS 0.2% gelatin for 30 min.
- Dilute primary antibody in blocking solution and deposit 50 µL of the dilution on a parafilm in a
humid chamber
- Put the coverslips cells face down on the drops
- Incubate 1 h at RT
- Rinse coverslips by repeated plunging in a beaker with PBS and deposit in 24-well plate with fresh
PBS for 5 min.
- Dilute secondary antibody in blocking solution and deposit 50 µL of the dilution on the parafilm
- Put the coverslips cells face down on the drops
- Incubate 1 h at RT
- Rinse coverslips by repeated plunging in a beaker with PBS and deposit in 24-well plate with fresh
PBS for 5 min.
- Remove excess liquid from the coverslips by tipping their side on a Kleenex and mount in a drop of
mounting medium (Prolong Antifade) on a glass slide
- Let the mounting medium harden one night at RT
2.2.5.3-EM
- Prepare freshly a fixative solution of 4% glutaraldehyde (EM grade), 0.6% OsO4 (very toxic, use
under hood).
- Take off the medium from the cells in the 6 cm or 10 cm dish and replace it with a 1:1 (v/v) diluted
medium with fixative solution.
- Incubate cells on the dish in fixative for 1h (under the hood).
- Scrape cells off the dish with a scraper, collect in an Eppendorf and pellet 5 min, 12,000 rpm.
- Wash the cell pellet twice with PBS.
- Send sample to the PFU plateform, CMU.
- The sample will be contrasted and embedded at the EM platform and section laid on grids.
65 66 3-Results
67 3.1-Establishment of an isolation procedure for the mycobacteria-containing compartments
In order to analyse the proteome of the mycobacteria-containing compartments, a procedure to isolate
these compartments had to be established. We decided to adapt the already well-established procedure
to isolate latex bead-containing phagosomes.
3.1.1-Latex-beads phagosomes isolation procedure
Previously in the lab, a procedure has been established and optimised to isolate homogeneous fractions
of latex bead-containing phagosomes (Dieckmann, Gopaldass et al. 2008). It consists in feeding cells
with latex beads. At different phagosome maturation times, the cells are homogenised. Then, the low
density of the latex allows to float up the latex bead-containing phagosomes on a sucrose gradient and
to separate them from other organelles of the cell. This is a powerful technique permitting the isolation
of a high number of phagosomes. Its use allowed to study the temporal profile of different phagosomal
proteins along the phagosomal pathway both by WB, IF, 2D and 2D-DIGE (Gotthardt, Warnatz et al.
2002, Gotthardt, Blancheteau et al. 2006, Dieckmann, Gueho et al. 2012, Gopaldass, Patel et al. 2012).
Furthermore, it also allowed the phagosome proteome analysis by MS (Boulais, Trost et al. 2010).
This technique allowed me to contribute to the Figure 6 in the paper “Role of magnesium and
phagosomal P-type ATPase in intracellular bacterial killing” from Lelong et al. 2011 presented in
appendix (Lelong, Marchetti et al. 2011).
This procedure has clearly proved its efficiency, and since our goal was to study the proteome
of mycobacteria-containing compartments, we were strongly interested to adapt it to isolate such
compartments.
3.1.2-Adaptation of the latex-beads phagosomes isolation procedure
To isolate mycobacteria-containing compartments, the strategy was to use the flotation properties of
the latex beads. The goal was to have both latex beads and mycobacteria in the same compartment and
then, float up the compartment on a sucrose gradient with the help of the latex buoyancy. To have both
beads and bacteria in the same compartment, the idea was to attach latex beads to mycobacteria. The
mycobacteria-containing compartments isolation strategy is presented in figure 1.
68 Figure 1: Strategy to isolate mycobacteria-containing compartments
1-Attachment of latex beads to mycobacteria
2-Preparation of pure inoculums of mycobacteria attached to latex beads
3-Infection of Dictyostelium
4-Homogenisation of the cells at times of interest
5-Flotation of compartments containing both mycobacteria and latex beads by ultracentrifugation of the
cell homogenate on a sucrose gradient
6-Recovery of mycobacteria-containing compartments
3.1.2.1-Formation of Bacteria+Beads Complexes (BBCs)
In order to confine both mycobacteria and latex beads to the same compartment, the best way was to
attach the latex beads directly onto mycobacteria. To do so, different approaches were considered such
as coupling the beads to antibodies against the mycobacteria cell wall, or to bind streptavidine-coated
beads to biotinylated mycobacteria. However, it appeared that the beads could be directly fixed to
mycobacteria without a linker, by hydrophobic adsorption. The use of a protocol normally used to fixe
proteins on latex beads was sufficient to stick the latex beads on mycobacteria (Figure 2A). After
several washes in borate buffer, the latex beads and the mycobacteria were then mixed together and
incubated for 2 h on a rotating wheel. Different sizes of beads (0.8 µm, 0.5 µm and 0.2 µm) and
different ratios of beads and bacteria were tested. The ratios were adapted according to the beads sizes:
ratio 2 and 15 for 0.8 µm beads, ratios 30 and 90 for 0.5 µm beads, and ratios 100 and 500 for 0.2 µm
beads. Interestingly, the highest ratios had an “individualising” effect on the mycobacteria during the
incubation. The high concentration of beads prevented the bacteria from sticking together to form big
bacteria clumps, and favored the formation of single-bacteria BBCs. The 0.8 µm beads formed big
BBCs, of twice the size of a single mycobacteria. But the 0.5 or 0.2 µm beads did not increase
drastically the size of the mycobacteria. To cover the whole bacterial surface, mycobacteria were
incubated with a large excess of beads and, at the end of the adsorption, not all the beads attached to
the bacteria. The BBCs were still mixed with free beads. However, our final goal was to isolate pure
fractions of compartments containing mycobacteria. In order to infect cells with a homogeneous
preparation of BBCs, the BBCs were separated from the free beads by ultracentrifugation of the
69 mixture on top of a one step sucrose gradient (Figure 2B). The BBCs collect at the 20%-60% sucrose
interphase, whereas the free latex beads stay at the 20% sucrose-medium interphase. Few bacteria not
coated with latex beads were found in the pellet. In a separate experiment, the BBCs were loaded on
the standard sucrose gradients usually used to isolate latex beads-phagosomes, and were observed to
float up to the 10%-25% sucrose interphase, like the latex-beads phagosomes (Figure 2C). This
confirmed that the BBCs could be used to isolate mycobacteria-containing compartments. However,
some BBCs were also found at the other interphases, probably too dense to float to the top of the
gradient. To increase the yield of BBCs floating to the top interphase, the sucrose gradient used for the
isolation of latex bead-phagosomes (10%, 25%, 35% and 60% sucrose) was adapted to 10%-30%40%-60% sucrose.
70 A
0.8)µm
2
15
30
90
100
500
0.8)µm
0.5)µm
0.2)µm
0.8)µm
0.5)µm
0.2)µm
0.5)µm
0.2)µm
B
Medium120%)sucrose
20%160%)sucrose
pellet
C
10%125%)sucrose
25%135%)sucrose
35%1sample
pellet
Figure 2: Latex beads adsorb onto mycobacteria and float them up on a sucrose gradient. A. M marinum-GFPBBCs were prepared with different beads size and with different beads:bacteria ratios. Prepared BBCs were
visualised by microcopy. Right panel; TRITC-couled Ab were adsorbed on beads during BBCs preparation.
Scale bar, 1 µ m. B. M marinum-GFP-BBCs were prepared with 0.8 µm, 0.5 µm or 0.2 µm latex beads at
beads:bacteria ratios of 15, 90 and 500 respectively. The prepared BBCs+free beads mixture was centrifuged on
71 a one step sucrose gradient. Fractions recovered at the different interphase were visualised by microscopy. C. M
marinum-GFP-BBCs prepared with 0.8 µm, 0.5 µm or 0.2 µm latex beads at beads:bacteria ratios of 15, 90 and
500 respectively were separated from free beads by ultracentrifugation on a one step sucrose gradient and loaded
on the sucrose gradient used to isolate latex-beads phagosomes. The majority of the BBCs float up at the 10%25% sucrose interphase.
3.1.2.2-BBCs are phagocytosed and are recognised as mycobacteria
BBCs are mycobacteria surrounded by beads. They are slightly bigger than single mycobacteria and
have an irregular shape. In order to test whether Dictyostelium cells could phagocytose them, an
infection with non-coated M. marinum or with M. marinum-BBCs prepared with the different bead
size was performed. Cells were infected by spinoculation. After washing the extracellular bacteria or
BBCs, samples of each condition were taken. The use of GFP-expressing strains allowed to evaluate
the number of infected cells by FACS (Fluorescence-activated cell sorting). By FACS analysis, the
events are plotted on a graph according to their side-scattering (FSC) and their fluorescence (FL-1).
These parameters allowed to distinguish three different populations on the graph: a population of nonfluorescent cells, corresponding to the non-infected cells, a population a fluorescent cells
corresponding to the infected cells, and a third fluorescent population, corresponding to extracellular
mycobacteria or BBCs. The percentage of infected cells was dependent on the particle size (Figure
3A). Indeed, the BBCs prepared with 0.8 µm beads infected 58% less cells than the non-coated M.
marinum, and the ones prepared with 0.5 µm beads, 50% less cells. The BBCs prepared with 0.2 µm
beads infected only 23% less cells than non-coated M. marinum. These 0.2 µm-BBCs are the smallest
and have the closest size to non-coated M. marinum. It was not surprising that the percentage of
infection was the closest to infection performed with non-coated M. marinum. For this reason, the 0.2
µm-BBCs were chosen for the establishment of the procedure to isolate mycobacteria-containing
compartments. If not specified, the rest of the experiments were performed with 0.2 µm-BBCs. Cells
infected with 0.2 µm-BBCs were observed by EM at 21 hpi (hours post infection) and by IF at 12 hpi.
By EM, the M. marinum-BBCs appeared to reside in closed membranous compartments (Figure 3B).
The latex beads (arrowhead) were also visible inside the compartment. By IF, the membrane around
the M. marinum-BBCs was detected positive for the phagosomal marker p80 (arrowhead) (Figure 3C).
The latex beads (arrow) were clearly observed around the bacteria in phase contrast, confirming that
this was really a BBC-containing compartment and not a non-coated bacteria-containing phagosome.
Furthermore, both EM and IF pictures showed that the latex beads and the bacteria reside and remain
in the same compartment, even after several hours of infection (21 hpi for EM, 12 hpi for IF). This
confirms that the BBCs are ingested by phagocytosis and reside in a phagosome.
LmpB-null cells are specifically defective for the ingestion of mycobacteria. They show a 40%
defect for the ingestion of M. marinum compared to wild type cells (Sattler et al., in preparation;
figure in appendix). To determine whether the M. marinum-BBCs are recognised by Dictyostelium as
beads or as mycobacteria, the capacity of lmpB-null cells to ingest BBCs was tested. The cells were
incubated with BBCs prepared with GFP-expressing M. marinum. The fluorescence of the cells,
increasing proportionally with bacteria uptake, was followed by FACS. The ingestion of BBCs was
72 highly impaired in lmpB-null cells (Figure 3D). Indeed, they ingested 49% less BBCs than the wild
type cells. This indicated that BBCs were recognised as mycobacteria. Furthermore, before separating
the BBCs from the free beads during the BBCs preparation, an IF was performed on the mix
BBCs+free beads. In parallel, an IF was also performed on beads, which have not been incubated with
mycobacteria. To better visualise the beads, this experiment was performed with 0.8 µm beads. When
beads were incubated with mycobacteria, they were detected by a cocktail of antibodies antimycobacteria cell wall (Figure 3E). The beads adsorbed to the mycobacteria were even more strongly
detected. On the opposite, beads that have not been incubated with mycobacteria were not
significantly detected. This indicates that some material composing the mycobacteria cell wall seems
to diffuse around the adsorbed beads and cover them. Moreover, some of the mycobacteria cell wall
material seems to be shed in the incubating buffer and adsorbed onto the beads.
73 A
B
D
AX2
LmpB
120
100
80
60
40
20
0
100
80
60
40
20
0
0
20
40
AX2
LmpB
120
BBCs+uptake++(%)
M.+marinum+uptake++(%)
C
60
80
Time+(minutes)
100
0
20
40
60
80
100
Time+(minutes)
E
0.8$µm$beads
0.8$µm$beads$+$M.$marinum
0.2$µm$beads$
+
$M.$marinum
Figure 3: BBCs are recognised and phagocytosed as mycobacteria by Dictyostelium. A. Wild type cells were
infected by spinoculation with M. marinum-GFP or BBCs prepared with M. marinum-GFP and different size of
latex beads. The number of infected cells in each condition was determined by counting the number of
fluorescent cells by FACS. The values were normalised to the number of infected cells obtained for the infection
performed with non-coated M. marinum-GFP. B. Wild type cells were infected with 0.2 µm-M. marinum-GFPBBCs and at 21 hpi, processed for Transmission Electron Microscopy (TEM). Arrowheads indicate 0.2 µm latex
74 beads. Scale bar, 1 µm. C. Wild type cells were infected with 0.2 µm-M. marinum-GFP-BBCs. At 21 hpi, cells
were stained for p80 (red). Arrowhead indicates p80 positive membrane. Arrow indicates 0.2 µm latex beads on
the phase picture. Scale bar, 10 µm. D. The uptake of non-coated M. marinum and of 0.2 µm-M. marinum-BBCs
by wild type and LmpB-null cells was measured by FACS. Relative fluorescence was normalised to the value
obtained for wild type cells ate 90 minutes. The curve represents mean and SD of 2 independent experiments. E.
BBCs were prepared with M. marinum-GFP and 0.8 µm or 0.2 µm latex beads and stained for antimycobacterial cell wall material with a cocktail of antibodies (anti-M. leprae membrane antigens, anti-M. leprae
surface antigens, anti-M. leprae cell wall associated antigens) (Red). Scale bars, 1 µm.
3.1.2.3-The BBCs’mycobacteria are still alive and infectious
In order to test whether the various mechanical, osmotic and chemical stresses applied during the BBC
preparation affect the bacteria, their infectiosity was tested as follows. Dictyostelium was infected with
BBCs prepared with different mycobacteria strains and in parallel with the same mycobacterial strains
not coated with beads: the pathogenic strain M. marinum, the avirulent strain M. marinum-L1D and
the non pathogenic strain M. smegmatis. The use of GFP-expressing strains allowed to follow the
infection by FACS (Figure 4A). As previously described (Hagedorn and Soldati 2007), the population
of infected cells rapidly disappeared when Dictyostelium was infected with the avirulent M. marinumL1D strain or the non-pathogenic M. smegmatis strain. For the infection carried out with M.
smegmatis, the population of infected cells had clearly disappeared at 18 hpi on the FACS profile. For
the infection performed with M. marinum-L1D, the population of infected cells decreased between 0.5
and 6 hpi. This decrease was explained by the exocytosis of the avirulent mycobacteria within a few
hours. Indeed, the population of extracellular bacteria increased. However, at 18 hpi, the population of
infected cells had almost disappeared and the population of extracellular M. marinum-L1D was also
decreased. M. marinum-L1D is actually re-ingested several times before being killed. However, when
Dictyostelium was infected with the pathogenic strain M. marinum, the population of infected cells
remained, even at 21 hpi. The infections performed with BBCs showed similar behaviour.
Dictyostelium cells infected with M.marinum-L1D-BBCs or M. smegmatis-BBCs could cure the
infection in several hours, whereas, cells remained infected 21 hpi when Dictyostelium was infected
with M. marinum-BBCs. The fate of the different BBCs is really dependent on the mycobacteria
strain. These results were also confirmed by EM and IF. Indeed, a phagosome containing an inert
particle such as a bead, usually completes the whole maturation process in a few hours. Between 2 and
three hour after ingestion, the bead is exocytosed. However, when Dictyostelium was infected with M.
marinum-BBCs, infected cells were still detected by EM at 21 hpi (Figure 3B) or by IF at 12 hpi
(Figure 3C). These results clearly demonstrate that the mycobacteria incorporated in the BBCs are still
alive and infectious.
Furthermore, it has been shown that at late stages of infection (37 hpi), the pathogenic strain
M. marinum ruptures its compartment and escapes into the cytosol (Hagedorn and Soldati 2007). M.
marinum initially coming from BBCs were also observed by EM to escape their compartments at late
times of infection (21 hpi) (Figure 4B). Even at this late stage of infection, some latex beads were still
observed around the escaping M. marinum (Figure 4B, arrowhead). We conclude that the beads do not
75 disturb the normal infection cycle of M. marinum. The BBCs behave similarly to non-coated
mycobacteria.
A
0.5$hpi
Smeg* .460
6$hpi
Smeg* .473
104
104
103
10
2
101
10
0
10
102
0
10
1
2
10
10
FL1/H:*FL1/Height
3
10
10
2
10
1
10
0
SSC/H:*SSC/Height
10
SSC/H:*SSC/Height
ssc
SSC/H:*SSC/Height
M.$smeg
SSC-H: Side Scatter
103
FL01
100
101
102
103
FL1-H: green FL
104
3
10
2
101
10
0
101
10
10
10
3
10
3
10
2
10
1
102
103
FL1/H:*FL1/Height
104
10
0
10
1
2
10
10
FL1/H:*FL1/Height
3
10
10
2
10
1
10
0
4
10
0.5$hpi
12$hpi
10
3
10
3
103
103
103
2
10
2
10
2
10
1
10
1
10
1
10
0
10
0
10
0
4
10
0
10
ax2*L1D*0.2m*.367
1
2
10
10
FL1/H:*FL1/Height
3
10
FL01
4
10
104
103
103
103
2
SSC/H:*SSC/Height
SSC/H:*SSC/Height
104
SSC/H:*SSC/Height
104
10
102
101
10
100
10
1
10
10
1
2
10
10
FL1/H:*FL1/Height
FL01
3
10
4
10
1
2
10
10
FL1/H:*FL1/Height
3
10
10
10
1
2
10
10
FL1/H:*FL1/Height
FL01
3
10
4
101
102
103
FL1-H: green FL
FL01
104
4
10
3
10
2
10
0
10
1
2
10
10
FL1/H:*FL1/Height
FL01
3
10
4
10
102
101
102
103
FL1-H: green FL
FL01
0
104
3
10
2
3
10
10
FL1-H: FL1-Height
FL01
4
10
4
1
10
2
3
10
3
10
10
10
FL1-H: green FL
FL01
4
3
2
10
1
10
0
1
10
0
10
4
0
10
0
10
10
ax2 marWT 0.2m.335
101
10
3
2
104
102
101
2
10
10
FL1/H:*FL1/Height
100
100
ax2 marWT 0.2m.331
10
1
101
100
100
ax2 marWT 0.2m .326
100
0
100
4
FL01
101
100
0
0
ax2*L1D*0.2m.372
10
10
101
SSC-H: SSC-Height
10
102
SSC-H: SSC-Height
3
SSC-H: SSC-Height
2
10
10
FL1/H:*FL1/Height
FL01
ssc
1
M.$mar0BBC
10
SSC-H: Side Scatter
3
10
SSC-H: Side Scatter
10
SSC-H: Side Scatter
104
ssc
104
M.$mar
104
SSC/H:*SSC/Height
4
101
104
21$hpi
10
102
102
103
FL1/H:*FL1/Height
FL01
4
0
0
FL01
10
SSC/H:*SSC/Height
ssc
SSC/H:*SSC/Height
M.$mar0L1D
18$hpi
101
FL01
4
4
10
ssc
100
Smeg* 0.2m.477
10
ax2*L1D*0.2m.356
M.$mar0L1D0BBC
6$hpi
1
104
4
FL01
0.5$hpi
102
103
FL1/H:*FL1/Height
100
100
2
10
SSC/H:*SSC/Height
10
101
Smeg* 0.2m.472
SSC/H:*SSC/Height
10
SSC/H:*SSC/Height
100
ssc
M.$smeg0BBC
101
10
FL01
Smeg*0.2m*.461
4
3
100
100
4
18$hpi
Smeg* .475
4
103
10
104
10
0
10
1
10
2
3
10
10
FL1-H: FL1-Height
FL01
4
10
0
10
1
10
2
10
10
FL1-H: FL1-Height
4
FL01
B
Figure 4: BBCs are infectious and behave as non-coated mycobacteria. A. Wild type cells were infected by
spinoculation with M. marinum-GFP, M. marinum-L1D-GFP, M. smegmatis or 0.2 µm BBCs prepared with
these different mycobacteria strains. Extracellular bacteria or BBCs were removed and the cells were cultivated
for two days at 32ºC. At the indicated time points, samples were taken and the infection was monitored by
FACS. For each infection, representative SSC vs FL-1 plots are presented. Three different populations can be
distinguished on the SSC vs FL-1 plots and are circled on the SSC vs FL-1 plot on the left: non-infected cells
(red), infected cells (black) and extracellular bacteria or BBCs (green). B. Wild type cells were infected with 0.2
µm-M. marinum-BBCs. At 21 hpi, cells were processed for Transmission Electron Microscopy (TEM).
Arrowheads indicate 0.2 µm latex beads. Scale bar, 1 µm.
76 3.1.2.4-Isolation of BBCs-containing compartments
The isolation of BBCs-containing compartments is performed similarly as the isolation of latex beadsphagosomes. After preparation of the BBCs, Dictyostelium cells were infected. The extracellular
BBCs were washed away and then, the cells were incubated at 25ºC. At times of interest, the cells
were pelleted at 4ºC and resuspended in cold HESES buffer to block further maturation of the BBCscontaining compartments. The isolation of BBCs-containing compartments was first tested at 1 hpi.
After homogenisation of the cells with the help of a ball homogeniser, the homogenate was incubated
with ATP. This incubation step allows to detach from the BBCs-containing compartments all the
cytoskeletal proteins which are bound in an ATP dependent manner. This allows the isolation of
compartments without contamination with high amounts of cell cytoskeleton (Dieckmann, Gopaldass
et al. 2008). After ultracentrifugation of the cell homogenate on a sucrose gradient, the top interphase
(10%-30% sucrose) was recovered. The recovered interphase contained a high number of BBCs
(Figure 5A). Since extracellular BBCs were washed away during the infection, this recovered material
was likely entirely BBCs-containing compartments. To verify the presence of a membrane around the
recovered BBCs, a FM4-64 staining and IF were performed. FM4-64 is a lipid dye. A positive FM464 staining was observed around M. marinum (Figure 5B). By IF, several phagosomal markers were
detected around the different mycobacteria strains-BBCs (Figure 5C). These results confirmed the
presence of a membrane around the isolated BBCs and showed that mycobacteria-containing
compartments can be isolated. The IF pictures already revealed differences between the various
mycobacteria-containing compartments. At 1 hpi, the M. marinum-containing compartment has
already diverted from the normal phagosomal pathway but is still a young M. marinum-containing
compartment. To the contrary, M. marinum-L1D or M. smegmatis-containing compartments follow
the normal phagosomal pathway and at 1 hpi, they are “mature phagosomes” almost at the end of the
maturation process. This was confirmed by the IF results. Indeed, compared to M. marinum-L1D or
M. smegmatis-containing compartments, only few M. marinum-containing compartments were
positive for the late phagosomal markers p80 and vacuolin. Only 11% of the M. marinum-containing
compartments were positive for p80, whereas 50% were detected as p80 positive for M. marinumL1D-containing compartments and 35% for M. smegmatis-containing compartments. Similarly, for
vacuolin, only 7% of M. marinum-containing compartments were detected positive, whereas 33% of
the M. marinum-L1D-containing compartments and 24% of the M. smegmatis-containing
compartments were vacuolin positive. Interestingly, 8% of the M. marinum-containing compartments
were detected positive for the H+-vATPase. Even if some studies showed that the M. tuberculosiscontaining compartment fails to acidify (Sturgill-Koszycki, Schlesinger et al. 1994, Mwandumba,
Russell et al. 2004, Pethe, Swenson et al. 2004), this result is in agreement with a previous study,
which showed that more than 50% of the M. marinum-containing compartments transiently acquire
the H+-vATPase during the first hour of infection (Hagedorn and Soldati 2007). However, more M.
marinum-L1D (68%) or M. smegmatis-containing compartments (32%) were detected as H+-vATPase
positive, confirming that M. marinum-L1D or M. smegmatis-containing compartments follow the
normal phagosomal pathway. Overall, even if the percentages calculated for the isolated
compartments were lower than previously published (Hagedorn and Soldati 2007), they confirmed the
differences observed between the different mycobacteria strains in this previous study (Hagedorn and
Soldati 2007). In this study, the compartments positive for the different markers were counted on
77 intact infected cells. During the isolation procedure, the membrane of some BBCs-containing
compartments could be damaged. This could explain why the percentages obtained for the isolated
mycobacteria-containing compartments were lower.
A
B
FM4664
M.3marinum
Merge
Phase
C
M.3marinum
M.3marinum6L1D
p80
vATPase
vacuolin
Figure 5: Isolation of compartments containing different mycobacteria strains. Wild type cells were infected
with 0.2 µm-M. marinum-GFP-BBCs. At 1 hpi, cells were homogenised. After incubation with ATP, the cell
homogenate was ulcentrifuged on a sucrose gradient. A. The fraction recovered at the 10%-30% sucrose
78 interphase was observed by microscopy. Scale bar, 5 µm. B. The recovered fraction was stained FM4-64. C.
Compartments containing 0.2 µm BBCs prepared with M. marinum-GFP, M. marinum-L1D-GFP or M.
smegmatis-GFP (green) were isolated and stained for p80, VatM or vacuolin (red). The number of bacteria
positive for the different markers was counted for each mycobacteria strain. Scale bars, 1 µm.
By establishing this procedure, we were able to isolate mycobacteria-containing
compartments. We then divided the proteome analysis project of mycobacteria-containing
compartments into two distinct parts. One part consisted in analysing the temporal modification of the
M. marinum-containing compartment during the early phase of infection. The second part consisted in
comparing the proteome of M. marinum-containing compartments to the proteome of compartments
containing avirulent, non-pathogenic or attenuated mycobacteria strains during the early phase of
infection. The different proteomic comparisons performed are presented in table I.
M. marinum
M. marinum-containing compartment 3 hpi
M. marinum-containing compartment 6 hpi
vs
M. marinum-containing compartment 1 hpi
vs
M. marinum-containing compartment 1 hpi
Manipulated M. marinum containing compartment
vs
Non or less-manipulated mycobacteria-containing compartments
Compartments isolated at 1 hpi
Compartments isolated at 6 hpi
M. marinum
M. marinum
M. marinum
M. marinum
M. marinum
vs
vs
vs
vs
vs
M. marinum-L1D
M. smegmatis M. marinum RD1 M. smegmatis M. marinum RD1
Table I: Proteomic comparisons. The table lists the different isolated compartments and the different proteomic
comparisons performed during this study.
3.2-Proteomic characterisation of M. marinum-containing compartments during the early phase
of infection
As previously observed by CFU counting, during the first twelve hours of infection, M. marinum does
not replicate (Hagedorn and Soldati 2007). This phase has been designated as the manipulation phase.
Indeed, immediately after ingestion, M. marinum has to fight against the hostile environment
encountered in its containing phagosome. Furthermore, in order to establish an environment
permissive for its replication, it actively interacts with its host to modulate the maturation of the
phagosome. In contrast, the non pathogenic strain M. smegmatis and the avirulent strain M. marinumL1D are not able to block the phagosome maturation. They are rapidly killed. The modifications
inflicted to this compartment during the early phase of infection are poorly known. Studies performed
79 on tuberculosis infection revealed that Rab5, the early endosomal marker, is retained onto the
mycobacteria-containing compartment whereas some of its effectors like EEA1 or PI3K, and the H+vATPase are excluded. To better characterise the M. marinum-containing compartment during the
early phase of infection and to identify new markers of this compartment, the quantitative
modifications of the M. marinum-containing compartment proteome was analysed. The M. marinumcontaining compartment proteome was also compared to the proteome of non-manipulated
compartments containing avirulent, non pathogenic or attenuated mycobacteria.
3.2.1-The M. marinum-containing compartment proteome
In order to study the temporal modification of the M. marinum-containing compartment during the
early phase of infection, M. marinum-containing compartments were isolated at 1, 3 and 6 hpi.
Biological duplicates were prepared for each time points. The isolated compartments were then
reduced, alkylated, digested with trypsin and finally labelled with isobaric TMT tags. Equal amounts
of each sample were mixed and an OGE was performed on the mixed peptides to separate them in 24
fractions. Each fraction was then analysed by LC-MS/MS. 1461 proteins from Dictyostelium and 555
proteins from M. marinum were identified. Each protein was identified with at least two unique
peptides. These proteins constitute the total M. marinum-containing compartment proteome during the
first six hours of infection. The rest of the study will be focused on the host proteins. 1313 of the
identified Dictyostelium proteins were associated with a GO term and 1447 were matched in KEGG
pathways using the Orange freeware (http://orange.biolab.si/). A stringent p-value of 10-5 and a strong
enrichment score (over 3) were applied to select only significantly represented GO-terms (Table II).
The enriched GO-terms representing “Cellular components” reflected the endo-phagosomal nature of
the M. marinum-containing compartment and its interaction with cytoplasmic vesicles. Only two GOterms represented different components: nucleolus and ribosome. Their enrichment in the M.
marinum-containing compartment might be explained by the degradative function of phagosomes.
Indeed, the corresponding proteins might have been targeted to the phago-endosomal pathway by
autophagy to be degraded. The enriched GO-terms representing “Biological process” and “Molecular
functions” revealed the presence of actin, actin-binding proteins and proteins potentially involved in
actin cytoskeleton regulation on the M. marinum-containing compartment. Indeed, some studies
showed that the actin-binding protein Coronin is retained on pathogenic mycobacteria-containing
compartments (Ferrari, Langen et al. 1999, Fratti, Vergne et al. 2000, Deghmane, Soualhine et al.
2007). More recently, the importance of the actin coat around M. marinum-containing compartments
during the first hours of infection was demonstrated (Kolonko et al., in revision at Cell Micro). Among
the enriched GO-terms representing “Biological process” and “Molecular functions” were also GOterms confirming the presence of ribosomal proteins in the M. marinum-containing compartment.
Finally, the GO-terms “response to biotic stimulus”, “response to other organism” and “response to
bacterium” were found among the strongly enriched “Biological process” GO-terms. They reflected
the presence of M. marinum in the compartment and host-pathogen interaction. Furthermore, it also
confirmed the phagosomal nature of the compartment, which primarily functions in the digestion of
bacteria. These results obtained with the GO-terms were confirmed by the KEGG pathways in which
80 the proteins were matched. A p-value of 10-5 was applied to keep only significantly represented KEGG
pathways (Table II). The pathways “endocytosis” and “phagosome” confirmed the endo-phagosomal
nature of the M. marinum-containing compartment. Furthermore, these endosomal and phagosomal
proteins did not necessarily correspond to early endosomes or early phagosomes showing that the M.
marinum-containing compartment is more complex than just a phagosome arrested at an early step of
maturation. The KEGG pathways “ribosome”, “ribosome biogenesis in eukaryotes”, “protein
processing in ER” and “RNA transport” confirmed the presence of ribosomal proteins and more
generally, the presence of proteins involved in translation. Again, these proteins might have been
targeted to the phago-endosomal pathway by autophagy to be degraded. 25 proteins matched in the
KEGG pathway “oxidative phosphorylation”. Among these proteins are found mitochondrial proteins
but also the H+-ATPase. This clearly demonstrated that M. marinum do not totally avoid the
establishment of a hostile environment inside the phagosome and indeed, encounters stresses such as
acidic pH after being ingested. It is also interesting to notice that 7 proteins matched in the
“phosphatidylinositol signalling system”. Indeed, phagosome maturation is dependent of the
phosphatidylinositol composition of its membrane and it would be interesting to know the
phosphatidylinositol
composition
of
the
M.
marinum-containing
compartment
membrane.
Furthermore, it has been shown that PI3K is excluded from M. tuberculosis-containing compartment
and that M. tuberculosis secreted protein SapM directly modifies the phosphatidylinositol composition
of this compartment by dephosphorylating PI3P (Fratti, Backer et al. 2001, Chua and Deretic 2004,
Hestvik, Hmama et al. 2005, Vergne, Chua et al. 2005). The “citrate cycle (TCA cycle)” was also
found significantly represented in the M. marinum-containing compartment. M. tuberculosis is known
to divert its isocitrate from the TCA cycle to the glyoxylate cycle (McKinney, Honer zu Bentrup et al.
2000, Munoz-Elias and McKinney 2005). It is also known to use the host cholesterol (Pandey and
Sassetti 2008). The regulation of the host enzymes of the TCA cycle could divert the carbon flux into
the fatty acid biosynthesis. Then, similarly to M. tuberculosis, M. marinum could use the fatty acids
produced by the host. However, the presence of this pathway among the significantly represented
KEGG pathways could also simply reflect the nutrition function of the phagosome in Dictyostelium.
This first qualitative analysis of the M. marinum-containing compartment proteome was
followed by a temporal quantitative proteomic analysis. Furthermore, it also allowed a qualitative
comparison with the phagosome proteome, which was already deeply investigated in the group. The
results of these analyses will be detailed in the following sections.
81 GO Terms (cellular components):
GO Terms (biological process):
M. marinum-containing
compartment
Whole cell
P-value:
Enrichment
3.83
64 (4.87%)
105 (1.27%)
<0.00001
196 (14.93%)
324 (3.93%)
<0.00001
3.8
200 (15.23%)
332 (4.03%)
<0.00001
3.78
241 (18.35%)
425 (5.16%)
<0.00001
3.56
233 (17.75%)
412 (5.00%)
<0.00001
3.55
236 (17.97%)
418 (5.07%)
<0.00001
3.54
233 (17.75%)
413 (5.01%)
<0.00001
3.54
22 (1.68%)
40 (0.49%)
<0.00001
3.45
29 (2.21%)
53 (0.64%)
<0.00001
3.44
29 (2.21%)
53 (0.64%)
<0.00001
3.44
40 (3.05%)
74 (0.90%)
<0.00001
3.39
70 (5.33%)
131 (1.59%)
<0.00001
3.36
40 (3.05%)
75 (0.91%)
<0.00001
3.35
34 (2.59%)
66 (0.80%)
<0.00001
3.23
44 (3.35%)
91 (1.10%)
<0.00001
3.04
47 (3.58%)
98 (1.19%)
<0.00001
3.01
M. marinum-containing
compartment
Whole cell
P-value:
Enrichment
4.19
64 (4.87%)
96 (1.16%)
<0.00001
GO:0006364: rRNA
41 (3.12%)
66 (0.80%)
<0.00001
3.9
GO:0016072: rRNA
41 (3.12%)
67 (0.81%)
<0.00001
3.84
65 (4.95%)
107 (1.30%)
<0.00001
3.81
21 (1.60%)
36 (0.44%)
<0.00001
3.66
21 (1.60%)
36 (0.44%)
<0.00001
3.66
43 (3.27%)
79 (0.96%)
<0.00001
3.42
43 (3.27%)
79 (0.96%)
<0.00001
3.42
39 (2.97%)
73 (0.89%)
<0.00001
3.35
27 (2.06%)
54 (0.66%)
<0.00001
3.14
27 (2.06%)
55 (0.67%)
<0.00001
3.08
M. marinum-containing
compartment
Whole cell
P-value:
Enrichment
18 (1.37%)
20 (0.24%)
<0.00001
5.65
26 (1.98%)
49 (0.59%)
<0.00001
3.33
62 (4.72%)
120 (1.46%)
<0.00001
3.24
GO Terms (Molecular function):
GO:0030515: snoRNA binding
KEGG Pathways
M. marinum-containing
compartment
1
Whole cell
P-value
<0.000001
<0.000001
<0.000001
<0.000001
<0.000001
<0.000001
<0.000001
<0.000001
A
RNA transport
<0.000001
0.00001
Table II: Number and percentage of host proteins matched in GO-terms and KEGG pathways. Only
significantly represented GO-terms (p-value < 10-5, enrichment > 3) and significantly represented KEGG
pathways (p-value > 10-5) are listed.
82 3.2.2-Comparison of the early M. marinum-containing compartment proteome to the
total phagosome proteome
Like for M. marinum-containing compartments, a similar proteome analysis had been performed on
Dictyostelium phagosomes. For a number of studies in the group in the last decade, latex-beads
phagosomes have been isolated at different maturation times in order to cover the complete
phagosome maturation pathway. In some studies, the phagosomal proteins were separated by 2D-GE,
and in some others by 1D-SDS-PAGE. Then, all the separated proteins were identified by LC-MS/MS.
A list of all the identified phagosomal proteins in these different studies was compiled to constitute the
exhaustive reference Dictyostelium phagosome proteome. This list has been analysed and the results
have been published (Dieckmann, Gueho et al. 2012). By matching the phagosomal proteins to their
GO-term annotations and in KEGG pathways using the Orange freeware, I contributed to the table I
and the supplemental figure 2 of this article, presented in appendix.
A total of 1291 phagosomal proteins were identified. 1124 of them had known GO-terms
annotations and 1288 could be matched in KEGG pathways. A stringent p-value of 10-5 and an
enrichment of 2 were applied to select only significantly enriched GO-terms, and a p-value of 10-4 was
applied to select only significantly represented KEGG pathways (Table III). Among the enriched GOterms, the basic functions of the phagosome were represented, such as vesicle trafficking (“Vesiclemediated transport” and “protein intracellular transport”), reorganisation of the cytoskeleton (“actin
cytoskeleton organisation”), acidification by ATP dependent H+-pump (“ATP synthesis coupled
proton transport” and “purine nucleoside triphosphate metabolic process”) and finally bacteria killing
and digestion (“response to bacterium”). Not surprisingly, the KEGG pathways “phagosome” and
“endocytosis” were represented in the phagosome proteome. Several metabolic pathways were also
found significantly present, reflecting the nutrition function of the phagosome in Dictyostelium.
Surprisingly, the KEGG pathways “Protein processing in ER”, “protein export” and “proteasome”
were also among the most significantly represented pathways. However, most of the proteins matched
to the ERAD part of the “Protein processing in ER” pathways. These pathways are actually involved
in the degradation of misfolded proteins. It is possible that the machinery and the products of this
degradation could be targeted to phagosomes by autophagy explaining the presence of these pathways
in the phagosome proteome. Alternatively, originally, this pathway now associated with the ER only,
might have contributed to the digestive function of the phagosome precursor organelle in primitive
eukaryotes (Cavalier-Smith 2009). The “protein processing to ER” pathway also reflected the
contribution of the ER membrane to the phagosomal membrane (Desjardins, Houde et al. 2005).
83 Table III: Number and percentage of Dictyostelium phagosomal proteins matched in GO-terms and KEGG
pathways (Dieckmann, Gueho et al. 2012). Significantly represented GO-terms (p-value < 10-5, enrichment > 2)
and significantly represented KEGG pathways (p-value < 10-4) are listed.
By comparing the enriched GO-terms and the significantly represented KEGG pathways of
the phagosome and of the early M. marinum-containing compartment, these two compartments did not
appear drastically different. Indeed, both of them have GO-terms and KEGG pathways related to the
phagosomal and endocytic pathways. Furthermore, their enriched GO-terms reflected the basic
function of the phagosome such as actin cytoskeleton reorganisation, vesicle trafficking or bacteria
killing. KEGG metabolic pathways reflecting the nutritive function of phagosomes were also found
for both compartments. GO-terms indicating the presence of transcriptional machinery in the M.
marinum-containing compartment and the detection of the KEGG pathway “protein processing in ER”
for both M. marinum-containing compartment and phagosome confirmed that both of them contain
cellular components which are targeted for degradation, probably by autophagy. Interestingly, the
KEGG pathway “phosphatidylinositol signalling system” was only found in the M. marinumcontaining compartment. This confirmed the interest in further investigating the membrane
composition of this compartment. The comparison of phagosomes and early M. marinum-containing
compartments using GO-terms and KEGG pathways did not reveal major qualitative differences.
However, the use of quantitative proteomic will help to find more subtle differences.
3.2.3-Proteomic analysis of the temporal modification of the M. marinum-containing
compartment during the early steps of infection
In order to study the temporal modifications of the M. marinum-containing compartment during the
early phase of infection, all the proteins identified in the mix containing M. marinum-containing
compartments isolated at 1 hpi, 3 hpi and 6 hpi were quantified with the help of the IsoQuant module
of the Easyprot platform (Gluck, Hoogland et al. 2013). Both M. marinum-containing compartments
isolated at 3 hpi and 6 hpi were compared to M. marinum-containing compartments isolated at 1 hpi
84 (Table I). Protein ratios were obtained for the two comparisons performed. Ratios thresholds of 0.67fold and 1.5-fold were applied to select only strongly decreased or enriched proteins on M. marinumcontaining compartments at 3 or 6 hpi compared to M. marinum-containing compartment at 1 hpi. Of
the 1461 proteins identified, 161 were found decreased at 3 hpi and 24 were enriched. Not
surprisingly, more differences were found between M. marinum-containing compartments isolated at 1
hpi and the ones isolated at 6 hpi. 207 proteins were decreased at 6 hpi, and 40 were enriched. 132
proteins were found commonly decreased at 3 hpi and 6 hpi and only 4 proteins were found commonly
enriched. To find pathways potentially excluded from the M. marinum-containing compartment
maturation or pathways potentially involved in the establishment of a compartment permissive for M.
marinum replication, the proteins commonly found differentially abundant at 3 hpi and 6 hpi were
matched to GO-terms and KEGG pathways. P-values of 0.05 and GO-term enrichment score of 3 were
applied to select significantly represented GO-terms and KEGG pathways. Among the 132 proteins
decreasing from the M. marinum-containing compartment during its maturation, 113 had GO-terms
annotations and 129 matched in KEGG pathways (Table IV). The GO-term “profilin binding” was
highly enriched. The proteins with this GO-term annotation were dynamin A, Formin-I and Formin-B.
All are actin-binding proteins, and formins are involved in actin polymerisation. The decrease of these
proteins from the M. marinum-containing compartment during its maturation could indicate that the
actin coat around the compartment slowly dissociates. However, confirming the results of a recent
study, this dissociation is much slower than for a normal phagosome, which looses its actin coat
within a minute after closure (Kolonko et al., in revision at Cell Micro). This actin coat would prevent
the fusion of the M. marinum-containing compartment with late acidic endosomes. The GO-terms
“lipid binding” and “enzyme regulator activity” were also found enriched among the proteins
decreasing during M. marinum-containing compartment maturation. It was not surprising that several
proteins were annotated with both of these GO-terms. Indeed, the GO-term “enzyme regulator
activity” regrouped regulators of small GTPases. These proteins are involved in intracellular
trafficking and usually bind to membranes. It is interesting that the amount of these regulators of small
GTPases decreases on the M. marinum-containing compartment during maturation. Indeed, this could
reflect a limited interaction of this compartment with other endosomal compartments of the cell.
Surprisingly, the GO-term “cis-trans isomerase activity” was found enriched among the proteins
decreasing on M. marinum-containing compartment during maturation. The eukaryotic cis-trans
isomerases are involved in different processes in the cell. They are notably regulating the cell cycle,
trafficking, signal transduction and transcription, stabilizing protein complexes and have an antioxidant activity (Min, Fulton et al. 2005, Shaw 2007). Any of these processes could play a role during
M. marinum infection. Finally, the GO-term “hydrogen ion transmembrane transporter activity” was
found enriched among the proteins decreasing in the M. marinum-containing compartment during
maturation. This would reflect a decreased acidification of the compartment. Interestingly, one protein
associated with this GO-term was the H+-vATPase subunit G, which is clearly involved in phagosome
acidification. Several other H+-vATPase subunits were actually found significantly decreased (pvalue<0.05) in the M. marinum-containing compartment during maturation. But, their ratios were too
high to pass the threshold applied. However, this clearly shows that the H+-vATPase is transiently
present in the M. marinum-containing compartment as demonstrated by a previous study (Hagedorn
and Soldati 2007). The other three of the four proteins annotated with this GO-term were actually
85 mitochondrial. However, some of them were also identified on the micropinosome suggesting that
they also have an endosomal localisation (Journet, Klein et al. 2012).
Six KEGG pathways with a p-value below 0.05 were found significantly represented among
the proteins decreasing in the M. marinum-containing compartment during maturation (Table IV).
Only 2 to 5 proteins were matched in the different pathways, which was probably too low to really
consider the pathways as highly represented. However, with 4 matched proteins, the “endocytosis”
pathway can be mentioned. The five other pathways were related to DNA replication DNA mismatch
repair and transcription. The presence of these proteins in phagosomal compartments might be
explained by fusion with autophagosomes. The decreasing abundance of these proteins in the M.
marinum-containing compartment during maturation probably reflects a limited interaction of this
compartment with autophagosomes.
GO Terms
KEGG pathways
DNA
M. marinum-containing
compartment
Whole cell
P-value
Enrichment
3 (2.65%)
4 (3.54%)
4 (3.54%)
9 (7.96%)
13 (11.50%)
13 (0.16%)
25 (0.30%)
40 (0.49%)
161 (1.95%)
250 (3.03%)
0.0128
0.0074
0.0345
0.0074
0.0021
16.84
11.67
7.3
4.08
3.79
M. marinum-containing
compartment
Whole cell
P-value
4 of 129
5 of 129
2 of 129
2 of 129
2 of 129
2 of 129
39 of 13310
89 of 13310
18 of 13310
24 of 13310
32 of 13310
34 of 13310
0.00061
0.00185
0.01348
0.02307
0.03903
0.04352
Table IV: Number and percentage of the proteins decreasing on the maturing M. marinum-containing
compartment matched in GO-terms and KEGG pathways. Proteins with ratios < 0.66 were matched to GO-terms
and KEGG pathways. Significantly represented GO-terms (p-value < 0.05, enrichment > 3) and KEGG pathways
(p-value < 0.05) are listed.
Among the 4 proteins increasing in the M. marinum-containing compartment during
maturation, all had a GO-term annotation and matched in KEGG pathways. However, due to the low
number of proteins, no KEGG pathways were found significantly represented. Two of the proteins had
a “hydrolase activity, acting on ester bonds” GO-term annotation. The 2 others had a “lipid binding”
GO-term annotation. Proteins with this annotation are usually associated to membranes. One of these
2 proteins was the Phox-domain containing protein DDB_G0289833 that appeared to be a homologue
of the human Sorting-nexin 4 (SNX4). Interestingly, several Sorting nexins are involved in
intracellular trafficking. The human SNX4 is involved in the recycling of the transferrin receptor from
early endosomes to the ERC (Leprince, Le Scolan et al. 2003, Traer, Rutherford et al. 2007, Skanland,
Walchli et al. 2009). From different studies, different SNX4 binding partners were identified: KIBRA,
86 dynein, tubulin, amphiphisin 2 and indirectly Dynamin, which is a binding partner of amphyphisin 2.
Interestingly, these partners are also present in the Dictyostelium genome and were also identified in
the early M. marinum-containing compartment proteome (Table V). Moreover, they were found
differentially abundant (p-value<0.05) on the M. marinum-containing compartment during its
maturation. With ratios close to 1, Dynein and Tubulin were not strongly differentially abundant.
However, DwwA which is the KIBRA homologue in Dictyostelium, a BAR-domain containing
protein, which is similar to amphyphisin 2, and dynamin were significantly decreased in the M.
marinum-containing compartment during maturation. It is worth noting that these 3 SNX4 partners
were decreasing whereas SNX4 was accumulating in the maturing M. marinum-containing
compartment. Interestingly, a recent study demonstrated that Legionella, an other intraphagosomal
pathogen, is able to inhibit retrograde trafficking from its containing vacuole. This action is actually
essential to promote intracellular replication (Finsel, Ragaz et al. 2013). In a similar way, M. marinum
could block cargo sorting from its compartment, resulting in the accumulation of SNX4 on the M.
marinum-containing compartment, whereas the SNX4 binding partners are not recruited. To test this
hypothesis, the role of SNX4 and of its binding partners in M. marinum infection will have to be
further investigated.
Mammal proteins
SNX4
Dictyostelium proteins
Phox domain-containing protein;
DDB number
DDB_G0289833
M. marinum-containing comparment
3 hpi/1 hpi
6 hpi/1 hpi
1.85
2.06
KIBRA
Dictyostelium WW domain-containing protein
DDB_G0281827
0.55
0.48
Dynein
Dynein heavy chain
DDB_G0276355
1.13
1.13
Tubulin
tubulin beta chain
DDB_G0269196
1.11
0.95
Tubulin
tubulin alpha chain
DDB_G0287689
1.13
NS
amphyphisin 2
BAR domain-containing protein
DDB_G0288895
0.66
0.73
Dynamin
DynaminA
DDB_G0277849
0.59
0.46
Table V: Protein ratios of SNX4 and its binding partners in the maturing M. marinum-containing compartment
The matching with GO-terms and KEGG pathways of proteins, which accumulate or decrease
in the M. marinum-containing compartment during its maturation, helped to highlight different
proteins, family of proteins or pathways, which might play a role during the establishment of a
compartment permissive for M. marinum replication. The role of these potential candidates will have
to be investigated more deeply. However, to differentiate among these candidates, proteins
representing general phagosomal functions from proteins specifically involved in the establishement
of the M. marinum-containing compartment, the manipulated M. marinum-containing compartment
was also compared to non or less-manipulated compartments in a second quantitative proteomic
analysis.
87 3.2.4-Quantitative proteomic comparison of non-manipulated phagosomes and
phagosomes manipulated by M. marinum
In order to identify potential M. marinum-containing compartment markers, and potential pathways
targeted by the pathogenic strain to divert the phagosomal pathway into the establishment of a
compartment permissive for mycobacterial replication, the M. marinum-containing compartment and
compartments containing avirulent (M. marinum-L1D), non pathogenic (M. smegmatis) or attenuated
(M. marinumΔRD1) mycobacteria were isolated at 1 hpi and 6 hpi. They were then subjected to
reduction, alkylation, trypsin digestion and finally isobaric labelling using TMT. The TMT sixplex kit
allows to analyse up to six samples at a time. Each time, biological duplicates of three different
mycobacteria strain-containing compartments were analysed. After TMT labelling, equal amounts of
the six samples were mixed. The mixed peptides were then separated in 12 or 24 fractions by OGE
and each fraction was analysed by LC-MS/MS. Three different experiments were performed in which
322, 1234 and 1023 Dictyostelium proteins were identified (Table VI). For the first experiment, the
mixed peptides were only separated in 12 fractions, explaining the lower number of identified proteins
(322) compared to the other 2 experiments. For each experiment, all the identified proteins were
quantified with the help of the IsoQuant module of the Easyprot platform (Gluck, Hoogland et al.
2013). Protein ratios were obtained for each comparisons performed. Ratios thresholds of 0.67-fold
and 1.5-fold were applied to select only strongly decreased or enriched proteins on the M. marinumcontaining compartment compared to compartments containing avirulent, non-pathogenic or
attenuated mycobacteria strains.
Number of proteins
Compartments isolated at 1 hpi
M. marinum
M. marinum
M. marinum
vs
vs
vs
M. marinum-L1D
M. smegmatis
M. marinum-RD1
Compartments isolated at 6 hpi
M. marinum
M. marinum
vs
vs
M. smegmatis
M. marinum-RD1
322
322
1234
1023
1023
Ennriched on M. marinumcontaining compartment
19
22
275
60
99
Decreased on M. marinumcontaining compartment
56
79
209
425
103
Table VI: Quantitative proteomic comparison of M. marinum-containing compartment with compartments
containing avirulent, non-pathogenic or attenuated mycobacteria strains. Number of identified proteins in each
experiments and numbers of proteins with ratios < 0.66 or ratios > 1.5 in each quantitative proteomic
comparisons performed.
M. marinumΔRD1 lacks the RD1 locus encoding for ESAT6 and CFP10. These proteins are
involved in phagosome escape of the mycobacteria and infection dissemination to other cells by
ejection of the mycobacteria from their infected cell. These events occur at a late time of infection, at
least 24 hpi. Before that, M. marinumΔRD1 is supposed to establish a quasi-normal infection, similar
to M. marinum wt. However, surprisingly, the quantitative proteomic analysis revealed numerous
differences between compartments containing M. marinum or M. marinumΔRD1, already at early
88 times of infection (1 hpi and 6 hpi). Indeed, at 1hpi, 209 proteins were decreased and 275 proteins
were enriched in M. marinum-containing compartments, and at 6 hpi, 103 proteins were decreased and
99 were enriched in M. marinum-containing compartments (Table VI). In contrast, numerous
differences were expected between M. marinum-containing compartments and compartments
containing M. marinum-L1D or M. smegmatis. The comparison of M. marinum-containing
compartments with M. smegmatis-containing compartments revealed 79 proteins decreased and 22
proteins enriched in M. marinum-containing-compartments at 1 hpi, and 425 proteins decreased and
60 proteins enriched in M. marinum-containing compartments at 6 hpi (Table VI). In the comparison
of M. marinum-containing compartment with M. marinum-L1D-containing compartment, 56 proteins
were found decreased, and 19 proteins were found enriched in M. marinum-containing compartment at
1 hpi (Table VI).
Interestingly, the proteins found differentially abundant in the M. marinum-containing
compartment correspond to protein categories, which were also found decreasing or accumulating in
the M. marinum-containing compartment during the 6 first hours of its maturation. Numerous small
GTPases and GTPases regulators were differentially abundant on the M. marinum-containing
compartment (Table VII). Not all the small GTPases ratios passed the strong thresholds applied.
However, they almost all indicated that these proteins were significantly (p-value<0.005) less
abundant on the M. marinum-containing compartment, with small exceptions when the comparison
was done with compartments containing the attenuated strain M. marinumΔRD1. The small GTPases
are involved in membrane trafficking. Their decreased abundance in the M. marinum-containing
compartment clearly indicates that this compartment poorly interacts with other intracellular vesicles.
Furthermore, as described for M. tuberculosis, the late endosomal marker Rab7a was less abundant in
M. marinum-containing compartments (Via 1997). However, unlike results described for M.
tuberculosis (Via 1997, Clemens, Lee et al. 2000, Fratti, Backer et al. 2001, Kelley and Schorey 2003,
Rohde, Yates et al. 2007), the early endosomal marker Rab5 was not enriched on M. marinumcontaining compartments. However, in Dictyostelium, Rab5 delivery to phagosomes is very transient
(Caroline Barisch, unpublished results). The M. marinum-containing compartment potentially stops its
maturation at a slightly later stage, when Rab5 is already gone.
89 ID
Q54TP9_DICDI
RAB14_DICDI
RB11A_DICDI
GACC_DICDI
ARL8_DICDI
RAB1A_DICDI
RB32A_DICDI
RAB7A_DICDI
Description
Arf GTPase activating protein;
Ras-related protein Rab-14
Ras-related protein Rab-11A
Rho GTPase-activating protein gacC
ADP-ribosylation factor-like protein 8
Ras-related protein Rab-1A
Ras-related protein Rab-32A
Ras-related protein Rab-7A
RapGAP/RanGAP domain-containing
Q54Q75_DICDI
protein;
RASC_DICDI
Ras-like protein rasC
RASG_DICDI
Ras-like protein rasG
RACE_DICDI
Rho-related protein racE
RASS_DICDI
Ras-like protein rasS
RAB4_DICDI
Ras-related protein Rab-4
RAPA_DICDI
Ras-related protein rapA
RB11C_DICDI Ras-related protein Rab-11C
RASB_DICDI
Ras-like protein rasB
RB32D_DICDI Ras-related protein Rab-32D
RAB6_DICDI
Ras-related protein Rab-6
RAB1D_DICDI Ras-related protein Rab-1D
RAB2B_DICDI Ras-related protein Rab-2B
RAB2A_DICDI Ras-related protein Rab-2A
RAB8A_DICDI Ras-related protein Rab-8A
Rap-GAP domain-containing protein
Y1809_DICDI
DDB_G0281809
Ras GTPase-activating-like protein
RGAA_DICDI
rgaA (DGAP1)
ARF1_DICDI
ADP-ribosylation factor 1
GACJJ_DICDI Rho GTPase-activating protein gacJJ
RABC_DICDI
Ras-related protein RabC
RAB6_DICDI
Ras-related protein Rab-6
Q54L90_DICDI RasGEF domain-containing protein;
RAB18_DICDI Ras-related protein Rab-18
RAB5B_DICDI Putative ras-related protein Rab-5B
RB32C_DICDI Ras-related protein Rab-32C
RAB21_DICDI Ras-related protein Rab-21
RABQ_DICDI
Ras-related protein RabQ
RACC_DICDI
Rho-related protein racC
RAB1C_DICDI Ras-related protein Rab-1C
RAC1A_DICDI Rho-related protein rac1A
Rac guanine nucleotide exchange
GXCJJ_DICDI
factor JJ (RacGEF JJ)
RAB5A_DICDI Ras-related protein Rab-5A
Circularly permutated Ras protein 2
CPAS2_DICDI
(DdiCPRas2)
Q8SSP5_DICDI Arf GTPase activating protein;
Probable serine/threonine-protein
ROCO5_DICDI
kinase roco5
Regulator of chromosome
Q55CR5_DICDI condensation domain-containing
protein;
RABG2_DICDI Ras-related protein RabG2
RAB1B_DICDI Ras-related protein Rab-1B
RACB_DICDI
Rho-related protein racB
Ras guanine nucleotide exchange
GEFO_DICDI
factor O
RB32B_DICDI Ras-related protein Rab-32B
Ras guanine nucleotide exchange
GEFP_DICDI
factor P
RABT2_DICDI Ras-related protein RabT2
RABG2_DICDI Ras-related protein RabG2
RapGAP/RanGAP domain-containing
Q54P30_DICDI
protein;
Compartments isolated at 1 hpi
M. marinum
M. marinum
M. marinum
vs
vs
vs
M. marinum-L1D M. smegmatis M. marinum-RD1
0.29
1.22
0.40
0.51
0.69
0.40
0.56
0.53
0.41
0.51
0.44
0.76
1.08
0.47
0.82
0.57
0.51
0.70
0.80
0.54
0.88
0.58
0.64
1.50
1.98
1.63
0.66
0.40
0.43
0.41
0.46
0.47
0.47
0.49
0.53
0.61
0.62
0.63
0.66
0.66
Compartments isolated at 6 hpi
M. marinum
M. marinum
vs
vs
M. smegmatis M. marinum-RD1
0.53
0.81
0.54
0.76
0.46
0.46
0.75
0.51
0.80
0.51
0.82
0.50
0.77
0.50
0.74
0.76
1.07
1.27
0.57
0.57
0.77
0.78
0.69
0.74
1.19
0.82
0.56
0.47
0.76
0.54
0.72
0.76
0.81
0.75
0.51
0.53
1.72
1.82
1.91
1.94
0.86
1.12
1.50
1.29
0.58
0.62
0.64
2.53
0.70
0.78
0.76
0.78
0.50
0.50
0.51
0.51
0.51
0.55
0.56
0.59
0.85
0.72
0.69
0.82
1.21
0.74
0.68
0.77
0.61
0.63
0.71
0.65
0.65
1.34
0.57
0.52
0.50
1.95
3.03
1.76
0.73
0.80
1.24
0.96
1.08
0.73
0.68
0.69
0.69
0.72
0.74
0.79
0.88
Table VII: Small GTPases and GTPases regulators differentially abundant in M. marinum-containing
compartments
90 Almost all the H+-vATPase subunits were found differentially abundant. They were
commonly decreased in the M. marinum-containing compartment in all the comparisons performed
(Table VIII). This suggests a limited acidification of the M. marinum-containing compartment as
described for the M. tuberculosis-containing compartment (Sturgill-Koszycki, Schlesinger et al. 1994,
Mwandumba, Russell et al. 2004, Pethe, Swenson et al. 2004).
ID
Description
vatA
vatB
vatC
vatD
vatE
vatF
vatG
vatH
vatL
vatM
vATPase subunit
Compartments isolated at 1 hpi
M. marinum
M. marinum
M. marinum
vs
vs
vs
M. marinum-L1D
M. smegmatis
M. marinum-RD1
0.32
0.57
0.55
0.50
0.85
0.62
1.25
0.39
0.66
0.42
Compartments isolated at 6 hpi
M. marinum
M. marinum
vs
vs
M. smegmatis
M. marinum-RD1
0.67
0.67
0.77
0.77
0.70
0.60
0.71
0.68
0.65
1.18
0.63
0.78
0.58
0.69
1.21
0.47
0.48
0.79
1.20
0.67
Table VIII: H+-vATPase subunits differentially abundant in M. marinum-containing compartments
Numerous actin-binding proteins were found differentially abundant on the M. marinumcontaining compartment. A list of proteins with strong ratios was constituted. The list was then
completed with some significant ratios values (p-value<0.05), which originally did not pass the strong
thresholds applied (Table IX). Previous studies showed that the M. tuberculosis-containing
compartment retains the actin-binding protein Coronin longer than a normal phagosome (Ferrari,
Langen et al. 1999, Fratti, Vergne et al. 2000, Deghmane, Soualhine et al. 2007). However, according
to the ratios obtained, Coronin A was decreased in the M. marinum-containing compartment. This
result is actually consistent with Schuller et al. study (Schuller, Neefjes et al. 2001). They show that
coronin was involved in the uptake of BCG but was then disscociated from the BCG-containing
compartment in 1 hour. Furthermore, a recent study showed that the actin coat is retained around M.
marinum-containing compartments (Kolonko et al., in revision at Cell Micro). Surprisingly, the ratios
obtained did not show a common enrichment of all the actin-binding proteins on the compartment
containing M. marinum. This result can be explained by the timing. Indeed, at 1 hpi, compartments
containing M. marinum-L1D are at the end of the phagosomal maturation pathway and close to
exocytosis. At that step, they re-acquire an actin coat. Similarly, some M. smegmatis-containing
compartments can be ready for exocytosis at 1 hpi. However, the maturation of these compartments is
slower than for M. marinum-L1D-containing compartments (Hagedorn and Soldati 2007). Most of the
M. smegmatis-containing compartments re-acquire an actin coat at 6 hpi, before exocytosis. The reacquisition of an actin coat by these compartments could explain why several actin-binding proteins
were found less abundant on the M. marinum-containing compartment. Indeed, the actin-binding
proteins decreased in the M. marinum-containing compartment were mainly proteins involved in actin
polymerisation and actin cytoskeleton folding (Arp 2/3 complex, TCP1 subunit, Formin B), and
91 proteins anchoring the actin cytoskeleton to the plasma membrane (Ponticulins). As expected, strict
actin-binding proteins were enriched in the M. marinum-containing compartment (CARMIL, Actin
binding protein F, Myosin IC) confirming the importance of the actin coat during the first hours of M.
marinum infection (Kolonko et al., in revision at Cell Micro).
ID
Description
T-complex protein 1 subunit zeta
(TCP-1-zeta)
FORB_DICDI
Formin-B
T-complex protein 1 subunit
TCPG_DICDI
gamma (TCP-1-gamma)
PONC5_DICDI Ponticulin-like protein C5
ACTNA_DICDI Alpha-actinin A
TALA1_DICDI
Talin-A
MYOC_DICDI
Myosin IC heavy chain
CARML_DICDI Protein CARMIL (dDcarmil) (p116)
ABPF_DICDI
Actin-binding protein F
COROA_DICDI Coronin-A
TBB_DICDI
Tubulin beta chain
TBA_DICDI
Tubulin alpha chain
LIME_DICDI
LIM domain-containing protein E
Adenylyl cyclase-associated
CAP_DICDI
protein (CAP)
PONC3_DICDI Ponticulin-like protein C3
PONA_DICDI
Ponticulin
PONB_DICDI
Ponticulin-like protein B
T-complex protein 1 subunit alpha
TCPA_DICDI
(TCP-1-alpha)
LIMD_DICDI
LIM domain-containing protein D
ACT1_DICDI
Major actin
Probable myosin light chain kinase
MYLKG_DICDI
DDB_G0275057
MYOE_DICDI
Myosin IE heavy chain
FORA_DICDI
Formin-A
CTXB_DICDI
Cortexillin-2
CTXA_DICDI
Cortexillin-1
MYOG_DICDI Myosin-G heavy chain
MYOA_DICDI
Myosin IA heavy chain
LIMB_DICDI
LIM domain-containing protein B
Actin-related protein 2/3 complex
ARPC4_DICDI
subunit 4 (p20-ARC)
MLR_DICDI
Myosin regulatory light chain
Actin-related protein 2/3 complex
ARPC3_DICDI
subunit 3 (p21-ARC)
ARP2_DICDI
Actin-related protein 2
T-complex protein 1 subunit theta
TCPQ_DICDI
(TCP-1-theta)
Actin-related protein 2/3 complex
ARPC2_DICDI
subunit 2 (p34-ARC)
ARP3_DICDI
Actin-related protein 3
DYNA_DICDI
Dynamin-A
Actin-related protein 2/3 complex
ARPC1_DICDI
subunit 1
TALB_DICDI
Talin-B
MYOK_DICDI
Myosin-K heavy chain
AP2A2_DICDI AP-2 complex subunit alpha-2
TCPZ_DICDI
Compartments isolated at 1 hpi
M. marinum
M. marinum
M. marinum
vs
vs
vs
M. marinum-L1D M. smegmatis M. marinum-RD1
0.34
0.76
0.46
0.66
0.47
0.48
0.57
1.54
1.55
1.66
1.78
0.77
0.53
1.10
1.58
1.08
0.54
1.51
1.76
2.16
1.63
0.58
Compartments isolated at 6 hpi
M. marinum
M. marinum
vs
vs
M. smegmatis M. marinum-RD1
0.88
1.22
1.14
0.81
0.66
0.67
0.87
1.09
0.67
0.74
0.51
0.93
1.42
1.36
1.28
1.51
0.62
1.53
0.82
0.69
1.27
0.63
0.86
0.81
0.59
0.65
0.69
0.79
1.20
0.70
0.41
0.47
0.52
0.86
0.31
0.55
0.57
0.59
0.73
0.63
0.65
1.69
1.86
2.01
0.50
0.63
1.34
1.34
1.25
1.67
2.22
2.48
0.83
0.90
0.64
0.88
0.47
0.47
0.49
0.55
0.77
0.89
0.56
0.57
0.95
1.27
0.58
0.59
0.61
1.31
1.16
0.71
1.23
0.65
0.65
0.65
Table IX: Actin-binding proteins differentially abundant in M. marinum-containing compartments
92 It has been shown that lipid metabolism is particularly important for M. tuberculosis
intraphagosomal growth (McKinney, Honer zu Bentrup et al. 2000, Munoz-Elias and McKinney
2005). Furthermore, the host cell lipid metabolism is deregulated, and lipid droplets containing
cholesterol accumulate in the cytosol of infected cells (D'Avila, Melo et al. 2006, Peyron,
Vaubourgeix et al. 2008). Interestingly, several proteins involved in lipid metabolism were found
differentially abundant (Table X). A list of these differentially abundant proteins was assembled.
Then, it was completed with some proteins with significant ratios (p-value<0.05), which did not pass
the first strong thresholds applied. However, these proteins were mainly decreased in the M. marinumcontaining compartment. During M. marinum infection, proteins involved in lipid metabolism could
be relocalised from the M. marinum-containing compartment to future lipid droplets to favour the
production and storage of lipids. Interestingly, at 1 hpi, proteins involved in lipid metabolism were
increased in M. marinum-containing compartment compared to M. marinumΔRD1-containing
compartments. As previously explained, M. marinumΔRD1 is supposed to establish a compartment
similar to M. marinum-containing compartment. 1 hpi is a very early stage of the infection. This
difference potentially indicates a small delay or an ineficiency at relocalisating the proteins involved
in lipid metabolism and consequently, a delay in the manipulation of its compartment by M.
marinumΔRD1 compared to M. marinum.
ID
CP51_DICDI
DGAT2_DICDI
SMT1_DICDI
OSB8_DICDI
Q553T0_DICDI
PLC_DICDI
Description
Probable lanosterol 14-alpha demethylase
(LDM)
Diacylglycerol O-acyltransferase 2
Probable cycloartenol-C-24methyltransferase 1
Oxysterol-binding protein 8
Putative uncharacterized protein;
1-phosphatidylinositol-4,5-bisphosphate
phosphodiesterase (PLC)
Phosphatidylinositol-3,4,5-trisphosphate 3-
Compartments isolated at 1 hpi
M. marinum
M. marinum
M. marinum
vs
vs
vs
M. marinum-L1D M. smegmatis M. marinum-RD1
Compartments isolated at 6 hpi
M. marinum
M. marinum
vs
vs
M. smegmatis M. marinum-RD1
0.34
0.39
0.66
0.39
1.62
0.86
0.57
0.74
0.50
0.39
0.42
0.75
0.58
1.52
0.65
1.53
1.10
1.75
0.71
0.67
0.48
0.47
0.52
1.54
2.94
2.11
0.60
0.58
0.54
0.41
0.83
0.57
2.87
0.63
0.50
1.81
PTEN_DICDI
SAC1_DICDI
FCSB_DICDI
Q75JL8_DICDI
Q8MMS1_DICDI
Q86IY4_DICDI
ERG24_DICDI
OSB7_DICDI
Y2012_DICDI
Q54PA2_DICDI
Q553T0_DICDI
Q556T4_DICDI
Q54UU9_DICDI
Q75JX1_DICDI
OSB6_DICDI
CAS1_DICDI
ERG2_DICDI
FCSA_DICDI
phosphatase PTEN
Phosphatidylinositide phosphatase SAC1
Fatty acyl-CoA synthetase B (LC-FACS 2)
Acyl-CoA oxidase;
Acyl-CoA oxidase;
Acetyl-CoA C-acyltransferase;
Delta(14)-sterol reductase
Oxysterol-binding protein 7
Putative elongation of fatty acids protein
DDB_G0272012
Putative uncharacterized protein;
Putative uncharacterized protein;
Delta 9 fatty acid desaturase;
Phosphatidylinositol 3-kinase;
Sphingosine kinase related protein;
Oxysterol-binding protein 6
Cycloartenol synthase
Protein erg2 homolog
Fatty acyl-CoA synthetase A (LC-FACS 1)
1.62
1.76
1.85
1.79
1.85
1.96
2.07
2.30
0.65
0.71
1.14
2.35
1.57
1.75
2.03
1.13
0.59
0.64
2.05
1.54
0.78
0.73
0.83
0.44
1.32
0.52
0.55
0.47
0.36
0.58
0.58
0.59
Table X: Proteins involved in lipid metabolism, differentially abundant in M. marinum-containing compartment
93 Probably due to the low number of proteins identified in the comparisons M. marinum/M.
marinum-L1D and M. marinum/M. smegmatis at 1 hpi, only few proteins involved in fusion or in
sorting were found differentially abundant. However, for the 3 other comparisons, numerous proteins
of the fusion machinery and proteins involved in sorting were identified and found differentially
abundant. These proteins were commonly decreased in the M. marinum-containing compartment in all
the comparisons (Table XI). This clearly confirms that the M. marinum-containing compartment
poorly interacts with other intracellular compartments. Not only does it not fuse with other
compartments but surprisingly, the sorting from this compartment also seems to be impaired. This
result, again, is in agreement with a recent study, which showed that blocking the retrograde
trafficking from Legionella-containing vacuole is necessary to promote intraphagosomal growth of the
pathogen (Finsel, Ragaz et al. 2013).
ID
Description
VPS35_DICDI Vacuolar sorting protein 35
Vacuolar protein sorting-associated
VPS26_DICDI
protein 26
Vacuolar protein sorting-associated
VPS29_DICDI
protein 29
Vacuolar protein sorting-associated
VPS4_DICDI
protein 4
Putative vacuolar protein sortingVP13A_DICDI
associated protein 13A
Putative vacuolar protein sortingVP13C_DICDI
associated protein 13C
Vacuolar protein sorting-associated
VPS45_DICDI
protein 45
NSF_DICDI
Vesicle-fusing ATPase
Gamma-soluble NSF attachment
SNAG_DICDI
protein (SNAP-gamma)
Alpha-soluble NSF attachment
SNAA_DICDI
protein (SNAP-alpha)
Vesicle transport through interaction
VTI1A_DICDI
with t-SNAREs homolog 1A
AP1M_DICDI AP-1 complex subunit mu
AP1G_DICDI AP-1 complex subunit gamma
AP2M_DICDI AP-2 complex subunit mu
AP1S2_DICDI AP-1 complex subunit sigma-2
AP2A2_DICDI AP-2 complex subunit alpha-2
AP1B_DICDI AP-1 complex subunit beta
STX7A_DICDI Syntaxin-7A
STX7B_DICDI Probable syntaxin-7B
Q54ZH1_DICDI t-SNARE family protein;
Q551H5_DICDI t-SNARE family protein;
Compartments isolated at 1 hpi
M. marinum
M. marinum
M. marinum
vs
vs
vs
M. marinum-L1D M. smegmatis M. marinum-RD1
0.71
0.44
Compartments isolated at 6 hpi
M. marinum
M. marinum
vs
vs
M. smegmatis M. marinum-RD1
0.63
0.45
0.37
0.56
0.37
0.66
0.91
0.72
0.25
0.59
0.69
0.68
0.72
0.51
0.82
0.53
0.46
0.69
0.55
0.63
0.49
0.63
0.49
0.51
0.53
0.58
0.61
0.71
0.67
0.64
0.77
0.46
0.64
0.65
0.66
0.56
0.57
0.53
0.70
0.51
Table XI: Proteins involved in fusion or sorting differentially abundant in M. marinum-containing compartment
As expected, late phagosomal markers and proteins involved in killing and digestion of
ingested particles are decreased in the M. marinum-containing compartment (Table XII). This again
indicates a maturation arrest or diversion of the M. marinum-containing compartment. The decreased
amount of lysosomal enzymes confirms the establishment of a compartment with a less hostile
environment for the mycobacteria.
94 ID
Description
Membrane-associated
sulfotransferase kil1
Q54F16_DICDI Cysteine protease;
VACB_DICDI Vacuolin-B
VACA_DICDI Vacuolin-A
CYSP4_DICDI Cysteine proteinase 4
P80_DICDI
Protein P80
KIL1_DICDI
Compartments isolated at 1 hpi
M. marinum
M. marinum
M. marinum
vs
vs
vs
Compartments isolated at 6 hpi
M. marinum
M. marinum
vs
vs
M. marinum-L1D
M. smegmatis
M. marinum-RD1
M. smegmatis
M. marinum-RD1
0.39
0.63
1.67
0.61
0.69
0.72
0.96
0.84
1.50
0.55
0.62
0.51
0.77
0.67
0.57
0.63
0.46
0.81
Table XII: Late phagosomal markers differentially abundant in M. marinum-containing compartment
As SNX4 appeared as one of the highest accumulated proteins on the maturing M. marinumcontaining compartment, protein ratios for SNX4 and its partners in the different proteomic
comparisons were listed (Table XIII). Both SNX4 and its partners, except the Dictyostelium KIBRA
homologue, were identified in the different comparisons. Surprisingly, at 1 hpi, SNX4 and its partners
were not found differentially abundant when the M. marinum-containing compartment was compared
to M. marinum-L1D or M. smegmatis-containing compartments. Only the tubulin appeared
differentially abundant. However, opposite ratio values were obtained in the 2 comparisons and it was
difficult to interpret. SNX4 was only found enriched in the M. marinum-containing compartment when
compared to M. marinumΔRD1-containing compartments. Some of the SNX4 partners were also
differentially abundant. This could again indicate a small delay between M. marinum and M.
marinumΔRD1 in the establishment of a compartment permissive for replication. However, at 6 hpi,
SNX4 was decreased in M. marinum-containing compartment compared to M. smegmatis-containing
compartment. SNX4 partners were also found decreased. It was surprising that this protein, which
strongly accumulates in the maturing M. marinum-containing compartment, was less abundant in the
M. marinum-containing compartment compared to the M. smegmatis-containing compartment. This
showed that even if this protein strongly accumulates in the M. marinum-containing compartment, it
accumulates even more in the M. smegmatis-containing compartment. However, this difference was
only observed at 6 hpi. SNX4 potentially accumulates similarly in both M. marinum and M.
smegmatis-containing compartments during the very first hours of infection, and then, this
accumulation slows down in the M. marinum-containing compartment when it starts to be highly
manipulated by the pathogenic mycobacteria. As SNX4 is involved in sorting events in mammals, its
decreased amount and the decreased amount of its partners in the M. marinum-containing
compartment again highlight an impaired sorting from this compartment. The blocking of sorting
events from compartments containing the pathogenic strain M. marinum will have to be further
investigated.
95 Compartments isolated at 1 hpi
Mammal
proteins
Dictyostelium proteins
DDB number
M. marinum/
M. smegmatis
SNX4
Phox domain-containing protein;
DDB_G0289833
KIBRA
Dictyostelium WW domain-containing
protein
DDB_G0281827
Dynein
Dynein heavy chain
DDB_G0276355
Tubulin
tubulin beta chain
DDB_G0269196
Tubulin
tubulin alpha chain
DDB_G0287689
amphyphisin 2 BAR domain-containing protein
DynaminA
Dynamin
M. marinum/
M.
M. marinum-L1D marinum RD1
M. marinum/
M. marinum/
M. smegmatis
M. marinum RD1
NS
1.52
0.86
1.24
NS
NS
1.25
0.68
NS
NS
1.51
NS
0.65
0.86
0.77
1.76
0.69
0.69
0.81
NS
0.72
NS
NS
0.59
1.27
NS
DDB_G0288895
DDB_G0277849
Compartments isolated at 6 hpi
M. marinum/
NS
NS
Table XIII: SNX4 and its binding partners.
Overall, both the proteomic analysis of the temporal modification of the M. marinumcontaining compartment and the quantitative proteomic comparison of this compartment with
compartments containing avirulent, non pathogenic or attenuated mycobacteria strains confirmed that
the M. marinum-containing compartment does not follow the normal phagosome maturation pathway.
M. marinum stops its maturation and avoids the establishment of a hostile environment. Indeed, this
compartment acquires less late phagosomal markers (Vacuolin), less proteins involved in acidification
(H+-vATPase subunits) and digestion (lysosomal enzymes). The maturation of a normal phagosome
implies fusion and fission events. M. marinum seems to block the maturation of its containingcompartment by limiting its interaction with other compartments. Indeed, the decreased abundance of
small GTPases and their regulators, of fusion machinery and of proteins involved in sorting, notably
SNX4 and its partners, clearly indicate that the M. marinum-containing compartment poorly interacts
with other intracellular compartments of the endocytic and secretory pathways. Interestingly, the
importance of the actin coat around the M. marinum-containing compartment during the first hours of
infection was also highlighted. Furthermore, the results confirmed the implication of the lipid
metabolism in the M. marinum infection. We decided to focus on 2 different aspects involved in the
establishment of a compartment allowing M. marinum replication. First, since a previous study
showed that the H+-vATPase is transiently delivered to the M. marinum-containing compartment
(Hagedorn and Soldati 2007), we initiated the further investigation of H+-vATPase retrieval from this
compartment. Then, we plan to study the modulation of cargo sorting from M. marinum-containing
compartments.
3.3-The H+-vATPase is retrieved from M. marinum-containing compartments in a WASHindependent manner during the early phase of infection
The different quantitative proteomic comparisons performed indicated that the H+-vATPase is
depleted from the M. marinum-containing compartment at both 1 and 6 hpi. Furthermore, they also
showed that the abundance of the H+-vATPase in the M. marinum-containing compartment decreases
during the 6 first hours of its maturation. However, a previous study showed that the H+-vATPase is
transiently delivered to the M. marinum-containing compartment. To better understand why the H+vATPase is depleted from the M. marinum-containing compartment, we decided to study H+-vATPase
96 retrieval mechanisms from mycobacteria-containing compartments. Since the WASH complex has
been shown to play a crucial role in the H+-vATPase retrieval from phagosomes, we decided first, to
confirm its retrieval function from phagosomes and then, to investigate its function during
mycobacteria infection.
3.3.1-WASH, a key factor for the H+-vATPase retrieval from phagosomes
Carnell et al. showed that the WASH complex is involved in the retrieval of the H+-vATPase from
phagosomes (Carnell, Zech et al. 2011). Neutralised compartments were not found in cells lacking
WshA. Furthermore, WASH is a nucleation-promoting factor. Its recruitment to phagosomes leads to
the formation of an actin coat. It is proposed that actin polymerisation induces budding of small
vesicles. By binding to the H+-vATPase, actin would recruit it to microdomains that finally bud into
vesicles, thereby inducing its retrieval. In collaboration with Robert Insall’s lab, the role of WshA and
of the WASH complex in phagosome reneutralisation was further investigated, especially in the
context of autophagy. WshA is part of a complex composed of 5 proteins: SWIP, Strumpellin, Fam21,
WshA and CCDC53 (Seaman, Gautreau et al. 2013). The role of Fam21, the hypothetical membranerecruiting subunit of the WASH-complex, was also investigated. In Dictysotelium, its role would be to
recycle the WASH complex from post-lysosomes to compartments earlier in the phagosomal pathway
(Park, Thomason et al. 2013). Latex-beads phagosomes from wt cells, wshA-null cells and fam21-null
cells were isolated after 3 hours of maturation. At this time, the wt phagosomes are at the end of the
maturation process, just before exocytosis, whereas the phagosomes from wshA-null cells and fam21null cells are blocked at two different subsequent steps of maturation, right before and right after H+vATPase retrieval (Park, Thomason et al. 2013). The phagosomes were subjected to reduction,
alkylation, trypsin digestion and finally isobaric labelling using TMT. After TMT labelling, equal
amounts of phagosomes from wt, wshA-null and fam21-null cells were mixed. The mixed peptides
were separated in 12 or 24 fractions by OGE and each fraction was analysed by LC-MS/MS. Two
independent experiments were performed in which 1073 and 951 proteins were identified. Each
identified proteins was then quantified with the help of the IsoQuant module of the Easyprot platform
(Gluck, Hoogland et al. 2013). Two comparisons were performed: phagosomes isolated from wshAnull cells and phagosomes isolated from fam21-null cells were compared to phagosomes isolated from
wt cells. Protein ratios were obtained for each comparison. Thresholds of 0.67-fold and 1.5-fold were
then applied to select proteins strongly enriched or strongly decreased in phagosomes isolated from
wshA-null cells or from fam21-null cells. A list a interesting proteins was assembled from the different
comparisosn. Then, it was completed with some ratios, which initially did not pass the strong
thresholds applied (Table XIV).
As expected, the results confirmed the role of WshA in the H+-vATPase retrieval. Indeed, in
cells lacking WshA, the H+-vATPase cannot be retrieve from phagosomes. This led to increased
amounts of all the H+-vATPase subunits on phagosomes isolated from WshA-null cells. On the
contrary, the H+-vATPase subunits were decreased in phagosomes isolated from Fam21-null cells.
Indeed, Fam21 is necessary for the recycling of the WASH complex. In its absence, the WASH
97 complex is hyperactive, stays on the phagosome and keeps retrieving the H+-vATPase. This induces
an even more complete and synchronous retrieval than in wt cells.
Interestingly, lysosomal enzymes and the lysosomal markers LmpA and LmpB were found
decreased in both phagosomes isolated from WshA-null cells and from Fam21-null cells. However,
lysosomal enzymes are supposed to be already delivered to phagosomes when the WASH complex is
recruited. This could indicate a trafficking defect in cells lacking a functionnal WASH complex.
Cells lacking either WshA or Fam21 are not able to complete the phagosome maturation. In
WshA-null cells, phagosomes are blocked before H+-vATPase retrieval and, in Fam21-null cells,
phagosomes are blocked after WASH complex recycling (Park, Thomason et al. 2013). Interestingly,
the results indicated that the phagosomes isolated from WshA-null cells and from Fam21-null cells
were at different maturation stages. Indeed, in the two comparisons performed, opposite ratios were
obtained for several small GTPases, proteins involved in fusion (SNARE, SNARE interacting protein
Vti1A, SNAP and NSF) and proteins involved in sorting (VPS13A and C, VPS45). These proteins
appeared mainly enriched on phagosomes isolated from WshA-null cells whereas they were decreased
in phagosomes isolated from Fam21-null cells. Among the small GTPases, there was Rab14, which is
involved in homotypic fusions after phagosome reneutralisation. The increased abundance of Rab14
and of proteins involved in fusion in phagosomes isolated from WshA-null cells could indicate that
phagosomes accumulate the machinery necessary for homotypic fusions which should occur after
reneutralisation. On the opposite, in Fam21-null cells, this machinery was decreased on the
phagosomes. Indeed, in Fam21-null cells, phagosomes are reneutralised and can undergo homotypic
fusion. However, they cannot recycle the WASH complex and cannot exocytose their content.
Blocked at this stage, they probably performe excessive homotypic fusion leading to the use of a large
amount of proteins involved in this process and the formation of huge post-lysosomes. Huge postlysosomes were indeed observed in Fam21-null cells (Park, Thomason et al. 2013).
Numerous other small GTPases and their regulators were found commonly enriched or
decreased on both phagosomes isolated from WshA-null cells and Fam21-null cells. As these proteins
are involved in endosomal trafficking, this confirms that this process is disturbed in cells lacking a
functionnal WASH complex.
Since the WASH complex is a nucleation-promoting factor, it was not surprising that
numerous actin binding proteins and proteins involved in actin cytoskeleton formation and folding
were found differentially abundant. Surprisingly, the results obtained showed mainly a decreased
amount of cytoskeleton proteins on both phagosomes isolated from WshA-null cells and from Fam21null cells. Indeed, it was shown that Fam21-null cells are able to polymerise actin around phagosomes
whereas WshA-null cells are not (Park, Thomason et al. 2013). However, this could simply reflects the
reaquisition of an actin coat by phagosomes in wt cells prior to exocytosis whereas phagosomes in
WshA-null cells and in Fam21-null cells are blocked before that stage.
98 WhsA-null phagosomes
ID
Description
Fam21 -null phagosomes
vs
vs
wt phagosomes
wt phagosomes
Experiment 1
Experiment 2
Experiment 1
1.52
1.12
0.89
0.68
1.56
1.22
1.36
0.74
V-type proton ATPase subunit C (V-ATPase subunit C)
1.25
1.28
V-type proton ATPase subunit d (V-ATPase subunit d)
1.68
1.11
VATD_DICDI
V-type proton ATPase subunit D (V-ATPase subunit D)
1.96
VATE_DICDI
V-type proton ATPase subunit E (V-ATPase subunit E)
1.42
1.31
VATF_DICDI
V-type proton ATPase subunit F (V-ATPase subunit F)
1.61
1.35
VATG_DICDI
V-type proton ATPase subunit G (V-ATPase subunit G)
1.24
1.40
0.87
VATH_DICDI
V-type proton ATPase subunit H (V-ATPase subunit H)
1.41
1.25
0.73
VATM_DICDI
Vacuolar proton translocating ATPase 100 kDa subunit
1.97
1.16
Q54R55_DICDI
Cathepsin Z;
0.44
CPVL_DICDI
Probable serine carboxypeptidase CPVL
0.44
CATD_DICDI
Cathepsin D
0.46
HEXA2_DICDI
Beta-hexosaminidase subunit A2
0.45
Q54F16_DICDI
Cysteine protease;
0.57
HEXA1_DICDI
Beta-hexosaminidase subunit A1
0.60
CYSP5_DICDI
Cysteine proteinase 5
CYSP4_DICDI
Cysteine proteinase 4
LMPB_DICDI
Lysosome membrane protein 2-B (LIMP II-2)
LYSG2_DICDI
Probable GH family 25 lysozyme 2
0.65
0.47
Q23857_DICDI
V
0.39
0.36
LMPA_DICDI
Lysosome membrane protein 2-A
0.76
PONB_DICDI
Ponticulin-like protein B [CHAIN 0]
CORO_DICDI
Coronin
0.47
0.56
MYOK_DICDI
Myosin-K heavy chain
0.61
0.44
ACT1_DICDI
Major actin [CHAIN 0]
0.62
0.73
CAPZB_DICDI
F-actin-capping protein subunit beta
0.64
VATB_DICDI
V-type proton ATPase catalytic subunit A (V-ATPase subunit
A)
V-type proton ATPase subunit B (V-ATPase subunit B)
VATC_DICDI
VA0D_DICDI
VATA_DICDI
Experiment 2
0.77
0.54
0.88
0.72
0.76
0.78
0.64
0.82
0.37
0.88
0.45
0.23
1.26
0.45
0.81
0.51
0.59
0.54
0.64
0.64
0.68
0.40
0.66
0.15
COF1_DICDI
0.40
0.93
0.56
0.72
MYLKD_DICDI
Probable myosin light chain kinase DDB_G0292624
MYOC_DICDI
Myosin IC heavy chain
0.76
0.69
ARPC1_DICDI
Actin-related protein 2/3 complex subunit 1
0.70
ARP3_DICDI
Actin-related protein 3
0.73
TALB_DICDI
Talin-B
MYOJ_DICDI
Myosin-J heavy chain
CAPZA_DICDI
F-actin-capping protein subunit alpha
MLR_DICDI
Myosin regulatory light chain
0.53
MLE_DICDI
Myosin, essential light chain
0.57
MYS2_DICDI
Myosin-2 heavy chain
0.59
TBA_DICDI
Tubulin alpha chain
2.46
0.41
0.44
0.72
0.73
0.51
0.82
0.76
0.59
0.75
0.64
0.86
0.64
3.16
3.19
0.56
3.01
0.65
2.84
PROF1_DICDI
CAP_DICDI
Adenylyl cyclase-associated protein (CAP)
0.51
1.73
SEVE_DICDI
Severin
0.84
0.69
1.42
0.66
FORI_DICDI
Formin-I
1.29
1.49
1.44
FORB_DICDI
Formin-B
0.87
TCPG_DICDI
T-complex protein 1 subunit gamma (TCP-1-gamma)
2.32
1.27
TCPZ_DICDI
T-complex protein 1 subunit zeta (TCP-1-zeta)
1.77
1.16
ACTNA_DICDI
Alpha-actinin A
LIME_DICDI
LIM domain-containing protein E
MYOE_DICDI
Myosin IE heavy chain
CARML_DICDI
Protein CARMIL (dDcarmil) (p116)
1.60
MYOG_DICDI
Myosin-G heavy chain
2.32
PROF2_DICDI
1.49
0.79
0.60
1.58
0.77
0.64
2.35
0.86
0.82
0.52
0.80
1.51
99 WhsA-null phagosomes
Fam21 -null phagosomes
vs
vs
wt phagosomes
wt phagosomes
ID
Description
Experiment 1
Q551H5_DICDI
3.32
NSF_DICDI
t-SNARE family protein;
Vesicle transport through interaction with t-SNAREs homolog
1A
Vesicle-fusing ATPase
SNAG_DICDI
Gamma-soluble NSF attachment protein (SNAP-gamma)
SNAA_DICDI
RAB14_DICDI
Experiment 2
Experiment 1
Experiment 2
0.71
0.80
1.17
0.49
0.78
1.27
1.18
0.45
0.72
1.39
1.09
0.51
Alpha-soluble NSF attachment protein (SNAP-alpha)
2.15
1.11
0.56
0.67
Ras-related protein Rab-14
1.76
1.25
0.61
0.81
Q54TP9_DICDI
Arf GTPase activating protein;
1.57
1.76
RAC1B_DICDI
Rho-related protein rac1B
1.61
Q54Q75_DICDI
RapGAP/RanGAP domain-containing protein;
1.63
RAB4_DICDI
Ras-related protein Rab-4
1.71
RB11A_DICDI
Ras-related protein Rab-11A
2.19
ARF1_DICDI
ADP-ribosylation factor 1 [CHAIN 0]
Q869V0_DICDI
RapGAP/RanGAP domain-containing protein;
RRAGA_DICDI
Ras-related GTP-binding protein A
0.44
0.65
RABQ_DICDI
Ras-related protein RabQ
1.34
0.56
0.71
RAB1C_DICDI
Ras-related protein Rab-1C [CHAIN 0]
1.28
0.57
0.72
RASC_DICDI
Ras-like protein rasC [CHAIN 0]
0.64
0.71
RAB1A_DICDI
Ras-related protein Rab-1A
0.58
0.38
Q8SSP5_DICDI
Arf GTPase activating protein;
RB11C_DICDI
Ras-related protein Rab-11C
0.88
0.40
0.78
KXCB_DICDI
Kinase and exchange factor for Rac B
1.17
1.39
2.65
RACD_DICDI
Rho-related protein racD
RGAP1_DICDI
RapA guanosine triphosphatase-activating protein 1
GACEE_DICDI
Rho GTPase-activating protein gacEE
1.28
GACHH_DICDI
Rho GTPase-activating protein gacHH
1.93
Q54DK9_DICDI
Arf GTPase activating protein;
2.12
GEFR_DICDI
Ras guanine nucleotide exchange factor R
1.53
Q54PK6_DICDI
Ras-related GTP-binding protein;
RAB6_DICDI
Ras-related protein Rab-6
0.91
RB32D_DICDI
Ras-related protein Rab-32D
0.81
0.54
0.64
RAB1D_DICDI
Ras-related protein Rab-1D
1.32
0.95
0.56
0.72
RAB8A_DICDI
Ras-related protein Rab-8A
0.80
0.57
0.71
RB32B_DICDI
Ras-related protein Rab-32B
1.11
0.85
0.59
0.74
RAPA_DICDI
Ras-related protein rapA [CHAIN 0]
RABG2_DICDI
Ras-related protein RabG2
1.15
ARL8_DICDI
ADP-ribosylation factor-like protein 8
0.86
1.05
0.61
RAB5A_DICDI
Ras-related protein Rab-5A
0.84
1.12
0.64
RB32A_DICDI
Ras-related protein Rab-32A
GEFV_DICDI
RASG_DICDI
Ras guanine nucleotide exchange factor V
RasGEF domain-containing serine/threonine-protein kinase
X
Ras-like protein rasG [CHAIN 0]
GXCJJ_DICDI
Rac guanine nucleotide exchange factor JJ (RacGEF JJ)
0.81
GACN_DICDI
Rho GTPase-activating protein gacN
1.07
VP13A_DICDI
Putative vacuolar protein sorting-associated protein 13A
1.49
1.72
0.86
1.07
VPS45_DICDI
Vacuolar protein sorting-associated protein 45
2.91
1.15
0.48
0.77
VP13C_DICDI
Putative vacuolar protein sorting-associated protein 13C
1.72
1.23
VTI1A_DICDI
GEFX_DICDI
0.88
1.21
0.45
0.93
0.81
0.76
1.33
1.15
2.35
0.54
1.44
1.38
0.61
2.54
0.76
0.76
1.23
2.35
2.00
3.11
0.33
0.46
0.60
0.61
0.68
0.64
0.73
0.54
2.47
3.50
1.11
0.93
0.68
0.65
0.57
3.08
0.82
Table XIV: Proteins differentially abundant on phagosomes isolated from WshA-null cells and Fam21-null cells.
H+-vATPase subunits (red), late endosomal markers and lysosomal enzymes (orange), cytoskeleton proteins
(blue), small GTPases and fusion machinery (green), sorting machinery (black).
To confirm the quantitative proteomic results and the role of WshA in the retrieval of the H+vATPase from phagosomes, the pH and the proteolytic activity of WshA-null cells phagosomes were
monitored. Latex-beads linked to fluorescent reporters were centrifuged on top of a cell monolayer to
synchronise their ingestion. The fluorescence of the reporters was then measured during 3 hours. It
100 should be noted that the reporter used for pH measurement is the FITC. It is perfect to trace kinetics
and pH variation between neutral and 5 but not lower. The lowest pH values measured might be even
lower. In wt cells, phagosomes rapidly acidified (Figure 6A). The pH reached the lowest value of 4.99
in 38 minutes. Then, phagosomes reneutralised to reach a pH of 6.20 at 130 minutes. In WshA-null
cells, phagosomes acidified slower and less efficiently. They reached only a pH value of 5.16 at 56
minutes after bead ingestion. Then, they barely reneutralised and failed to reach the same pH as in wt
phagosomes. This reneutralisation defect clearly demonstrates that WshA is involved in this process.
This result and the quantitative proteomic results, together demonstrate that WshA is necessary to
retrieve the H+-vATPase from phagosomes and that this retrieval then allows the reneutralisation of
the phagosomal pH. The proteolysis profiles were consistent with the acidification profiles (Figure
6B).
Indeed, the hydrolytic activity of lysosomal enzymes is, at least partly, dependent on the
phagosomal pH. In wt cells, the phagosomal proteolytic activity started at 20 minutes, when the
phagosomal pH reached a value of 5.30. In WshA-null cells, the phagosomal proteolytic activity
started later, at 30 minutes. Indeed, the WshA-null cells phagosomes acidified slower and reached a pH
of 5.40 at 30 minutes. Furthermore, the proteolytic activity was lower in WshA-null cells. This is
consistent with the quantitative proteomic results indicating a decreased amount of lysosomal enzymes
in WhsA-null cells phagosomes. Then, in wt cells, at 90 minutes, the phagosomal proteolytic activity
slowed down and finally reached a plateau indicating an arrest of the proteolytic activity. Indeed, at 90
minutes, phagosomes were in the reneutralisation phase. The phagosomal pH had already reached a
value of 5.8 and was not acidic enough for an optimal activity of lysosomal enzymes. In contrast, the
proteolytic activity in WshA-null cells phagosomes did not reach a plateau. Indeed, WshA-null cells
phagosomes failed to reneutralise and stayed at an optimal pH for lysosomal enzymes hydrolytic
activity.
A
B
7.0
Ratio-488/594
6.5
pH
8
AX2
Wash
6.0
5.5
5.0
4.5
0
50
100
Time-(minutes)
150
200
AX2
Wash
6
4
2
0
0
50
100
150
200
Time-(minutes)
Figure 6: WshA-null cells are defectives for phagosome reneutralisation. A. Phagosomes acidification profiles
for wild type and WshA-null cells. Cells were spinoculated with silica beads coupled to the pH-sensitive dye
FITC and the pH-insensitive dye Alexa 594. Fluorescences emitted by the two dyes were measured with the help
of a plate reader and ratios were calculated. Curves represent the mean and SEM of 7 measurements of 3
independent experiments. B. Phagosomal proteolysis profiles for wild type and WshA-null cells. Cells were
spinoculated with silica beads coupled to the self-quenched dye DQ-Green via BSA and to the reporter dye
101 Alexa 594. Digestion of the BSA allows dequenching of the DQ-Green and the increase of fluorescence.
Fluorescences emitted by the two dyes were measured with the help of a plate reader and ratios were calculated.
Curves represent the mean and SEM of 4 measurements of 2 independent experiments.
Part of the proteomic data was published in the paper “WASH is required for lysosomal
recycling and efficient autophagic and phagocytic digestion” from King et al., 2013 (King, Gueho et
al. 2013). It confirmed the trafficking defect of lysosomal enzymes in WshA-null cells. Indeed, despite
an increased amount of lysosomal enzymes and an increased proteolytic activity in the whole cell, the
amount of lysosomal enzymes and the proteolytic activity are reduced at the single phagosome level.
The following model has been proposed. Lysosomal enzymes are supposed to be recycled after
WASH complex retrieval. In WshA-null cells, the WASH complex is not retrieved from phagosomes
and lysosomal enzymes cannot be recycled. They get trapped in a dead-end compartment and are not
available for maturation of newly formed phagosomes
Figure 7: Model of the Dictyostelium endocytic pathway (King, Gueho et al. 2013). A. In presence of a
functional WASH complex. B. In absence of WshA.
102 3.3.2-WASH localises to mycobacteria-containing compartments
Its role in H+-vATPase retrieval from latex-beads phagosomes revealed WshA as a plausible candidate
to explain the decreased amount of H+-vATPase in M. marinum-containing compartments. In order to
study the role of WshA in the context of a mycobacterial infection, its localisation and recruitment
were investigated by microscopy on both live and fixed cells. Proteomic comparisons revealed that the
H+-vATPase subunits were already less abundant in the M. marinum-containing compartment at 1 hpi.
So, WshA-GFP expressing cells were observed at 1 hpi after infection with M. marinum-mCherry10
or M. smegmatis-DsRed. WshA-positive mycobacteria-containing compartments were detected both in
M. marinum and in M. smegmatis infected cells at 1 hpi (Figure 7A). The recruitement of WshA to
these mycobacteria-containing compartments was then followed by live microscopy. WshA
recruitment around mycobacteria-containing compartments was quantified. The quantification was
then normalised to the lowest fluorescence value measured (Figure 7B). An early wave of recruitment
of WshA to both M. marinum and M. smegmatis-containing compartments was observed during the
very first minutes after mycobacteria ingestion (Figure 7C). In both cases, this recruitment was
transient, WshA rapidly dissociated from mycobacteria-containing compartments and was totally
absent at 10 mpi. After this first phase, a second wave of recruitment started immediately in M.
smegmatis infected cells. M. smegmatis-containing compartments became strongly WshA positive at
25-30 mpi. This might correspond to the beginning of the reneutralisation phase and confirms the role
of WhsA in this process. However, this second wave of WshA recruitment was delayed in M.
marinum-infected cells and the maximum of WshA recruitment was reached at 70 mpi. Furthermore,
before reaching this maximum recruitment, smaller waves of transient recruitment of WshA were also
observed. This could indicate a balance, a tug-of-war, between the host response to infection and the
manipulation of the host by M. marinum. Indeed, before succeeding in phagosome maturation arrest,
several waves of H+-vATPase delivery to the M. marinum-containing compartment could occur. In
response, WshA could be recruited several times to retrieve the H+-vATPase and prevent the
acidification of the M. marinum-containing compartment.
103 M. marinum
1 hpi
Relative-fluorescence-intensity
B
A
M.#marinum#
M.#smegmatis
2.0
1.5
1.0
0.5
0
5
10
15
M. smegmatis
Relative-fluorescence-intensity
Time-(minutes)
M.#marinum#
M.#smegmatis
2.5
2.0
1.5
1.0
0.5
0
20
40
60
80
100
Time-(minutes)
C
M. marinum
0s
M. smegmatis
D
30 s
0s
60 s
90 s
90 s
120 s
120 s
60 mpi
M. marinum
M. smegmatis
30 mpi
30 s
60 s
Figure 8: WshA is recruited to mycobacteria-containing compartments. A. WshA-GFP expressing cells were
infected with M. marinum-mCherry10 (Blue) or M. smegmatis DsRed (Blue). At 1 hpi, cells were stained for the
H+-vATPase subunit VatM (Red) and for GFP (Green). DNA was visualised using DAPI (Grey). Both M.
marinum and M. smegmatis were detected in WshA-positive compartments. Scale bar, 1 µm. B. WshA-GFP
expressing cells were infected with M. marinum-mCherry10 or M. smegmatis DsRed and immediately processed
104 for live microscopy. The WshA-GFP fluorescence intensity around mycobacteria was quantified using ImageJ.
C. Snapshots from movies illustrating the early delivery of WshA to mycobacteria-containing comparments
(arrowheads). Scale bar, 5 µm. D. Snapshots from movies illustrating the second wave a WshA recruitement to
mycobacteria-containing compartments (arrowhead), delayed for M. marinum-containing compartments. Scale
bar, 5 µm.
3.3.3-WASH is not a susceptibility factor
It was hypothesised that absence of the WASH complex could lead to a defect in the retrieval of the
H+-vATPase from M. marinum-containing compartments. Consequently, the resulting acidification of
these compartments would harm or kill the mycobacteria and prevent the establishment of an
infection. In order to test the hypothetical function of the WASH complex during M. marinum
infection, WshA-null cells were infected with M. marinum-lux. The mycobacteria growth was then
followed by measuring the luminescence emitted by M. marinum-lux. Surprisingly, M. marinum grew
similarly in wt and in WshA-null cells indicating that WshA is not required for the establishment of a
normal infection (Figure 8A). To avoid a hostile environment in whsA-null cells phagosomes, M.
marinum might escape from their compartment to the cytosol at earlier times. In order to test whether
the growth was due to an earlier escape of the mycobacteria, wshA-null cells were infected with M.
marinum. An IF was performed to verify the presence of p80 or Vacuolin around the mycobacteria.
Both p80 and Vacuolin are late phagosomal markers and Vacuolin strongly accumulates on M.
marinum-containing compartment at late times of infection. At 21 hpi, the majority of the
mycobacteria were still observed in compartments in WshA-null cells (Figure 8B). However,
compared to wt cells, less vacuolin was detected on these compartments. Furthemore, no strong p80
positive M. marinum-containing compartments were detected. In the absence of WshA, M. marinum
seems to establish a compartment with a slightly modified composition. However, as shown by the
infection results, this compartment is also permissive for M. marinum replication.
105 Luminescence,fold,increase,(RLU)
A
6
AX2
Wash
4
2
0
0
10
20
30
40
50
Time,(hours,post,infection)
B
AX2 cells
WshA-null cells
21 hpi
Figure 9: WshA-null cells are not resistant to M. marinum infection. A. Wild type and WshA-null cells were
infected with M. marinum-lux. At indicated time points, the mycobacterial growth was monitored by measuring
luminescence. Luminescence values were normalised to the value measured at 0.5 hpi. The curve represents the
mean fold increase of luminescence and SEM from 4 independent experiments. B. Wild type and WshA-null
cells were infected with M. marinum-GFP (blue) and stained for the phagosomal marker p80 (red) and the niche
marker vacuolin (green) at 21 hpi. In wild type cells, M. marinum was in p80 and/or vacuolin positive
compartments. A majoriy of M. marinum were detected in vacuolin-positive compartments in WshA-null cells.
Strong p80 positive M. marinum-containing compartments were hardly detected in WshA-null cells. Scale bar,
10 µm.
3.3.4-The H+-vATPase retrieval from mycobacteria-containing compartments is WASHindependent
Infection results showed that the WASH complex is not necessary for the establishment of a M.
marinum infection. To test whether, like in phagosomes, WshA is really involved in the retrieval of
H+-vATPase from mycobacteria-containing compartments or if an other machinery has the same
106 function, the pH of these compartments was measured during the first hours of infection.
Mycobacteria were first labelled with 2 different dyes; the pH-sensitive dye FITC as a pH reporter and
the pH-insensitive dye TRITC as a reference. Labelled M. marinum were centrifuged on top of a
monolayer of wt cells or of WshA-null cells. The emission of the 2 dyes was then measured during 5
hours with the help of a plate reader. As a control, in parallel, wt cells and WshA-null cells were also
infected with labelled M. marinum-L1D. Indeed, M. marinum-L1D-containing compartments follow
the normal phagosome maturation pathway. As expected, in wt cells, M. marinum-L1D-containing
compartments were rapidly acidified (Figure 9A). This acidification phase was even so fast that it
could not be entirely measured for technical reason. After 20 minutes, the pH of the M. marinum-L1Dcontaining compartment reached the lowest value of 5.09 and started to reneutralise to reach a pH of
6.xx at 5 hpi. This reneutralisation was slower than for a latex bead-containing phagosome, which was
already reneutralised 130 minutes after ingestion of the bead (Figures 9A and 6). In contrast, M.
marinum-containing compartments in wt cells barely acidified. The pH only dropped to 6.07 at 15
minutes to reach back a pH of 6.2 at 90 minutes. This clearly demonstrates that M. marinum prevents
acidification of its containing-compartment, as shown for M. tuberculosis (Sturgill-Koszycki,
Schlesinger et al. 1994, Mwandumba, Russell et al. 2004, Pethe, Swenson et al. 2004). In WshA-null
cells, the pH of M. marinum-L1D-containing compartments and of M. marinum-containing
compartments behaved as in wt cells. Indeed, M. marinum-L1D-containing compartments reached the
lowest pH value of 5.05 at 15 minutes and then reneutralised to reach a pH of 6.07 at 5:45 hpi. The
only difference was that the reneutralisation was slightly slower than for wt cells. However, a
reneutralisation phase was indeed observed, demonstrating that WshA is not essential for the retrieval
of the H+-vATPase from mycobacteria-containing compartments. The slower reneutralisation could
indicate that WshA participates in this step but that one or several other proteins have a similar
function. As in wt cells, the pH of M. marinum-containing compartments in WshA-null cells hardly
dropped. It reached a value of 6.02 at 10 mpi, and then reneutralised to reach a pH of 6.20 at 4 hpi.
Again, this reneutralisation is slower than in wt cells.
(&'
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Figure 10: WshA is not involved in mycobacteria-containing compartments reneutralisation. Mycobacteriacontaining compartments acidification profiles for wild type and WshA-null cells. Cells were spinoculated with
M. marinum or M. marinum-L1D labelled with the pH sensitive dye FITC and the pH insensitive dye TRITC.
Fluorescences emitted by the two dyes were measured with the help of a plate reader and ratios were calculated.
Curves represent the mean and SEM of 3 to 4 measurements of 2 independent experiments.
107 108 4-Discussion
109 The aim of this thesis was to understand how M. marinum manipulates the maturation of the
compartment where it resides. Studies performed on the M. tuberculosis-containing compartment
revealed only few qualitative characteristics of this compartment, based on the presence or absence of
different phagosomal markers. This compartment was shown to retain some early endosomal markers
such as coronin (Ferrari, Langen et al. 1999, Fratti, Vergne et al. 2000, Deghmane, Soualhine et al.
2007) or rab5 (Via 1997, Clemens, Lee et al. 2000, Fratti, Backer et al. 2001, Kelley and Schorey
2003, Rohde, Yates et al. 2007) whereas late phagosomal markers such as Rab7 or the H+-vATPase
(Sturgill-Koszycki, Schlesinger et al. 1994, Mwandumba, Russell et al. 2004, Pethe, Swenson et al.
2004) and lysosomal markers (Xu, Cooper et al. 1994, Sturgill-Koszycki, Schaible et al. 1996, Malik,
Iyer et al. 2001) were excluded. However, the full proteomic composition and the quantitative and
dynamic modification of the mycobacteria-containing compartment during the very early stage of
infection are poorly known.
By establishing a procedure to isolate mycobacteria-containing compartments, we were able to
purify compartments containing various mycobacteria strains at informative times during the early
phase of infection. This allowed us to compare the proteome of the isolated compartments using
quantitative proteomic. Thus, we managed to observe the qualitative and quantitative modifications
occurring to the M. marinum-containing compartment during the 6 first hours of infection and we able
to compare and contrast the composition of the M. marinum-manipulated compartment with non or
less-manipulated phagosomal compartments.
4.1-The M. marinum-containing compartment, a phagosomal compartment
The analysis of the proteome of the early M. marinum-containing compartment did not reveal major
differences from a standard phagosome in terms of composition. In both M. marinum-containing
compartments and phagosomes, the identified proteins represent basic functions of the phagosome
such as actin cytoskeleton organisation, vesicle trafficking or bacteria killing. However, some proteins
not directly related to phagosomal functions were also identified such as ribosomal proteins in both M.
marinum-containing compartments and in phagosomes, proteins of the translational machinery in M.
marinum-containing compartments, and proteins of the ERAD pathway in phagosomes. These
proteins could be targeted to phagosomes by autophagy to be degraded and would actually reflect the
degradative function of phagosomes. Interestingly, these proteins were also present in M. marinumcontaining compartments indicating that these compartments interact with the autophagosomal
pathway. It was shown that autophagic markers are recruited to mycobacteria-containing
compartments and that autophagy modulates mycobacterial survival (Gutierrez, Master et al. 2004,
Ponpuak, Davis et al. 2010, Castillo, Dekonenko et al. 2012). Autophagy would favour the fusion of
mycobacteria-containing compartments with phagolysosomes leading to the killing of mycobacteria.
Furthermore, it has been shown that ribosomal proteins and ubiquitin-derived peptides can be
processed into anti-mycobacterial compounds that can be delivered to mycobacteria-containing
compartments via autophagy (Alonso, Pethe et al. 2007, Ponpuak, Davis et al. 2010). In that sense, the
ribosomal proteins that we found in the M. marinum-containing compartment could reflect the host
defense response to infection. However, no proteins directly involved in autophagic function have
110 been identified in our M. marinum-containing compartment proteome. This proteome only represents
the M. marinum-containing compartment during the first six hours of infection and the autophagy
machinery could be recruited later. Furthermore, only three Atg proteins were identified in the
phagosomal proteome. This potentially indicates that autophagy proteins are hardly identifiable
probably because of a low abundance or of low solubilisation properties. It would be interesting to
further study the temporal recruitment of the autophagy machinery to mycobacteria-containing
compartments during infection. Furthermore, since autophagy seems to contribute to mycobacteria
killing in more than one way, by fusing mycobacteria-phagosomes with autophagolysosomes and by
delivering anti-mycobacterial compounds, the contribution of each of these pathways to mycobacteria
killing should be evaluated. The high number of proteins not directly related to phagosomal function
in our M. marinum-containing compartment proteome could represent a list of potential candidates
with anti-mycobacterial properties. The role of these proteins in mycobacterial infection should be
further investigated.
The major qualitative difference between M. marinum-containing compartments and
phagosomes was the significant representation of the KEGG pathway “phosphatidylinositol signalling
system” only in the M. marinum-containing compartment proteome. Consistent with this result, it is
known that PI3K is excluded from M. tuberculosis-containing compartments (Fratti, Backer et al.
2001, Chua and Deretic 2004, Hestvik, Hmama et al. 2005). PI3K is responsible for the
phosphorylation of PI(4,5)P2 present in the membrane of forming phagosomes to PI(3,4,5)P3. Its
exclusion from M. tuberculosis-containing compartments leads to a different lipidic composition of
the compartment. Furthermore, the mycobacterial protein SapM has been shown to directly
dephosphorylate PI(3,4,5)P3 (Vergne, Chua et al. 2005). Not only mycobacteria manipulate the
maturation of their containing-compartment by modifying their proteomic composition but they also
affect their membrane lipid composition. To better understand the manipulation mechanisms exerted
by mycobacteria on their containing-compartments, the proteomic analysis of these compartments
should be complemented with a lipidomic analysis. Furthermore, mycobacterial cell wall lipids have
been shown to modulate the maturation of the mycobacteria-containing compartment. However, their
mode of action is not known. It has been proposed that ManLAM incorporates into the membrane of
the mycobacteria-containing compartment. Lipidomic analysis of the mycobacteria-containing
compartment membrane would determine whether mycobacterial cell wall lipids incorporate into the
phagosomal membrane.
If the qualitative comparison of the M. marinum-containing compartment proteome with the
phagosome proteome did not reveal major differences, the quantitative proteomic comparison of M.
marinum-containing compartments with compartments containing avirulent, non pathogenic or
attenuated mycobacteria strains showed that M. marinum clearly affects the proteomic composition of
its compartment. Furthermore, our study confirms that M. marinum resides in a compartment with
much lower degradative properties than a non-manipulated phagosome, characterised by decreased
amounts of H+-vATPase and of lysosomal enzymes.
111 4.2-The M. marinum-containing compartment poorly interacts with other endosomal
compartments
Our proteomic analysis of mycobacteria-containing compartments clearly demonstrates that the M.
marinum-containing compartment poorly interacts with other endosomal compartments. Numerous
small GTPases were found to be less abundant on compartments containing the pathogenic strain M.
marinum compared to compartments containing the non-pathogenic strain M. smegmatis or the
avirulent or attenuated M. marinum strains. Furthermore, the fusion machinery was also found to be
decreased. GEF and GAP proteins were also at lower abundance in the M. marinum-containing
compartment during its maturation. The variety of these depleted proteins indicates that the fusion is
not selectively blocked with one kind of compartment, but that the overall fusion property of the M.
marinum-containing compartment is impaired. This result is not surprising since blocking the fusion
of the compartment seems to be the easiest way to prevent its maturation. However, it was surprising
that the sorting from M. marinum-containing compartments is also blocked. Indeed, intuitively, sorting
might appear as a way to remove from the compartment the proteins that are detrimental to the
bacteria. Interestingly, it was described that the retrograde transport is blocked during Legionella
infection. This block is even necessary to allow the intracellular growth of the bacteria (Finsel, Ragaz
et al. 2013). The authors also describe a decreased recruitment of several SNX proteins of the retromer
complex to the Legionella vacuole. Similarly, we see a decreased amount of SNX4 to the M.
marinum-containing compartment at 6 hpi. However, SNX4 is not associated with retromer function
(Hettema, Lewis et al. 2003). In mammals, it has been shown to localise to recycling endosomes and
to sort the transferrin receptor to the ERC (Traer, Rutherford et al. 2007, van Weering, Verkade et al.
2012). SNX4-binding partners have also been identified on the M. marinum-containing compartment.
As SNX4, they are depleted confirming that the sorting pathway regulated by SNX4 is really affected
during M. marinum infection. However, SNX4 binding partners were already decreased at 1 hpi
whereas SNX4 appeared decreased at 6 hpi. As described in Legionella infection (Finsel, Ragaz et al.
2013), a mycobacterial effector could directly act on SNX4 to prevent the recruitment of its partners.
The role of SNX4 during M. marinum infection will have to be further investigated with the help of a
knock-out strain. Furthermore, the conservation of its role in sorting will have to be evaluated in
Dictyostelium.
4.3-The role of lipids in mycobacteria infection
Our study clearly indicates that host lipid metabolism is affected during M. marinum infection.
Consistent with this, it was already shown that M. tuberculosis disturbs host lipid metabolism. It
induces the accumulation of cholesterol into cytosolic structures called lipid droplets (D'Avila, Melo et
al. 2006, Peyron, Vaubourgeix et al. 2008). The depletion of proteins involved in lipid metabolism
from the M. marinum-containing compartment could indicate a relocalisation of these proteins to
cytosolic lipid droplets. The accumulation of lipid droplets during mycobacterial infection should also
be confirmed in Dictyostelium. Furthermore, we can wonder how mycobacteria access the content of
these lipid droplets from inside their containing-compartment. The most plausible would be a fusion
even transient or partial of M. marinum-containing compartments with these lipid droplets. However,
112 our results indicate an abrogation of the overall fusion property of the M. marinum-containing
compartment. We could imagine that the M. marinum-containing compartment keeps fusion
machinery specifically involved in interaction with lipid droplet. An other possibility is the autophagy
of these lipid droplets and delivery of the autophagosome to the mycobacteria-containing
compartment. Mycobacteria access to lipid droplets content should definitely be investigated more
deeply
4.4-The WASH complex is not the key player of H+-vATPase retrieval during mycobacteria
infection
Our quantitative proteomic analysis revealed that the amount of H+-vATPase was decreasing on the
M. marinum-containing compartment during the 6 first hours of its maturation. This was confirmed by
the proteomic comparison of the M. marinum-containing compartment with compartments containing
avirulent, non-pathogenic or attenuated mycobacteria strains, which showed that the H+-vATPase was
indeed found commonly decreased in the M. marinum-containing compartment in all the comparisons
we performed. As a previous study in the lab indicated that the H+-vATPase is actually transiently
delivered to the M. marinum-containing compartment in the first 20 minutes of infection, we decided
to focus on the retrieval mechanism of the H+-vATPase (Hagedorn and Soldati 2007). The role of the
WASH complex in phagosome reneutralisation in Dictyostelium designated it as a good candidate to
study the H+-vATPase retrieval in mycobacterial infections (Carnell, Zech et al. 2011). We confirmed
the role of WshA in the retrieval of the H+-vATPase. Indeed, WshA-null cells were not able to
reneutralise their phagosomes and WshA-null cells phagosomes were enriched in H+-vATPase
compared to wt cells. We could show that the WASH complex was recruited to mycobacteriacontaining compartments. A first wave of recruitment was observed to both, M. marinum and M.
smegmatis-containing compartments during the first minutes after bacteria ingestion. The fact that it is
observed for both pathogenic and non-pathogenic mycobacteria strains indicates that this recruitment
might represent a basic feature of early phagosome maturation. Given its role in H+-vATPase retrieval,
WASH is associated to H+-vATPase positive compartments. Interestingly, the timing of this first wave
of recruitment corresponds to the time when H+-vATPase starts to be delivered to phagosomes. This
early WASH recruitment to the phagosomal compartment could simply reflect the fusion of the
phagosome with compartments carrying the H+-vATPase and still decorated with some patch of
WASH. Interestingly, the second wave of WASH recruitment was different for M. marinum and M.
smegmatis and could represent an event specific to phagosomes containing pathogenic mycobacteria.
In cells infected with the non-pathogenic strain M. smegmatis, the second wave of WASH recruitment
matches with the reneutralisation phase of the M. smegmatis-containing phagosome preceding
exocytosis. This is consistent with a canonical role of WASH in the reneutralisation process. However,
in cells infected with M. marinum, this second wave of recruitment was delayed, and was preceded by
smaller waves of WASH recruitment. We could imagine that these several waves of WASH
recruitment represent a balance between the host response to infection and the bacteria host
manipulation. Indeed, the host could try several times to deliver the H+-vATPase to the M. marinumcontaining compartment, while each time, the bacteria responds by inducing the recruitment of WASH
113 to the compartment to retrieve the H+-vATPase and prevent the acidification of its containingcompartment. These data were quite consistent with a role of WASH in the establishment of a
mycobacterial infection. However, the WshA-null cells infections with M. marinum performed did not
confirm this hypothesis. Indeed, M. marinum was able to grow in cells lacking WshA. Since WASH is
the major actor of phagosome reneutralisation, we were interested to monitor the pH experienced by
M. marinum during infection to better understand why WshA does not seem to be involved in
mycobacteria infection. Our results indicate that the pH of the M. marinum-containing compartment
stays quite neutral. However, the small pH drop and reneutralisation observed during the first hour of
infection indicate indeed some H+-vATPase delivery and retrieval. It was already shown that the
myocbacterial effector PtpA dephophorylates Vps33b to prevent HOPS assembly and H+-vATPase
delivery (Bach, Papavinasasundaram et al. 2008, Wong, Bach et al. 2011). We could imagine that the
barely modified pH in the M. marinum-containing compartment reflects a balance between the
decreased delivery and the retrieval of the H+-vATPase. Furthermore, other unidentified proteins could
also be involved in the H+-vATPase retrieval. This would explain why knock-out of WshA was not
sufficient to impact on M. marinum infection. Other proteins might compensate the absence of WshA.
Further investigations will be necessary to identify new proteins involved in H+-vATPase retrieval to
better understand the equilibrium between decreased H+-vATPase delivery and active retrieval.
Our phagosomal pH results obtained with mycobacteria also raised the question of the model
particles to use to study phagocytosis. Indeed, pH measurement of latex beads-phagosomes clearly
indicates that WshA and the WASH complex are the major actors of phagosome reneutralisation.
However, phagosomal pH measurement performed with the avirulent strain M. marinum-L1D
indicated that WASH was not or at least not the only actor of phagosome reneutralisation. Bacteria are
not inert particles. They have specific bacterial antigens, which are recognised by phagocytic cells,
and once ingested, some of them secrete proteins, which can interact with host proteins. Live bacteria
can clearly activate or interfere with different pathways of the host cells, whereas these pathways will
clearly not be activated with inert latex beads. Latex beads represent a good model to study the core
mechanisms involved in phagosome maturation. However, only bacteria will allow to understand the
entirety of the phagosomal pathway.
Overall, our results show that M. marinum affects the proteomic composition of its
containing-compartment, indicating that the maturation of this compartment is clearly affected.
Podinovskaia et al. have actually shown that the maturation of all the phagosomes of a cell infected
with M. tuberculosis is affected (Podinovskaia, Lee et al. 2013). This clearly indicates that pathogenic
mycobacteria do not only manipulate their containing-compartment from inside. They secrete
effectors, which access the cytosol and can affect the whole metabolismand homeostasis of the
infected cell. Furthermore, we could show that a variety of host pathways are affected: lipid
metabolism, phagosomal fusion, phagosomal sorting, actin cytoskeleton organisation. This indicates
that there must be many more secreted mycobacterial effectors than the few already identified. In the
future, the response of the mycobacteria to infection should be investigated in more depth, and notably
the secretome of the bacteria. During mycobacteria-containing compartments isolation, we did not
114 separate the isolated compartments from their containing-bacteria. This allowed us to also identify
mycobacterial proteins in addition to the host proteins during our proteomic analysis. This might
constitute a list of potential mycobacterial virulence factors and reflect the metabolic state of the
intraphagosomal mycobacteria. This list will be further dissected.
115 116 5-Appendix
117 5.1-Role of magnesium and phagosomal P-type ATPase in intracellular bacterial killing
Contributed Figure 6
118 Cellular Microbiology (2011) 13(2), 246–258
doi:10.1111/j.1462-5822.2010.01532.x
First published online 2 November 2010
Role of magnesium and a phagosomal P-type ATPase
in intracellular bacterial killing
Emmanuelle Lelong,1 Anna Marchetti,1
Aurélie Guého,2 Wanessa C. Lima,1
Natascha Sattler,2 Maëlle Molmeret,3
Monica Hagedorn,2† Thierry Soldati2 and
Pierre Cosson1*
1
Département de Physiologie Cellulaire et Métabolisme,
Faculté de Médecine de Genève, Centre Médical
Universitaire, CH1211 Geneva 4, Switzerland.
2
Département de Biochimie, Université de Genève,
CH1211 Geneva 4, Switzerland.
3
Université de Lyon1, IFR128, INSERM, U851,
Hospices Civils de Lyon, Faculté de Médecine Laënnec,
69007 Lyon, France.
Summary
Bacterial ingestion and killing by phagocytic cells
are essential processes to protect the human body
from infectious microorganisms. However, only
few proteins implicated in intracellular bacterial
killing have been identified to date. We used
Dictyostelium discoideum, a phagocytic bacterial
predator, to study intracellular killing. In a random
genetic screen we identified Kil2, a type V
P-ATPase as an essential element for efficient
intracellular killing of Klebsiella pneumoniae bacteria. Interestingly, kil2 knockout cells still killed
efficiently several other species of bacteria, and
did not show enhanced susceptibility to Mycobacterium marinum intracellular replication. Kil2 is
present in the phagosomal membrane, and its
structure suggests that it pumps cations into the
phagosomal lumen. The killing defect of kil2
knockout cells was rescued by the addition of
magnesium ions, suggesting that Kil2 may function as a magnesium pump. In agreement with this,
kil2 mutant cells exhibited a specific defect for
growth at high concentrations of magnesium. Phagosomal protease activity was lower in kil2 mutant
cells than in wild-type cells, a phenotype reversed
Received 13 August, 2010; revised 22 September, 2010; accepted 24
September, 2010. *For correspondence. E-mail pierre.cosson@
unige.ch; Tel. (+41) 22 379 5293; Fax (+41) 22 379 5338.
†
Present address: Bernhardt-Nocht Institute for Tropical Medicine,
Bernhardt-Noch-strasse 74, 20359 Hamburg, Germany.
by the addition of magnesium to the medium. Kil2
may act as a magnesium pump maintaining magnesium concentration in phagosomes, thus ensuring optimal activity of phagosomal proteases and
efficient killing of bacteria.
Introduction
Phagocytic cells are a key element of the immune system.
Monocytes and macrophages ingest and kill microorganisms, thus preventing the development of harmful infections (Segal, 2005). Phagocytosis of large particles
(typically > 0.5 mm) relies on the complex interplay of specific receptors at the cell surface, the actin cytoskeleton
and actin-binding proteins (Underhill and Ozinsky, 2002).
Following uptake, the next stages in the phagocytic
process involve notably changes in the protein composition (Garin et al., 2001; Haas, 2007) and ionic content
(Segal, 2005) of phagosomes (e.g. delivery of lysosomal
enzymes and acidification), as well as production of
superoxide ions (Sumimoto et al., 2005), culminating ultimately with the killing of the ingested microorganisms.
The capacity to escape or resist cellular killing mechanisms is a major virulence determinant for many pathogenic bacteria. Therefore, the understanding of cellular
killing mechanisms would greatly advance our understanding of host–pathogen interactions.
The mechanisms by which phagocytic cells kill internalized bacteria have been studied intensely in the last
decades (reviewed in Segal, 2005; Haas, 2007). Because
newly formed phagosomes were observed to progressively acidify and acquire lysosomal enzymes, it was initially proposed that lysosomal enzymes digest and kill
bacteria in the acidic environment of phagolysosomes.
The discovery of the critical role of the NADPH oxidase in
bacterial killing then led to the notion that superoxide and
other free radicals were the primary means by which
phagocytic cells kill bacteria. More recently, it has been
proposed that the main function of NADPH oxidase is to
regulate the ionic composition of the phagosome, and
to activate lysosomal enzymes (Reeves et al., 2002).
However, the ionic composition of maturing phagosomes
is still poorly characterized to date, as well as the importance of various ions in bacterial killing. It is also not
established how many distinct mechanisms exist for
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P-type ATPase, magnesium and intracellular killing
intracellular bacterial killing, and which ones are at play
for the killing of various types of bacteria. For both ethical
and technical reasons, mammals are not easily amenable
to genetic analysis, and no large-scale search for host
genes involved in intracellular bacterial killing has been
performed so far.
Dictyostelium discoideum amoebae are phagocytic
bacterial predators present in the soil. They can be easily
grown and manipulated, and their small, haploid, fully
sequenced and annotated genome allows simple genetic
analysis (Eichinger et al., 2005). They have been used
extensively to study the structure and dynamics of
the endocytic (Neuhaus et al., 2002) and phagocytic
(Gotthardt et al., 2006a) pathways, as well as the intracellular fate of pathogenic bacteria (Cosson and Soldati,
2008). Two Dictyostelium mutants defective for intracellular killing of Gram-negative Klebsiella pneumoniae bacteria (phg1a and kil1) have been described (Benghezal
et al., 2006), suggesting that this model system allows the
genetic analysis of intracellular killing mechanisms.
Phg1a is a member of the TM9 family of proteins, and Kil1
a sulfotransferase, and the direct or indirect role of these
two proteins in bacterial killing remains to be established.
In these two mutants, killing defects resulted in an inability
to use bacteria as nutrients, suggesting that new killingdefective mutants may be identified by screening for
mutants unable to grow on bacteria.
Here we describe the identification, in a random genetic
screen, of Dictyostelium kil2 mutant cells defective for
growth on Klebsiella bacteria. The kil2 gene encodes a
P-type ATPase essential for intracellular killing of Klebsiella. Analysis of kil2 knockout cells suggests a specific
role for magnesium in intracellular killing of Klebsiella.
247
Fig. 1. Isolation of kil2 mutant cells.
A. Dictyostelium mutants were obtained by restriction enzyme
mutagenesis insertion and screened for growth on Klebsiella.
Individual Dictyostelium clones were transferred onto a Klebsiella
lawn (black). Growing Dictyostelium cells create phagocytic plaques
(white) in the bacterial lawn. In this picture, two Dictyostelium
clones defective for growth on Klebsiella are visible (arrowheads).
B. In the kil2 restriction enzyme mutagenesis insertion mutant, the
mutagenic plasmid was inserted in the coding sequence of
DDB_G0279183 gene, 2121 nucleotides downstream from the start
codon.
C. The kil2 knockout mutant was obtained by deleting 1647
nucleotides in the kil2 coding sequence, and replacing them with a
blasticidin-resistance (BSR) cassette.
D. The Kil2 protein is not expressed in kil2 mutant cells.
Arrowhead indicates the Kil2 protein detected by Western blot. The
asterisk indicates a non-specific protein recognized by the
antiserum.
Results
kil2, an essential gene for growth on K. pneumoniae
To identify new genes involved in intracellular killing, we
performed a random insertional mutagenesis (Guerin and
Larochelle, 2002) and selected Dictyostelium mutants
growing normally in liquid HL5 medium, but defective for
growth on a lawn of Klebsiella (Fig. 1A). In the kil2 mutant,
the mutagenic plasmid was inserted in the coding
sequence of the DDB_G0279183 gene (hereafter named
kil2), 2121 nucleotides downstream from the start codon
(Fig. 1B). To confirm that the selective growth defect of the
kil2 mutant was due exclusively to the disruption of the kil2
gene, we deleted the kil2 coding sequence in the wild-type
DH1 strain by homologous recombination: 1647 nucleotides were deleted from position 798 to 2445 (Figs 1C and
S1). These cells no longer expressed the Kil2 protein
(Fig. 1D), exhibited a specific growth defect on Klebsiella
like the original kil2 mutant (see below) and were used to
further characterize the kil2 mutant phenotype.
The predicted Kil2 protein is composed of 1158 amino
acid residues, with 10 putative transmembrane domains.
It exhibits a strong similarity to members of the P-type
ATPase superfamily, which are involved in the active
transport of a variety of cations across membranes. The
five characteristic domains of P-type ATPases (Catty
et al., 1997; Axelsen and Palmgren, 1998) are notably
conserved in the Kil2 protein (Fig. 2A): the LTGES motif
(1) (position 295), the DKTGTLT phosphorylation domain
(2) (pos. 467), the ATP binding sites KGA(S)PE (3) (pos.
623) and ML(V)TGD (4) (pos. 717) and the GDGxND
hinge sequence (5) (pos. 857). In addition, Kil2 exhibits a
PPxxP motif (V) (pos. 424) and two cysteine residues
flanking the hinge sequence (Fig. 2B). These two features, as well as a long luminal loop between transmembrane domains 1 and 2 are typical of type V P-ATPases
(Figs 2 and S2). Type V family members are only found in
eukaryotes, and they form the most poorly characterized
family of P-type ATPases (see Discussion).
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248 E. Lelong et al.
range of other cellular functions. Phagocytosis of Klebsiella and of latex beads was unaffected in kil2 mutant
cells compared with wild-type cells (Fig. 4A) ruling out the
possibility that the kil2 growth defect might be due to
decreased bacterial internalization. In order to assess if
Fig. 2. The Kil2 protein is a type V P-ATPase.
A. The predicted topology of Kil2 shows 10 transmembrane
domains. Numbers 1 to 5 indicate the position of the five specific
motifs of P-type ATPases: (1) LTGES (2) DKTGTLT
(phosphorylation domain) (3) KGA(S)PE and (4) ML(V)TGD (ATP
binding sites) (5) GDGxND (hinge sequence). The PPxxP motif
specific for type V P-ATPases is also indicated (V).
B. Kil2 presents a high sequence similarity to two type V
P-ATPases: yeast YPK9 (YOR291w) and human AT132 (also
named ATP13A2). The presence of a PPxxP motif and of two
cysteine residues flanking the GDGxND hinge motif is also typical
of type V P-ATPases.
In order to characterize the phenotype of kil2 mutant
cells, we tested their ability to grow on a range of bacterial species. For this we applied increasing numbers
of Dictyostelium cells on a bacterial lawn to monitor
quantitatively their ability to form phagocytic plaques
(Alibaud et al., 2008; Froquet et al., 2009) (Fig. 3A). Kil2
mutants were defective for growth on Klebsiella, as well
as on a mucoid strain of Escherichia coli, but showed
no growth defect on many other Gram-negative or
Gram-positive bacteria, notably Bacillus subtilis or a nonvirulent Pseudomonas aeruginosa strain (Fig. 3A and B).
Interestingly, these results were very similar to those
seen with the previously characterized phg1a mutant
cells (Benghezal et al., 2006).
In addition, kil2 and phg1a mutants both grew on a lawn
of dead Klebsiella, killed either by heat inactivation or by
the addition of antibiotics (Fig. 3C). This result suggested
that both mutants do not grow on live Klebsiella because
they are unable to kill these bacteria efficiently.
Kil2 is essential for efficient intracellular killing
of Klebsiella
Defective growth on bacteria could conceivably be due to
defects in bacterial ingestion, in bacterial killing or in a
Fig. 3. kil2 mutant cells are specifically defective for growth on
Klebsiella.
A. In order to quantify the ability of Dictyostelium strains to grow
on bacteria, 10 000, 1000, 100 or 10 Dictyostelium cells were
applied onto a bacterial lawn (black). Wild-type Dictyostelium cells
created a phagocytic plaque (white). kil2 cells grew normally on B.
subtilis and on avirulent P. aeruginosa but presented an important
growth defect on Klebsiella.
B. Growth of kil2 cells on several bacterial species was tested as
described in A. Normal growth of Dictyostelium cells is indicated in
white, defective growth in black. Wild-type Dictyostelium grew on a
collection of Gram-negative and Gram-positive bacteria and did not
grow on virulent bacteria like wild-type P. aeruginosa. The kil2 and
phg1a mutants exhibited the same specific growth defects on
Klebsiella and on the mucoid E. coli B/r strain. (As: Aeromonas
salmonicida, Bs: Bacillus subtilis, Ec: Escherichia coli, Kp:
Klebsiella pneumoniae, Ml: Micrococcus luteus, Pa: Pseudomonas
aeruginosa.)
C. Normal growth of kil2 and phg1a cells was restored when
Klebsiella were boiled before the growth test or deposited on
SM-agar containing antibiotic (gentamycin 25 mg ml-1).
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P-type ATPase, magnesium and intracellular killing
249
Fig. 4. Inefficient intracellular killing of Klebsiella in kil2 mutant
cells.
A. Phagocytosis is not defective in kil2 mutant cells. Wild-type or
kil2 cells were incubated with fluorescent latex beads or
fluorescently labelled Klebsiella for 20 min. The cells were then
washed and the internalized fluorescence measured by flow
cytometry. The average and SEM of six independent experiments
are indicated.
B. Dictyostelium cells were incubated with Klebsiella and the
number of surviving bacteria (total or cell-associated) was
determined at different times by killing the Dictyostelium and plating
the bacteria on LB plates. Wild-type Dictyostelium cells killed
Klebsiella rapidly and very few viable cell-associated bacteria were
detected. On the contrary, kil2 mutant cells killed Klebsiella
inefficiently and a significant number of live intracellular bacteria
were detected in these cells. This experiment was repeated four
times with equivalent results.
C. Dictyostelium cells were incubated in the presence of
Klebsiella-GFP for 90 min, and then fixed. The endosomal p80
marker was revealed by immunofluorescence (white) and cells
observed by confocal microscopy. Live fluorescent Klebsiella
(green) were always observed inside p80-positive compartments
and accumulated more prominently in kil2 mutant cells (scale bar:
5 mm).
D. In cells analysed as described in C, the number of fluorescent
bacteria was quantified within 100 cells. Each bar indicates the
average and SEM of five independent experiments. The statistical
significance of these results was established using the Student’s
t-test (*P < 0.001).
kil2 cells killed efficiently bacteria, live Klebsiella were
incubated with Dictyostelium cells for up to 6 h. At 0, 1, 2,
4 and 6 h, the total number of remaining viable bacteria
was determined, as well as the number of live cellassociated bacteria. Wild-type Dictyostelium ingested and
killed bacteria rapidly (Fig. 4B). Intracellular killing in wildtype cells was so fast that viable intracellular bacteria
were hardly detectable at any time, suggesting that under
these experimental conditions the limiting factor for killing
was the rate of phagocytosis. On the contrary, kil2 mutant
cells killed Klebsiella slowly, and live intracellular bacteria
accumulated in these cells (Fig. 4B).
The fluorescence of GFP-expressing bacteria disappears rapidly when they are killed (Benghezal et al.,
2006). Therefore, to visualize the intracellular killing activity, we incubated GFP-expressing Klebsiella with Dictyostelium cells for 90 min. Cells were then fixed and
endosomal compartments visualized by immunofluorescence using a monoclonal antibody to endosomal p80
(Ravanel et al., 2001). In both wild-type and kil2 mutant
cells, live fluorescent bacteria were always observed in
phagosomal compartments delimited by a p80-positive
membrane (Ravanel et al., 2001) (Fig. 4C). However, two
times more fluorescent bacteria accumulated within kil2
mutant cells than within wild-type cells (Fig. 4C and D).
This result further suggested that intracellular killing of
Klebsiella was inhibited in kil2 mutant cells.
In order to assess the specificity of the killing defect
observed in kil2 mutant cells, we tested the ability of these
cells to kill other bacterial strains. Like phg1a mutant cells,
kil2 mutants killed Klebsiella more slowly than wild-type
cells did, but showed no defect in killing of B. subtilis and
P. aeruginosa (Fig. 5A). This result is in agreement with
the observation that kil2 mutant cells grew readily on the
latter two bacteria, and confirmed the specificity of the kil2
killing defect for Klebsiella.
In a previous study, Klebsiella mutants were selected
based on their ability to support growth of killing-defective
phg1a mutant cells (Benghezal et al., 2006). Two of these
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250 E. Lelong et al.
Fig. 5. The killing defect of kil2 mutant cells is specific for certain bacterial species. Dictyostelium cells were incubated with Klebsiella (Kp), B.
subtilis (Bs) or avirulent P. aeruginosa (Pa PT531) and the total number of viable bacteria was determined at the indicated times as described
in the legend to Fig. 4. The mean and SEM of three independent experiments are shown.
A. kil2 mutant cells killed inefficiently Klebsiella, but normally B. subtilis and P. aeruginosa. A similar phenotype was observed in phg1a mutant
cells.
B. kil2 mutant cells killed efficiently Klebsiella mutants with altered surface biosynthesis (waaQ and wbbM).
Klebsiella strains were mutated in genes involved in the
biosynthesis of the bacterial cell wall (waaQ and wbbM),
and they were more easily killed by phg1a mutant cells
than wild-type Klebsiella. Interestingly, these two Klebsiella strains were also more easily killed by kil2 mutant
cells (Fig. 5B), further reinforcing the resemblance
between the phenotypes of phg1a and kil2 mutants. This
result strongly suggests that the particular composition of
the Klebsiella surface allows them to withstand killing by
kil2 and phg1a mutant cells.
The killing defect exhibited by kil2 knockout cells could
be due to a gross defect in the organization and/or
dynamics of the endocytic/phagocytic pathway. Therefore, we assessed several key parameters of the
endocytic/phagocytic pathway. Macropinocytosis of fluid
phase containing a fluorescent dextran was measured
by flow cytometry, and was as efficient in kil2 cells as in
wild-type cells (110.27% ⫾ 5.83; n = 3). The pH of lysosomal and post-lysosomal compartments, as well as the
speed at which ingested fluid phase was transferred
from acidic lysosomes to more neutral post-lysosomes
was identical in wild-type and kil2 cells (Fig. S3). Quantitative immunofluorescence analysis during phagocytosis of latex beads revealed that the residency time of
beads in lysosomes (p80-positive, H+-ATPase-positive)
before their transfer to post-lysosomes (p80-positive,
H+-ATPase-negative) was also very similar in wild-type
and kil2 mutant cells (Fig. S4A). The activity of several
lysosomal enzymes was virtually identical in wild-type
and kil2 knockout cells (Fig. S4B). Finally, the size
and numbers of lysosomes and post-lysosomes were
determined after immunofluorescence staining, and were
very similar in both cell types (Table S1). Overall, these
results show that the killing defect of kil2 mutants is
not accompanied by a strong alteration in the general
organization or dynamics of the endocytic/phagocytic
pathway.
Co-purification of Kil2 with phagosomal membranes
Because our results implicated the Kil2 protein in bacterial
killing, we hypothesized that Kil2 localizes to phagosomes, the organelle in which killing is achieved. To test
this hypothesis, we allowed Dictyostelium cells to phagocytose latex beads, and purified on a sucrose gradient
latex beads containing phagosomes at various stages of
maturation (Gotthardt et al., 2006b; Dieckmann et al.,
2008). We then used specific antibodies to assess the
protein content of phagosomes. Kil2 was readily detectable in early phagosomes, and accumulated further in
maturing phagosomes (Fig. 6), a profile resembling that of
Fig. 6. Co-purification of the Kil2 protein with phagosomes. Cells
were incubated with latex beads during 5 or 15 min, washed to
eliminate non-internalized beads and incubated further for 15 or
45 min, as indicated. Phagosomes containing latex beads were
purified on sucrose gradients and analysed by Western blot in
order to detect the presence of different proteins: Kil2, endosomal
p80 and mitochondrial porin. Kil2 was detectable in very early
phagosomes and gradually accumulated during the maturation of
phagosomes. This pattern was extremely similar to that of
endosomal p80. Only trace amounts of mitochondrial porin were
detected in phagosomal fractions.
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P-type ATPase, magnesium and intracellular killing
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Fig. 7. Exogenous magnesium restores efficient killing in kil2 mutant cells.
A. Intracellular bacterial killing was assessed as described in the legend to Fig. 5, but with one of the following salts added to the extracellular
medium: MgCl2 (10 mM), CaCl2 (10 mM), NaCl (10 mM), KCl (10 mM), MnCl2 (10 mM), FeSO4 (10 mM), ZnCl2 (1 mM), NiCl2 (0.5 mM). The kil2
killing defect is the difference between the percentages of remaining bacteria in wild-type and kil2 cells, after 2 h of incubation. Exogenous
magnesium was the only ion abolishing the killing defect of kil2 mutant cells.
B. A typical killing experiment, showing the killing of Klebsiella by wild-type or kil2 knockout cells in the presence or absence of extracellular
MgCl2. In the presence of 10 mM MgCl2, efficient killing of Klebsiella by kil2 mutant cells was restored.
C. The percentage of remaining live bacteria after 2 h is shown for wild-type, kil2 and phg1a cells. Addition of 10 mM MgCl2 in the medium
restored the killing capacity of kil2 mutant cells whereas phg1a mutant cells were still unable to kill Klebsiella efficiently. It also slightly
accelerated killing in wild-type cells.
D. Killing was assessed as described above but in the presence of increasing concentrations of MgCl2 (0.01, 0.1, 1 or 10 mM). The effect of
exogenous MgCl2 on intracellular killing was dose-dependent and visible at concentrations above 100 mM.
E. Wild-type or kil2 mutant cells were incubated with GFP-expressing Klebsiella for 90 min. The cells were then fixed, and the number of
intracellular fluorescent bacteria was determined as described in the legend to Fig. 4. The intracellular accumulation of live bacteria in kil2
cells was abolished by the addition of 10 mM MgCl2 in the medium.
All experiments were repeated at least three times (average and SEM are indicated). The statistical significance of the results was determined
using the Student’s t-test (*P < 0.05 and **P < 0.01).
the p80 endosomal marker (Fig. 6). Mitochondrial porin
was virtually absent from phagosomal preparations
(Fig. 6). The presence of Kil2 in purified phagosomal
membranes suggests that it may participate in creating an
appropriate environment for efficient intracellular killing of
ingested Klebsiella.
A role for magnesium in bacterial killing
Assuming that Kil2 pumps a specific cation into the phagosomal lumen that is essential for Klebsiella killing, then
providing this ion in the extracellular medium may restore
its intra-phagosomal concentration and consequently efficient killing in kil2 mutant cells. A similar strategy has been
used previously to determine the ability of various ions to
induce expression of bacterial genes within phagosomes
(Martin-Orozco et al., 2006). However, this approach is
limited by the fact that certain ions have toxic or inhibitory
effects on cells. We thus tested the effect of each ion at
the highest concentration at which it did not inhibit killing
of Klebsiella by wild-type Dictyostelium. Extracellular
addition of CaCl2 (10 mM), NaCl (10 mM), KCl (10 mM),
MnCl2 (10 mM), FeSO4 (10 mM), ZnCl2 (1 mM) or NiCl2
(0.5 mM) did not increase killing of Klebsiella in kil2 cells
(Fig. 7A). However, efficient killing was restored in kil2
cells by the extracellular addition of MgCl2 (10 mM)
(Fig. 7A). In the presence of magnesium, kil2 mutant cells
killed Klebsiella as rapidly as wild-type cells (Fig. 7B).
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252 E. Lelong et al.
Fig. 8. kil2 knockout cells are sensitive to
high magnesium concentrations. Growth of
wild-type and kil2 mutant cells was tested in
HL5 medium supplemented or not with
100 mM MgCl2. In HL5, kil2 knockout cells
grew as well as wild-type cells. In the
presence of 100 mM MgCl2, kil2 cells grew
more slowly than wild-type cells. This
experiment was reproduced three times with
similar results.
Remarkably, addition of extracellular magnesium had no
effect on the killing defect of phg1a mutant cells (Fig. 7C).
This contrasting behaviour rules out the possibility that the
observed effect was due to a non-specific toxic effect of
magnesium on bacteria. The effect of magnesium on the
killing ability of kil2 cells was dose-dependent, and detectable at concentrations above 100 mM (Fig. 7D). This
result was also confirmed by measuring intracellular accumulation of live Klebsiella-GFP: in the presence of 10 mM
MgCl2, kil2 cells no longer accumulated high numbers of
undigested intracellular bacteria (Fig. 7E).
The ionic specificity of a P-type ATPase can also be
determined by measuring the ability of a mutant strain to
grow when exposed to very high concentrations of
various ions (Schmidt et al., 2009). For this we used ionic
concentrations that slowed down significantly but not
completely growth of wild-type Dictyostelium cells. Mutant
kil2 cells grew as well as wild-type cells in HL5 medium,
but significantly more slowly in a medium supplemented
with 100 mM MgCl2 (Fig. 8). No significant difference
between wild-type and kil2 cells were observed when
cells were grown in medium supplemented with KCl
(200 mM), NaCl (150 mM), CaCl2 (80 mM), MnCl2
(15 mM), CdCl2 (50 mM) or NiCl2 (350 mM) (data not
shown). These observations suggest that Kil2 may participate in sequestration of magnesium ions in endosomal
compartments.
In order to understand how phagosomal magnesium
may influence bacterial killing, we measured in living cells
the activity of phagosomal proteases. For this, latex
beads coupled to fluorescently labelled BSA were fed to
amoeba, and the fluorescence recorded as a function of
time. The fluorescence attached to the beads is
quenched, but increases when it is released from the
beads upon proteolysis of BSA (see Experimental procedures). This assay revealed that phagosomal protease
activity was significantly lower in kil2 knockout cells than
in wild-type cells (Fig. 9A), a defect fully compensated by
the addition of magnesium in the medium (Fig. 9B). This
result suggests that Kil2 controls the activity of phagosomal proteases by maintaining an appropriate magnesium
concentration in phagosomes.
Kil2 is not implicated in intracellular bacterial replication
In addition to its role in bacterial killing, the ionic composition of phagosomes is also a key determinant in the
intracellular replication of some bacterial pathogens. This
has been established particularly clearly by studying cells
defective for the function of Nramp1, a putative phagosomal iron transporter. The lack of activity of Nramp1
(natural resistance-associated macrophage protein)
renders mammalian phagocytic cells more susceptible to
intracellular replication of several different intracellular
pathogens, notably Salmonella, mycobacteria and Leishmania (reviewed in Papp-Wallace and Maguire, 2006). In
Dictyostelium, intracellular replication of mycobacteria
and of Legionella is facilitated in nramp1 knockout cells
Fig. 9. Phagosomal proteolysis is defective in kil2 mutant cells, but
restored by exogeneous magnesium. Wild-type or kil2 mutant cells
were incubated with latex beads coupled to fluorescent BSA.
Fluorescence is quenched at the surface of beads, but increases
when the fluorophore is released in the lumen upon digestion of
BSA.
A. In phosphate buffer, protease activity was lower in kil2 mutant
cells than in wild-type cells.
B. Addition of 10 mM MgCl2 to the phosphate buffer restored
normal protease activity in kil2 mutant cells. The curves represent
the average and SEM of 13 independent measures in 6
independent experiments. The values attained after 3 h were
significantly different for wild-type and kil2 mutant cells in
phosphate buffer (P < 0.01; Student’s t-test), but not in buffer
supplemented with 10 mM MgCl2 (P > 0.25). The protease activity
was also not significantly different in wild-type cells incubated in
phosphate buffer or in magnesium-enriched buffer (P > 0.25).
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P-type ATPase, magnesium and intracellular killing
(Peracino et al., 2006). To determine if nramp1 and kil2
mutant cells present similar phenotypes, we investigated
first if intracellular replication of bacteria was facilitated in
kil2 cells, and second if intracellular killing of Klebsiella
was inhibited in nramp1 cells.
Dictyostelium is an established host model for pathogenic Mycobacterium marinum, and the replication of
GFP-expressing M. marinum can be visualized and quantified with precision (Hagedorn and Soldati, 2007). During
infection of wild-type cells, internalized M. marinum are
found in a p80-positive replication compartment, from
which they eventually escape to the cytosol. The replication niche of M. marinum was indistinguishable in wildtype and kil2 cells, and accumulation of p80, as well as
vacuole rupture were frequently observed (Fig. S5A).
Flow cytometry analysis of infected cells also revealed a
virtually identical rate of intracellular replication in wildtype and mutant cells (Fig. S5B). Finally, an avirulent M.
marinum mutant (L1D) was unable to replicate in kil2
mutant cells as well as in wild-type cells (Fig. S5B). These
results indicate that the Kil2 protein is not involved in
intracellular replication of M. marinum in Dictyostelium
cells, neither as a limiting factor, nor as a facilitating
element.
Conversely, we did not observe any defect in the ability
of nramp1 mutant cells to kill Klebsiella compared with
wild-type cells (Fig. S6), indicating that Nramp1 does not
play a critical role in intracellular killing of Klebsiella in
Dictyostelium. Together, these results suggest that different host mechanisms are involved in intracellular killing
of Klebsiella and during intracellular replication of M.
marinum.
Discussion
In this study we identified in a random genetic screen Kil2,
a new gene product involved in intra-phagosomal killing of
Klebsiella in Dictyostelium. In kil2 knockout cells, the
overall organization of the endocytic pathway is essentially unaffected, but intra-phagosomal killing of Klebsiella
is very inefficient. Kil2 exhibits all the sequence characteristics of a type V P-ATPase and is present in the phagosomal membrane, it may thus be expected to pump
cations into the phagosomal lumen. Efficient killing was
restored in kil2 mutant cells when magnesium ions were
added to the extracellular medium together with the bacteria, presumably restoring an adequate magnesium concentration in phagosomal compartments. The simplest
interpretation of our results is that Kil2 transports magnesium into the phagosome, and that magnesium is required
for optimal protease activity and efficient intracellular
killing of Klebsiella.
Not much is known about type V P-ATPases. Although
it is assumed that, like most other P-type ATPases, they
253
are cation pumps, their substrate specificity is not established with certainty (Axelsen and Palmgren, 1998;
Schultheis et al., 2004). In human, mutations in ATP13A2,
a lysosomal member of the family, are responsible for a
hereditary form of Parkinsonism, but the cellular function
of ATP13A2 is unclear (Ramirez et al., 2006). At the cellular level, type V P-ATPases were mostly studied in Saccharomyces cerevisiae. In this organism, there are 16
P-type ATPases (Catty et al., 1997), two of which belong
to the type V: Cod1/Spf1/YEL031w and Ypk9p. Ypk9 has
been localized to the vacuole, and its absence renders
cells more sensitive to heavy metal ions (e.g. Cd2+ and
Ni2+), suggesting that it may sequester these ions in the
vacuole (Gitler et al., 2009; Schmidt et al., 2009). Cod1
plays a role in the ionic homeostasis of the ER, and its
loss leads notably to alterations of the secretory pathway
(e.g. protein stability, glycosylation) (Cronin et al., 2002).
Interestingly, there are indications that Cod1 may transport magnesium ions (Cronin et al., 2002). Indeed magnesium was the only ion capable of stimulating Cod1p
ATPase activity in vitro. The effect was seen at relatively
high concentrations (1 to 10 mM, i.e. 200 times more than
the ATP concentration), suggesting that it could not be
attributed to ATP-bound magnesium ions (Cronin et al.,
2002). In summary, although the ion specificity of type V
P-ATPases remains to be firmly established, they may be
involved in the transport of heavy metal ions, and of
magnesium. Our results reinforce the notion that at
least some type V P-ATPases may act as magnesium
transporters.
The role of magnesium in bacterial survival within phagosomes has been a debated subject, particularly its role
in intracellular replication of Salmonella. It was initially
observed that magnesium depletion in vitro induces
the expression of several Salmonella genes essential for
replication within phagosomes, notably phoPQ (Garcia
Vescovi et al., 1996). Another gene, mgtC, was shown to
be essential for growth of Salmonella in magnesiumdepleted medium in vitro, as well as in phagosomes (Alix
and Blanc-Potard, 2007). These observations led to the
idea that bacteria encounter conditions of magnesium
depletion in phagosomes. This was proposed to be part of
a general cellular strategy aimed at depriving internalized
bacteria of nutrients, in order to control their replication
and to facilitate their killing (Appelberg, 2006). However, it
was later shown that phagosomal acidification, and not
magnesium depletion, may be the critical element inducing PhoPQ in phagosomes (Martin-Orozco et al., 2006).
Other studies also suggested that the two functions of
MgtC (growth in magnesium-deprived medium and intracellular replication) can be dissociated (Alix and BlancPotard, 2007). Finally, direct measurement revealed a
stable concentration of magnesium (approximately 1 mM)
in phagosomes (Martin-Orozco et al., 2006), suggesting
© 2010 Blackwell Publishing Ltd, Cellular Microbiology, 13, 246–258
126
254 E. Lelong et al.
the existence of a mechanism actively maintaining magnesium concentration in this compartment. Our results
extend this notion one step further: they suggest that the
presence of magnesium in phagosomes is not beneficial
to all internalized bacteria, and is actually critical to
achieve efficient killing of internalized Klebsiella. Our
results suggest that a minimal concentration of magnesium may be necessary for optimal activity of phagosomal
enzymes (e.g. proteases). Exogenous magnesium even
stimulates the killing of Klebsiella by wild-type cells,
maybe because an optimal phagosomal magnesium concentration is more readily attained in these conditions.
While the simplest interpretation of our results would be
that Kil2 itself is a magnesium transporter, there are a
number of alternative interpretations. One possibility is
that Kil2 participates only indirectly in magnesium homeostasis in phagosomes, for example, if its activity is
coupled to that of other magnesium-specific channels.
Another scenario is that there may be several redundant
killing mechanisms, one Kil2-dependent, the other
magnesium-dependent. Magnesium would then only be
necessary for killing when the Kil2-dependent killing is
inactivated, i.e. in kil2 but not in wild-type cells.
From a more general perspective, our results confirm
previous observations suggesting that the killing of Klebsiella mobilizes a specific set of gene products, which is
not essential for the killing of other types of bacteria (e.g.
B. subtilis or P. aeruginosa). Indeed the growth and
killing defects of kil2 mutant cells are very similar to
those observed for phg1a and kil1 mutant cells previously (Benghezal et al., 2006), and are restricted to a
small subset of Gram-negative bacteria (Klebsiella and a
mucoid E. coli isolate). This subset of bacteria is most
likely defined by specific bacterial surface determinants,
as suggested by the fact that kil2 and phg1a mutant cells
kill readily Klebsiella mutants with defective cell wall synthesis. This result stresses the importance of the nature
of the bacterial surface in determining resistance to specific killing mechanisms. Intracellular replication of mycobacteria is not affected in kil2 mutant cells, suggesting
that Kil2 is involved neither in limiting mycobacterial replication, nor in facilitating it. Conversely, we observed that
Nramp1, which inhibits replication of mycobacteria (Peracino et al., 2006), was dispensable for efficient killing of
Klebsiella. Overall, these results indicate that distinct
host mechanisms are involved in the killing of different
types of bacteria, and in limiting the replication of pathogens. An extensive genetic analysis will be necessary to
determine the mechanisms involved in the intracellular
killing or survival of various types of bacteria. In this perspective, Dictyostelium amoebae are attractive model
phagocytic cells, as they are amenable to genetic analysis, and their interactions with bacteria can be studied
relatively easily.
Experimental procedures
Cells and reagents
Unless otherwise specified, all mutant Dictyostelium strains used
in this study were derived directly from the subclone DH1-10
(Cornillon et al., 2000) of the DH1 strain (Caterina et al., 1994),
referred to here as wild-type for simplicity. The phg1a mutant was
described previously (Cornillon et al., 2000). The nramp1 mutant
and the corresponding AX2 parental strain were a kind gift of Dr
S. Bozzaro (University of Turin, Italy) (Peracino et al., 2006).
A polyclonal antibody recognizing the Kil2 protein was
obtained by immunization of rabbit (Covalab, France) with two
peptides corresponding to sequences in the C-terminal portion of
Kil2: KSKRKLKQKQNSDP and IIAKNTVNERYTSLN. The H161
monoclonal antibody recognizing the p80 endosomal protein and
the monoclonal antibody 70-100-1 recognizing the mitochondrial
porin were described earlier (Troll et al., 1992; Ravanel et al.,
2001).
Bacterial strains were a K. pneumoniae laboratory strain and
isogenic mutants (Benghezal et al., 2006), the isogenic P. aeruginosa strains PT5 and PT531 (rhlR-lasR avirulent mutant)
(Cosson et al., 2002), the P. aeruginosa strain PT894 and the
isogenic DP5 (trpD) and DP28 (pchH) avirulent mutants (Alibaud
et al., 2008), the E. coli strains DH5a (Invitrogen), and B/r
(Gerisch, 1959), non-sporulating B. subtilis 36.1 (Ratner and
Newell, 1978), Micrococcus luteus (Wilczynska and Fisher,
1994), and the avirulent Aeromonas salmonicida JF2397 strain
(Froquet et al., 2007).
Cell culture and mutagenesis
Dictyostelium discoideum cells were grown at 21°C in HL5
medium (Mercanti et al., 2006) and subcultured twice a week to
maintain a density < 106 cells ml-1.
To test the effect of various ions on growth of Dictyostelium,
cells were cultivated in HL5 complemented with increasing concentrations of various ions, to attain a concentration at which
growth was slowed down but not fully inhibited (MgCl2: 100 mM,
KCl: 200 mM, NaCl. 150 mM, CaCl2: 80 mM, MnCl2: 15 mM,
CdCl2: 50 mM, NiCl2: 350 mM). Growth curves were obtained in
these conditions by seeding the cells at 10 000 cells ml-1 and
recording their growth over up to 3 weeks.
To isolate killing-deficient mutants, cells were mutagenized
by restriction enzyme-mediated integration of the pSC plasmid
(Cornillon et al., 2000; Guerin and Larochelle, 2002), and transfected cells selected in the presence of blasticidin (10 mg ml-1). A
cell sorter was used to clone single cells into individual wells of
96-wells plates. A Replica Plater for 96 wells plate (SigmaAldrich) was used to transfer 2 ml of each clone to a lawn of
Klebsiella. Overall, 2000 individual clones were tested for their
ability to grow efficiently on Klebsiella and five were unable to.
Mutant cells were expanded and their genomic DNA was
extracted. The inserted pSC plasmid was recovered with the
genomic flanking regions after a ClaI digestion and the insertion
site determined by sequencing (Cornillon et al., 2000). Further
studies were focused on one of these clones (kil2), which was the
only one corresponding to an insertion within a coding sequence.
A new knockout vector was constructed to delete the sequence of
the kil2 gene (Figs 1C and S1). Transfected cells were cloned by
limiting dilution, and screened by PCR (Fig. S1) (Charette and
© 2010 Blackwell Publishing Ltd, Cellular Microbiology, 13, 246–258
127
P-type ATPase, magnesium and intracellular killing
Cosson, 2004). Two independent mutant clones were used with
identical results in all experiments presented in this study.
Growth of Dictyostelium on bacteria
Procedures to test growth of various Dictyostelium strains on
bacteria have been described previously (Froquet et al., 2009).
Briefly, bacteria were grown overnight in LB, then 50 ml of the
culture was deposited and dried on 2 ml of SM-Agar (10 g l-1
peptone, 1 g l-1 yeast extract, 2.2 g l-1 KH2PO4, 1 g l-1 K2HPO4,
1 g l-1 MgSO4:7H2O, 10 g l-1 glucose, 20 g l-1 Agar) in one well of
a 24-well plate. When indicated, an antibiotic was added to the
SM-agar (gentamycin 25 mg ml-1) or bacteria were boiled at 95°C
during 4 h before depositing them on SM-Agar. Variable numbers
of wild-type or mutant Dictyostelium amoebae (10 000, 1000, 100
or 10) were deposited on the bacterial lawn, and allowed to grow
at 21°C for 4–5 days, i.e. until individual colonies of wild-type
Dictyostelium became visible.
Intracellular killing of bacteria by Dictyostelium
Phagocytosis and killing of bacteria were assessed as described
previously (Benghezal et al., 2006). Briefly, 1.5 ¥ 104 bacteria
from an overnight liquid culture (in LB for Klebsiella or B. subtilis;
in SM deprived of glucose for P. aeruginosa PT531) were mixed
with 106 Dictyostelium cells in 500 ml of phosphate buffer (2 mM
Na2HPO4, 14.7 mM KH2PO4, pH 6.5) and incubated at 21°C with
shaking. When indicated, salts were added to the phosphate
buffer: MgCl2 (10 mM), CaCl2 (10 mM), NaCl (10 mM), KCl
(10 mM), MnCl2 (10 mM), FeSO4 (10 mM) or ZnCl2 (1 mM) or NiCl2
(0.5 mM). After 0, 1, 2, 4 or 6 h of incubation, a 10 ml aliquot of the
suspension was collected, diluted in 40 ml of ice-cold sucrose
(400 g l-1). Two hundred microlitre phosphate buffer containing
0.5% saponin was added, before plating on a LB-agar plate and
incubating at 37°C. This procedure was previously shown to kill
Dictyostelium cells, without affecting bacterial viability. The bacterial colonies were counted 24 h later. When indicated, the
number of viable bacteria associated with Dictyostelium cells
(intracellular fraction) was determined by washing the cells twice
with ice-cold HL5 medium before diluting in sucrose.
In order to visualize live intracellular bacteria, 25 ¥ 106 GFPexpressing Klebsiella from an overnight culture were mixed with
5 ¥ 105 Dictyostelium cells in 500 ml of HL5 medium. After 60 min
of shaking incubation, cells were allowed to attach on a glass
coverslip for 30 min, fixed and processed for immunofluorescence as described previously (Mercanti et al., 2006) using the
H161 monoclonal antibody and an Alexa-546-coupled secondary
antibody to reveal the endosomal p80 marker. Cells were analysed with a LSM510 confocal microscope (Carl Zeiss).
Intracellular replication of Mycobacterium was determined as
described (Hagedorn and Soldati, 2007), Dictyostelium cells
were incubated with M. marinum expressing GFP and the replication of bacteria was followed during 2 days using a flow cytometer. When cells were analysed by confocal microscopy, they
were fixed and the p80 marker revealed by immunofluorescence.
Purification of phagosomes
Phagosomes containing latex beads were purified as described
(Gotthardt et al., 2006b). Briefly, Dictyostelium cells were incu-
255
bated with 0.8 mm latex beads during a pulse of 5 or 15 min, then
washed and incubated further for 15 or 45 min. Phagosomes
containing latex beads were purified by flotation on sucrose
gradient. Proteins were then separated in SDS-polyacrylamide
gels, transferred to nitrocellulose, detected with specific antibodies, horseradish-peroxidase-coupled secondary antibodies
(Bio-Rad), and visualized by ECL.
Endosomal and lysosomal pathways
Endosomal pH was measured as described in (Marchetti et al.,
2009), by following at various times after internalization the
fluorescence levels of two internalized dextrans, one coupled to
a pH-sensitive fluorophore (Oregon green), and one to a
pH-insensitive fluorophore (Alexa 647). Lysosomes and postlysosomes were detected by co-immunofluorescence with antibodies to H+-ATPase and p80, and their number and size
analysed as previously described (Charette and Cosson, 2007).
Transfer of internalized latex beads from lysosomes to postlysosomes was measured as described previously (Charette and
Cosson, 2007) to determine the rates of transfer between these
two compartments. Briefly, Dictyostelium cells were incubated
15 min with FITC-latex beads, washed to eliminate uningested
beads, and then incubated for different times before fixation and
immunofluorescence. The number of beads in lysosomes (H+ATPase-positive, p80-positive) and post-lysosomes (H+-ATPasenegative, p80-positive) was determined, and the fraction present
in post-lysosomes is indicated.
The activity of lysosomal enzymes in cells and in the extracellular medium was measured using a colorimetric assay as
described previously (Froquet et al., 2008).
Kinetic analysis of phagosomal proteolytic activity was performed using a fluorescence plate reader (Synergy Mx, Biotek)
as described (Russell et al., 2009), based on the principle of dye
dequenching induced by proteolysis of the carrier protein. Briefly,
the proteolytic reporter Self-Quenched BODIPY® Dye Conjugates of Bovine Serum Albumin (DQ Green BSA, Molecular
Probes) was coupled to 3 mm carboxylate-modified silica particles (Kisker Biotech). As a reference dye, particles were also
coupled with Alexa Fluor 594-SE (Molecular Probes). Dictyostelium cells were plated as a monolayer in clear bottom black wall
96-well dishes (Costar) and allowed to adhere in phosphate
buffer. The fluorescent beads were added to the cells at a ratio of
1:2 and the plate was centrifuged for 30 s. Non-ingested beads
were removed immediately by washing twice with phosphate
buffer supplemented, when indicated with 10 mM MgCl2. The
emission fluorescence was measured in the same buffer at
490 nm and 450 nm excitation every 90 s over a period of
180 min. The 490/450 nm ratio reflects the bulk proteolytic activity within the bead-containing phagosomes.
Acknowledgements
This work was supported by grants from the Fonds National
Suisse de la Recherche Scientifique (http://www.snf.ch) to P.C.
and T.S. The P.C. research group is supported by the
Doerenkamp-Zbinden Foundation (http://www.doerenkamp.ch)
and the Fondation E. Naef pour la Recherche in Vitro (http://
www.fondation-naef.com). The P.C. and T.S. research groups are
part of the NEMO network supported by the 3R Research Foun-
© 2010 Blackwell Publishing Ltd, Cellular Microbiology, 13, 246–258
128
256 E. Lelong et al.
dation Switzerland (http://www.forschung3r.ch). We thank Drs
Brian VanderVen and David G. Russell for the opportunity to
learn the proteolytic activity assay in their lab, supported by an
‘Individual short visit fellowship’ to MH (Swiss National Science
Foundation IZK0A3-121674). We thank S.J. Charette (Université
Laval, Canada) for critical reading of the manuscript.
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Supporting information
Additional Supporting Information may be found in the online
version of this article:
Fig. S1. Isolation of kil2 knockout mutants. A. Schematic representation of the kil2 gene in wild-type or kil2 cells and of the kil2
knockout vector. Arrows indicate the position of oligonucleotides
used to construct the knockout vector and to identify knockout
257
mutant cells. B. Sequences of the oligonucleotides and their
position on the kil2 genomic sequence are indicated. C. kil2
knockout mutants were identified by PCR with three different
pairs of oligonucleotides. The PCR fragment observed after
amplification of wild-type genomic DNA with pairs 1 + 4 and 5 + 6
is absent when kil2 mutant cells are tested. In kil2 mutant cells,
a specific PCR amplification is seen with oligonucleotides
1 + BSRa. The results obtained with wild-type cells and three
independent kil2 knockout clones are shown.
Fig. S2. Phylogenetic tree of P-type ATPases in five eukaryotic
organisms. Protein sequences of P-type ATPases from five fully
sequenced eukaryotic genomes were aligned with CLUSTALX
2.0 program. The distance-based phylogenetic tree was generated using the neighbour-joining algorithm (as implemented in
the PHYLIP package), and bootstrap assessment of the tree
topology was performed with one thousand replicates. The
branch lengths are proportional to the number of amino acid
substitutions per site. Numbers at the nodes represent percentage of bootstrap support (only values > 60% are indicated).
Uniprot protein IDs are shown, followed by the organism abbreviation: D. discoideum (DI), H. sapiens (HU), A. thaliana (AT), S.
cerevisiae (YST) and S. pombe (SPO). Kil2 clusters with P-type
ATPases from group V (Axelsen and Palmgren, 1998).
Fig. S3. The endosomal pH is very similar in wild-type and kil2
cells. Endosomal pH was measured in wild-type and kil2 mutant
cells following a protocol described previously (Marchetti et al.,
2009). For this, Dictyostelium cells were incubated for 20 min in
the presence of a mixture of dextran coupled to Oregon green(OG, pH-sensitive) and to Alexa 647 (A-647, pH-insensitive). The
cells were then washed and incubated further in HL5. The intracellular fluorescence was measured by flow cytometry at each
indicated time. This experiment was repeated three times with
equivalent results. A. The cell-associated fluorescence of both
probes exhibited the same profiles in wild-type and kil2 cells. B.
The fluorescence ratio of the two probes provides an estimate of
the pH of endosomes at various times following endocytosis. The
endosomal pH is virtually identical at all times in wild-type and kil2
mutant cells. C. A calibration curve was obtained in parallel by
incubating cells having endocytosed dextrans in medium at a
defined pH, in the presence of sodium azide and ammonium
chloride. Approximate pH values can be obtained by comparing
the values in B with the calibration curve. These results demonstrate that in wild-type and kil2 cells, fluid phase is endocytosed,
transferred from acidic lysosomes to less acidic post-lysosomes,
and recycled to the extracellular medium with virtually identical
kinetics. It also indicates that the pH of the various endocytic
compartments is indistinguishable in both cell types.
Fig. S4. The general organization of the endocytic/phagocytic
pathway is not altered in kil2 mutant cells. A. In order to
measure the kinetics of maturation of phagosomes, Dictyostelium cells were incubated with 1 mm latex beads for 15 min,
washed to eliminate non-internalized beads and incubated
further for 15, 45 and 75 min (total incubation time of 30, 60 and
90 min). Cells were then fixed, and p80 and H+-ATPase were
detected by immunofluorescence in order to determine if beads
were present in lysosomes (p80-positive, H+-ATPase-positive) or
post-lysosomes (p80-positive, H+-ATPase-negative). At each
time, 30 internalized beads were analysed, and the percentage
of beads present in post-lysosomes was determined. The
average and SEM of three independent experiments are indicated. In wild-type and in kil2 cells, all internalized beads were
© 2010 Blackwell Publishing Ltd, Cellular Microbiology, 13, 246–258
130
258 E. Lelong et al.
found initially in lysosomes, then transferred with very similar
kinetics to post-lysosomes. B. Cells were grown for 3 days in
HL5 medium, and recovered by centrifugation. The activity of
two lysosomal enzymes (NAG: N-acetyl b-glucosaminidase;
MAN: a-mannosidase) was determined in cell pellets and in
supernatants. The total activity of lysosomal enzymes (full bars)
was similar in wild-type and kil2 mutant cells. In both cells, only
a small fraction of lysosomal enzymes was released in the
medium (empty bars). Each bar indicates the average and SEM
of three independent experiments.
Fig. S5. Mycobacterium marinum replicates normally in kil2
mutant cells. Dictyostelium cells were infected with wild-type or
mutant (L1D) M. marinum expressing GFP (green). A. At 37 h
post-infection, cells were fixed and p80 revealed by immunofluorescence (white). Mycobacterium replication vacuoles were indistinguishable in wild-type and kil2 cells: several bacteria were
found within p80-positive replication vacuoles (upper panel) from
which they also escaped into the cytosol (lower panel) (scale bar:
5 mm). B. Intracellular replication of M. marinum was measured
by flow cytometry over a period of 2 days. The total amount of
fluorescence (intracellular and extracellular) is indicated. Wildtype M. marinum replicated with similar kinetics in kil2 and in
wild-type cells. L1D mutant mycobacteria were incapable of replicating in either cell type.
Fig. S6. nramp1 mutant cells kill Klebsiella efficiently. Wild-type
or nramp1 cells were incubated with Klebsiella and the number of
remaining live bacteria was determined at different times by
plating an aliquot on LB plates and counting bacterial colony
forming units, as described in the legend to Fig. 5. Wild-type and
nramp1 mutant cells killed Klebsiella with very similar kinetics.
Table S1. Endocytic compartments are very similar in wild-type
and kil2 cells. Endocytic compartments were analysed by confocal microscopy in wild-type or kil2 cells: endosomal p80 and
H+-ATPase were detected by immunofluorescence in order to
differentiate lysosomes (Ly, p80-positive, H+-ATPase-positive)
and post-lysosomes (PL, p80-positive, H+-ATPase-negative).
Compartments were counted, and their size determined in at
least 20 cells. The average and SEM values in three independent
experiments are indicated.
Please note: Wiley-Blackwell are not responsible for the content
or functionality of any supporting materials supplied by the
authors. Any queries (other than missing material) should be
directed to the corresponding author for the article.
© 2010 Blackwell Publishing Ltd, Cellular Microbiology, 13, 246–258
131
5.2-The balance in the delivery of ER components and the vacuolar proton pump to the
phagosome depends on myosin IK in Dictyostelium
Contributed Table I
132 Research
© 2012 by The American Society for Biochemistry and Molecular Biology, Inc.
This paper is available on line at http://www.mcponline.org
The Balance in the Delivery of ER Components
and the Vacuolar Proton Pump to the
Phagosome Depends on Myosin IK in
Dictyostelium*□
S
Régis Dieckmann‡**, Aurélie Guého‡, Roger Monroy‡, Thomas Ruppert§,
Gareth Bloomfield¶, and Thierry Soldati‡储
In Dictyostelium, the cytoskeletal proteins Actin binding
protein 1 (Abp1) and the class I myosin MyoK directly
interact and couple actin dynamics to membrane deformation during phagocytosis. Together with the kinase
PakB, they build a regulatory switch that controls the
efficiency of uptake of large particles. As a basis for further functional dissection, exhaustive phagosome proteomics was performed and established that about 1300
proteins participate in phagosome biogenesis. Then,
quantitative and comparative proteomic analysis of phagosome maturation was performed to investigate the impact of the absence of MyoK or Abp1. Immunoblots and
two-dimensional differential gel electrophoresis of phagosomes isolated from myoK-null and abp1-null cells
were used to determine the relative abundance of proteins during the course of maturation. Immunoblot profiling showed that absence of Abp1 alters the maturation
profile of its direct binding partners such as actin and the
Arp2/3 complex, suggesting that Abp1 directly regulates
actin dynamics at the phagosome. Comparative twodimensional differential gel electrophoresis analysis resulted in the quantification of mutant-to-wild type abundance ratios at all stages of maturation for over one
hundred identified proteins. Coordinated temporal
changes in these ratio profiles determined the classification of identified proteins into functional groups. Ratio
profiling revealed that the early delivery of ER proteins to
the phagosome was affected by the absence of MyoK and
was coupled to a reciprocal imbalance in the delivery of
the vacuolar proton pump and Rab11 GTPases. As direct
functional consequences, a delayed acidification and a
reduced intraphagosomal proteolysis were demonstrated
From the ‡Départment de Biochimie, Faculté des Sciences, University de Genève, Sciences II, 30 quay Ernest Ansermet, CH-1211
Genève-4, Switzerland; §Core Facility for Mass Spectrometry and
Proteomics, Zentrum für Molekulare Biologie der Universität Heidelberg (ZMBH), Im Neuenheimer Feld 282, D-69120 Heidelberg, Germany; ¶MRC Laboratory of Molecular Biology, Hills Road, Cambridge
CB2 0QH, UK
Received February 2, 2012, and in revised form, May 22, 2012
Published, MCP Papers in Press, June 26, 2012, DOI 10.1074/
mcp.M112.017608
886
in vivo in myoK-null cells. In conclusion, the absence of
MyoK alters the balance of the contributions of the ER and
an endo-lysosomal compartment, and slows down phagosome acidification as well as the speed and efficiency of
particle degradation inside the phagosome. Molecular &
Cellular Proteomics 11: 10.1074/mcp.M112.017608, 886 –
900, 2012.
Professional phagocytes, ranging from phagotrophic protozoa to specialized cells of the innate immune system such
as macrophages, neutrophils, or dendritic cells, ingest and
digest large particles (⬎ 250 nm). Indeed, the core machineries acting in phagocytosis have been conserved as its basic
purpose evolved from predation and feeding to antigen presentation (1). Particle recognition by the phagocytes initiate the
internalization process and triggers remodeling of the plasma
membrane and its underlying cytoskeleton, to project a circular cup-shaped lamella around the particle. The phagocytic
cup finally encloses the particle into a de novo membranebound vacuole, the phagosome. Endomembrane compartments, such as early and late endosomes (2– 4) and the endoplasmic reticulum (5), are recruited to provide membrane to
form the phagosome. Modifications of the environment within
the closed phagosome generates a microbicidal milieu and
leads to particle degradation. Maturation of the phagosome
is, basically, a linear process driven by a flux of incoming and
outgoing vesicular trafficking, and is characterized by a progressive decrease in pH and successive fusions with early
endosomes, late endosomes, and lysosomes (6, 7). The trafficking events are controlled by small GTPases and the soluble N-ethylmaleimide-sensitive factor attachment protein
receptor (SNARE) machinery (6, 8), and phagosome acidification is driven by the activity of the vacuolar H⫹-ATPase proton
pump (V-ATPase)1 (4).
1
The abbreviations used are: V-ATPase, vacuolar H⫹-ATPase proton pump; Abp1, Actin binding protein 1; MyoK, myosin IK; 2D-DIGE,
two-dimensional fluorescent differential gel electrophoresis; ER, endoplasmic reticulum; ERAD, ER-associated degradation machinery;
ERGIC, ER-Golgi intermediate compartments.
Molecular & Cellular Proteomics 11.10
133
MyoK Modulates the Early Delivery of ER to the Phagosome
In Dictyostelium, a synchronized pulse/chase protocol for
the isolation of latex bead-containing phagosomes was developed that covers the entire maturation process (8, 9). Temporal quantitative profiling by immunoblot and two-dimensional electrophoresis (2-DE) was used to characterize the
protein content of the phagosome and follow the time-dependent variations driving maturation (8, 10). Five successive
phases in the maturation program were clearly identified i.e.
uptake, metabolism/ion exchange, late endosomal, digestive,
and exocytic stages. The uptake phase was the more complex. Abundant proteins characteristic of this phase were
involved in signaling, actin cytoskeleton, membrane dynamics
and the emergence of a degradative phase. Although a clear
sequence of maturation events could be established, the
discontinuity of individual profiles revealed that a protein,
required at different steps of maturation, can be retrieved
in-between (10). A similar plasticity was observed in macrophages by temporal profiling of isolated phagosomes after
stable isotope labeling (11).
Having precisely characterized the maturation process in
wild type Dictyostelium, we used proteomic profiling to investigate the impact on phagocytosis of an ablation of either
Actin binding protein 1 (Abp1) or the class I myosin, MyoK.
Together with the kinase PakB, MyoK and Abp1 build a regulatory switch that regulates the efficiency of uptake of large
particles. These two proteins interact and couple actin dynamics to membrane deformation but exert opposite regulatory roles on phagocytic uptake (12, 13). Compared with wild
type cells, myoK-null mutants display 20% decrease whereas
abp1-null display 35% increase in the rate and extent of
uptake of large particles. Despite their direct interaction,
MyoK and Abp1 localize independently at the phagocytic cup
and on early phagosomes (12). Like other class I myosins,
MyoK is essential for the maintenance of resting cortical tension and provides a force to push on negatively curved membranes (12, 13). Although there is no evidence yet that MyoK
might share these functions with other class I myosins, MyoB
in Dictyostelium (14), and Myosin IB in human (15), for example, are involved in membrane trafficking in the pinocytic
pathway, potentially contributing to membrane deformation
both at the endosomal level and at the plasma membrane
(16). The function of Abp1 itself is not well characterized although its interaction network is well described. It contributes to
vesicle uptake, interacting notably with 1) the membrane binding and force generating class I myosins motors such as MyoK
in Dictyostelium and Myo5p in yeast, 2) the Arp2/3 actin nucleation complex in yeast, and 3) dynamin, a protein involved in
membrane constriction, in Dictyostelium and mammals (12, 17).
Although the role of Abp1 in vesicle formation has been mainly
studied during uptake at the plasma membrane, it might perform similar tasks on endomembranes. Indeed, Abp1 is the
main phagosomal F-actin binding protein (18).
The initial goal of the study was to understand how the
removal of one member of a regulatory complex of the actin
Molecular & Cellular Proteomics 11.10
cytoskeleton remodeling can decrease or increase uptake,
and what the downstream consequences are on phagosome
maturation. On one hand, the uptake rate and extent are
strictly regulated and can be significantly decreased or increased. The Abp1-MyoK-PakB regulatory loop is not the only
illustration of this concept. Mutation of the interacting partners CH-Lim or LimF induce respectively an increase or a
decrease in phagocytosis and the balance is regulated by the
small GTPase Rab21 (19). Increased uptake as a result of a
gene knockout might appear counter-intuitive, however,
knockout mutants for dynamin A, the actin cross-linking protein, Abp34, and both profilins I and II show increased phagocytosis (20, 21). On the other hand, actin polymerization and
Abp1 play an active role in phagosome maturation. Indeed,
the formation of a transient actin coat prevents phagosomelysosome fusion and delays phagocytosis (22) and the presence of Abp1 regulates phagosome binding to actin in vitro
(18). Nevertheless, the impact of a regulatory switch of actin
polymerization on the downstream maturation process has
not been investigated.
As a targeted approach, the maturation profiles of Abp1,
MyoK and their direct binding partners were first determined
by quantitative immunoblots to delineate the impact of myoKor abp1-knockouts. Quantitative profiling of maturation by
2D-DIGE was then used to measure mutant to wild type
protein abundance ratios of more than one hundred identified
phagosomal proteins over the entire maturation process in
both myoK-null and abp1-null mutants. Coordinated changes
were observed in the temporal profiles of functionally related
proteins predicting a delay in phagosome acidification resulting from compensatory trafficking defect in the myoK-null
mutant and an alteration of the phagosomal protease composition in abp1-null cells. Alterations in phagosome acidification and proteolysis were observed in vivo as these predictions were verified at a functional level.
EXPERIMENTAL PROCEDURES
Cell Culture—Dictyostelium cells of the parent wild type strain AX-2
were grown at 22 °C in HL-5c medium (Formedium) supplemented
with 100 units/ml penicillin and 100 ␮g/ml streptomycin (Invitrogen).
myoK null cells (13) and abp1 null mutants were maintained with
additional 5 ␮g/ml blasticidin (12).
Electron Microscopy—Phagosomes directly collected from the
gradient were diluted and normalized for concentration in HESES
(0.25 M sucrose, 20 mM HEPES-KOH, pH 7.2). Phagosomes in equivalent number for each fraction were mixed 1:1 (v/v) in freshly prepared
fixative solution (0.4 M sodium cacodylate, pH 7.2, 4% glutaraldehyde, 0.6% OsO4). Samples were fixed for 1h on a rotating wheel.
Phagosomes were pelleted 15 min, 14⬘000 rpm on a mini-centrifuge
and washed in PBS until the supernatant was clear. Phagosome
pellets were then contrasted and embedded (23, 24).
Flow Cytometry-based Uptake Assay—Uptake was measured as
described (10) with the following modifications. Fluorescent beads of
0.5, 1.0 and 4.5 ␮m in diameter (Fluorescent YG carboxylated beads,
Polyscience) were added to the cell at 800:1, 200:1 and 10:1 ratios,
respectively. Flow cytometry was performed with a FACScalibur (Beckton Dickinson). Data were analyzed with the FlowJo software (TreeStar).
887
134
MyoK Modulates the Early Delivery of ER to the Phagosome
Phagosome Isolation—Latex bead-containing phagosomes were
isolated via flotation on sucrose step gradients and processed as
described (9, 10).
Relative Abundance Profiling by Immunoblotting—Quantitative immunoblots were performed as described (10) with following modifications. The chemiluminescent signal was detected in a UVP
EpiChem II Darkroom equipped with a digital camera. Signals were
quantified with Labworks 4.0 software. Bands were defined as boxes
inside lanes and total box volume was quantified after “filtered profile”
background subtraction. Signal intensities were then normalized, setting the maximum signal intensity to 100%, resulting in a relative
abundance profile during phagosome maturation time. To measure
mutant-to-wild type ratios, phagosomes were isolated in parallel in
the mutant and wild type cells at early time-points (5⬘/0, 15⬘/0).
Immunoblot signal intensities were measured and averaged (n ⫽ 3).
The signal at 5⬘/0 in wild type phagosomes was set as 100% and
signal intensity values in the mutant were normalized accordingly.
Mutant-to-wild type ratios were consistent over the 5⬘/0 and 15⬘/0
time-points. Thus, the whole mutant profile was normalized
accordingly.
Antibodies—Antibodies raised against the following Dictyostelium
antigens were obtained from: (1) actin (mAb, 224-236-1), calreticulin
(mAb, 251-67-1) (gift of Dr. G. Gerisch, MPI for Biochemistry, Martinsried); (2) profilin II (mAb, 174-380-3) (gift of Dr. M. Schleicher, AdolfButenandt-Institute, Munich); (3) Arp3 (pAb; gift of Dr. R.H. Insall,
CR-UK Beatson Institute for Cancer Research, Glasgow), (4) dynamin
A (gift of Dr. D. J. Manstein, Hannover Medical School). The antiMyoK and anti-Abp1 antibodies were described in (12). For immunoblots, goat-anti-rabbit IgGs or goat-anti-mouse IgGs conjugated to
HRP (BioRad) were used at dilutions between 1:2,000 and 1:10,000.
Two-dimensional Gel Electrophoresis (2D-DIGE)—Cell were synchronized for growth in suspension over a 3 days period, harvested
by centrifugation (5 min, 1200 rpm, GH3.8A) and washed twice in
Sørensen buffer. Cell pellets were resuspended in 10 ␮l of 14x complete protease inhibitors (Roche) water solution, flash frozen in liquid
nitrogen and stored at ⫺80 °C. Cells were then lysed in DIGE sample
buffer and samples were labeled directly after determination of the
protein concentration [⬇ 5 ␮g/␮l, 2D-Quant kit (Amersham Biosciences)]. Phagosome pellets were resuspended in a minimal volume of
DIGE sample buffer (7 M urea (Amersham Biosciences), 2 M thiourea
(Amersham Biosciences), 4% 3-((3-Cholamidopropyl)dimethylammonio)-1-propanesulfonate (CHAPS), 15 mM 1,2-diheptanoyl-snglycero-3-phosphatdiyl choline (DHPC) (Avanti Lipids), 30 mM Tris
(Sigma Aldrich), pH 8.5). Protein samples (50 ␮g) were adjusted to the
same concentration (ⱖ 2.5 ␮g/␮l) and labeled in parallel in the DIGE
sample buffer with the respective minimal fluorescent CyDye-NHS
(Cy5, Cy3, Cy2), according to manufacturer’s instructions (Amersham
Biosciences). Labeled samples were mixed and co-migrated with
additional 200 –300 ␮g of corresponding unlabeled sample. Total
protein sample was adjusted to 450 ␮l in two-dimensional sample
buffer (7 M urea, 2 M thiourea, 65 mM DTE 1,4-Dithioerythritol (DTE),
4% CHAPS, 15 mM DHPC, 2% Resolyte ampholytes, Tris 30 mM, pH
8.5). Samples were separated as described (10, 25) with the following
modifications. First dimension was performed in NL 3–10, 24 cm
strips (Amersham Biosciences) at 15 °C with a current of 75 ␮A/strip
and up to 45 kVh. Second dimension was run on polyacrylamide gel
of 12.5%T, 2.6%C (National Diagnostics) at 15 °C, 15 mA/gel, O/N in
an Ettan Dalt Twelve chamber (Amersham Biosciences). Gels were
poured in a six-gel chamber between low-fluorescence plates according to manufacturer’s instructions (Amersham Biosciences). Gels
were scanned in an Ettan DIGE imager (Amersham Biosciences).
Scan times were kept identical in a comparative gel run and adjusted
for each dye to result in the best signal to noise ratio and the most
extensive range of signal intensity.
888
Gel Analysis and Mutant/Wild Type Ratio Profiling—Gels were analyzed with Melanie Image Master 2D Platinum version 6 for DIGE.
The initial number of detected spots was set to 2⬘500 and then spots
were filtered according to spot area, percentage volume and saliency
depending on gel signal-to noise ratio and manually curated. For
phagosome samples, gels corresponding to the same time point were
matched pair-wise. Gels were then matched within the pulse/chase
series, setting the 5⬘/0 time point as a reference. To match spot
absent from the 5⬘/0 time point, matched spots were withdrawn from
analysis and remaining spots were then matched to the 15⬘/105⬘ time
point as a reference. Merging of matched pairs and time-series were
performed in Excel using the SuperCombine add-in (gift from Marianne Tardif and Jérôme Garin, Institute of Life Sciences Research and
Technologies, Grenoble) and resulted in a list containing all matched
spots and their respective mutant/WT ratios during maturation i.e. all
the spot ratio profiles. In case of conflict, matches associated with a
known mutant/WT ratio value and/or the most complete time-series
were favored on the final matching table. Differential spots fulfilled
three criteria: 1) their differential ratio was higher than ⫾1.6, 2) standard deviation of the mean should not overlap, 3) spot detection
should not introduce any potential bias. For analysis of cell lysates, a
ratio of 1.3 (30% difference) was experimentally determined as being
a reasonable cut off to consider bona fide differential spots. As the
spot ratio variability was slightly higher in phagosome samples compared with cell lysates, the cut off was set twice as high (1.6, 60%
difference). Ratio profiles of proteins identified in multiple neighboring
spots (i.e. spot groups or spot trains) are very similar (see supplementary Data). Thus, only an averaged profile of all isoforms is shown
here (i.e. VatB, VatA, CrtA, PDI1, PDI2, …).
Protein Identification by Mass Spectrometry and Data Analysis—
Protein identification was performed essentially as described (10, 26).
In brief, proteins were in-gel digested with trypsin and extracted from
Coomassie stained one-dimensional or two-dimensional gel pieces in
a liquid handling robot (DigestPro MS, Intavis AG) according to standard procedures.
For peptide fingerprinting (PMF) by MALDI-TOF mass spectrometry (Ultraflex, Bruker), samples were analyzed as described (10). Protein sequencing was performed either on an ESI-Q-TOF (QSTAR
Pulsar, PE Sciex) (10) or an Orbitrap (LTQ Orbitrap, Thermo Scientific)
as described (26). From the MS and MS/MS spectra, peak lists were
generated without peak removal after centroiding and deisotoping
(analyst QS 1.1; Applied Biosystems; combined with Mascot script
version 1.6b13; Matrix Science). The peak list was then applied to a
database search against Dictybase, the Dictyostelium protein database (27) (dicty_primary_decoy downloaded on 22.02.2010, 54294
entries), using the Mascot software, version 2.2.4 (Matrix Science).
The Scaffold software (Proteome Software Inc.), version 3.0.8, was
used for editing of MS/MS identifications. The algorithm was set to
use trypsin as enzyme, allowing at maximum for one missed cleavage
site. Iodoacetamide derivative of cysteine was specified in Mascot as
a fixed modification. Deamidation of asparagine and glutamine and
oxidation of methionine were specified in Mascot as variable modifications. For PMF identifications, mass tolerance was set to 100 ppm
using already described parameters and threshold for peak-picking
(10). Protein hits were considered identified if the Mascot score exceeded the significance level (p ⱕ 0.05). For MS/MS protein identifications from two-dimensional spots, mass tolerance was set to 0.1
Da for precursor and fragment ions. Protein identifications were accepted if they could be established at greater than 99.0% probability
and contained at least 2 identified peptides with a peptide identification probability greater than 20%. The threshold was set to maximize
sensitivity and minimize the overlap between correct and incorrect
identity score distribution in the Scaffold software. The relative abundance of each protein in the spot was evaluated. The total number of
Molecular & Cellular Proteomics 11.10
135
MyoK Modulates the Early Delivery of ER to the Phagosome
spectra (NSn) of each protein was normalized to the total number of
spectra of the most abundant protein in the spot (NS1). Proteins with
a relative abundance over 20% of the most abundant protein in the
sample (NSn/NS1 ⬎ 0.2) were excluded from further analysis because
their contribution to the differential fluorescent signal in-gel is minor.
For MS/MS protein identifications from one-dimensional bands, mass
tolerance was set at 10 ppm for precursor ions and 0.5 Da for
fragment ions. Protein identifications were accepted if their protein
identification probability was over 99%, their peptide identification
probability was greater than 95.0% and they were identified by at
least two unique peptides. The protein false discovery rate was 0.1%
and the peptide false discovery rate was 5.3% as calculated with a
probabilistic method. Proteins that contained similar peptides and
could not be differentiated based on MS/MS analysis alone were
listed without further filtering, except redundant entries for actin (31
entries), which were grouped into one common entry (act15,
DDB_G0272520). All protein identifications and the corresponding
assigned peptide sequences are listed in supplemental Tables 3 and
4, respectively.
The list of phagosomal protein (supplemental List S1) is compiled
from published data (1, 10), 2D-DIGE spot identifications (2D MSMS
and 2D PMF, this study) and all proteins identified in the major bands
of a phagosomal sample separated on a 10% one-dimensional SDSPAGE (one-dimensional MSMS) (9). Phagosomal proteins were associated to GO terms and matched in KEGG pathways (http://
www.genome.jp/kegg/) (28, 29) using the Orange freeware (http://
orange.biolab.si/(30)). A stringent p value was applied to select only
significantly represented GO terms (p ⬍ 10⫺5) and KEGG pathways
(p ⬍ 10⫺4). Only functionally non-redundant GO terms are represented. Transmembrane domains and signal peptides in phagosomal
protein sequences were predicted using different prediction programs including TMHMM (31) and SignalP (32).
Microarray Analysis—Cells were synchronized for growth as for
proteomics. 15–20 ⫻ 107 cells were lysed by eight passages in a 5 ml
Dounce homogenizer with a small void clearance (S, 10 –30 ␮m)
pestle and processed with the Qiagen RNA isolation Midi kit following
manufacturer’s instructions. Three biological replicates of total cell
RNA were isolated for each strain. Each mutant strain was compared
with its respective wild type in two technical duplicates, yielding six
direct wild type-to-mutant comparisons for each strain analyzed.
Mutants were compared in two genetic backgrounds (DH1–10 and
AX-2) and data were cross-correlated. Samples and data were processed as described (33). Analysis was performed on a total of 9247
probes, representing 8579 D. discoideum specific genes.
Measure of pH and Proteolytic Digestion in Phagosomes—The
kinetics of intraphagosomal acidification and proteolytic activity were
monitored using a fluorescence plate reader (Synergy Mx, Biotek)
over a period of 120 min at intervals of 1 measurement/1.5 min. The
indicator beads for the pH measurement were prepared as described
(57). Briefly, 3 ␮m carboxylated silica beads (Kisker Biotech, Steinfurt,
Germany) were coupled with BSA and labeled with the pH-sensitive
fluorochrome carboxyfluorescein succinimidyl ester (S.E.) and the
pH-insensitive internal reference fluorochrome Alexa Fluor 594-S.E.
(Molecular Probes). The measurement of proteolytic activity is based
on the principle of dye dequenching induced by proteolysis of the
carrier protein. The indicator beads were prepared as described (58),
In brief, the proteolytic reporter Self-Quenched BODIPY ® Dye Conjugates of Bovine Serum Albumin (DQTM Green BSA, Molecular
Probes) and the reference dye Alexa Fluor 594-S.E. were coupled to
3 ␮m carboxylate-modified silica particles. Dictyostelium cells were
plated as a monolayer in clear bottom black wall 96-well dishes
(Costar) and allowed to adhere in LoFlo medium (Formedium). The
fluorescent indicator beads were added to the cells at a ratio of 1:2
and the plate was centrifuged for 30 s. Non-ingested beads were
Molecular & Cellular Proteomics 11.10
removed immediately by washing twice with Soerensen-buffer. For
pH monitoring, the fluorescent emission of carboxyfluorescein at 520
nm when excited with 490 nm is pH-sensitive, whereas emission at
520 nm, when excited at 450 nm, is largely unaffected by changes in
pH. Conversion from the excitation ratio to pH was achieved through
polynomial regression of a standard curve generated by calculation of
the excitation ratio of the fluorochrome in environments of known pH.
For proteolysis monitoring, the emission fluorescence was measured
in these buffers at 490 nm and 450 nm excitation. The ratio 490/450
nm reflects the bulk proteolytic activity within the bead-containing
phagosomes.
RESULTS
Relative Contribution of Protein Classes to the Composition
of Pure and Intact Isolated Phagosomes—Molecular characterization of the phagosome maturation process in wild type
Dictyostelium cells is based on a powerful phagosome purification strategy associated with a synchronized pulse/chase
particle feeding protocol (Fig. 1A, scheme) (8, 10). Pre-adsorption of latex beads on the cells in the cold, followed by
rapid re-warming, generates a strong and synchronous wave
of uptake (9). Phagosomes isolated at 5 min contained single
particles surrounded by an intact membrane as shown by
electron microscopy and FM4 – 64 lipid dye staining (supplemental Fig. S1). Phagosome maturation can be followed not
only molecularly but also morphologically in isolated phagosomes (Fig. 1A, images). Internal phagosomal vesicles were
observed with the successive appearance of electron-lucent
vesicles, multi-vesicular bodies and electron-dense vesicles,
reminiscent of early endosomes, late endosomes and lysosomes, respectively. Then, morphological complexity diminished so that only electron-lucent internal vesicles were observed at pre-exocytosis time points. Large multi-bead
phagosomes, indicative of intense phagosome-phagosome
fusion, were also visible, but only at late time-points.
An exhaustive list of proteins identified in phagosomes isolated from wild type cells was built by compiling all our data
acquired in this study and others (1, 10). It comprises 1291
proteins identified from two-dimensional and one-dimensional
gels (Fig. 1B, supplemental List S1). Its content in predicted
transmembrane proteins (18%) and secreted proteins (11%
with a signal peptide) as well as in proteins of unknown function
(20%) are similar to what is found in the whole Dictyostelium
proteome (22% of transmembrane proteins, 12% of proteins
with signal peptides, ⬎30% of proteins of unknown function).
Therefore, this list offers the most representative picture to date
of the diverse biological processes participating in phagosome
biogenesis and maturation in Dictyostelium. Protein classification illustrates the central but uncharacterized role of metabolic
enzymes in nutrient assimilation (⬎10%), the importance of
signaling cascades (⬇ 10%) downstream of a small set of
potential receptors (⬇ 2%), and the complexity of membrane
trafficking, targeting and remodeling factors such as Rab
GTPases, SNAREs and cytoskeletal proteins, respectively (⬇
20% in total). Novel categories emerged from the present analysis: 1) lipid metabolism (4.1%), indicative of the intense remod-
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MyoK Modulates the Early Delivery of ER to the Phagosome
FIG. 1. Analysis of phagosome maturation in wild type cells and the regulatory roles of Abp1 and MyoK. A, The major steps of
phagosome maturation in Dictyostelium are depicted along a time axis that indicate the six time points used to isolate phagosomes following
a pulse/chase protocol. In brief, beads are first adsorbed on cells 15⬘ in the cold, followed by an uptake pulse of 15⬘ and a chase of up to 165⬘
for a total of 3 h. Ultrastructural changes were visualized by electron microscopy at the indicated pulse/chase times. The successive formation
of electron-lucent internal phagosomal vesicles (5⬘/0), multi-vesicular bodies (15⬘/0), and electron-dense vesicles (15⬘/15⬘) was observed. At
(15⬘/15⬘) and (15⬘/45⬘), phagosomes contain a complex mix of dense vesicles and multivesicular bodies. Complexity of internal vesicles then
decreased. Only electron-lucent vesicles were observed in the last stage of maturation (15⬘/165⬘). Large multi-bead phagosomes were
observed from (15⬘/45⬘) onward. Each panel shows a representative picture of a mixed population. Scale bar, 0.5 ␮m. B, Functional
classification of an exhaustive wild-type phagosome proteome of 1291 proteins identified by a combination of 1D and 2D PAGE separation and
MS sequencing (supplemental List S1). The number of identified proteins belonging to each class and its percentage to the total are indicated.
The same protein classification terms were used as in our initial study (10). C, Phagocytic uptake of 0.5, 1.0, and 4.5 ␮m fluorescent latex beads
by myoK null (green) and abp1 null (red) cells relative to wild type. Data are expressed as a percentage of beads ingested by wild-type cells
at 90 min. D, MyoK and Abp1 have differential temporal profiles during phagosome maturation as shown by immunoblots of phagosomes
purified at the indicated times. Immunoblot signals were quantified and normalized to 100% according to the maximum value in wild type.
Mutant profiles are normalized according to mutant-to-wild type ratios at early time-points (5⬘/0, 15⬘/0) as described in details in the Material
and Methods section.
eling of phagosomal membrane lipid composition and the degradation of the prey by lipases, 2) cell defense, including
proteins involved in immunity-related functions and particle
degradation, possibly representing proteins involved in specific
bacteria-sensing and killing machineries (34, 35).
890
A majority of the identified proteins have a known GO term
annotation (1124/1291, 87.1%) or can be matched in the
KEGG pathways (1288/1291, 99.8%). Significant GO terms
encompass the basic functions of the phagosome i.e. vesicle
trafficking, reorganization of the cytoskeleton, energy-depen-
Molecular & Cellular Proteomics 11.10
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MyoK Modulates the Early Delivery of ER to the Phagosome
TABLE I
Number and percentage of identified phagosomal proteins matching Gene Ontology (GO) terms and KEGG pathways
GO term:
GO:0015986:
GO:0009617:
GO:0009144:
GO:0030036:
GO:0006886:
GO:0016192:
ATP synthesis coupled proton transport
response to bacterium
purine nucleoside triphosphate metabolic process
actin cytoskeleton organization
intracellular protein transport
vesicle-mediated transport
Phagosome
Whole cell
Enrichment
14 (1.25%)
40 (3.56%)
29 (2.58%)
42 (3.74%)
41 (3.65%)
56 (4.98%)
16 (0.21%)
53 (0.71%)
47 (0.63%)
107 (1.43%)
109 (1.46%)
157 (2.10%)
5.82
5.02
4.11
2.61
2.5
2.37
KEGG pathways:
Phagosome
Whole cell
Enrichment
p value
Ribosome
Protein processing in endoplasmic reticulum
Protein export
Phagosome
Oxidative phosphorylation
Metabolic pathways
Endocytosis
Citrate cycle (TCA cycle)
Biosynthesis of secondary metabolites
Alanine, aspartate and glutamate metabolism
Proteasome
Glycolysis/Gluconeogenesis
42 (3.26%)
22 (1.71%)
11 (0.85%)
18 (1.40%)
30 (2.33%)
114 (8.85%)
19 (1.48%)
15 (1.16%)
40 (3.11%)
11 (0.85%)
14 (1.09%)
12 (0.93%)
86 (0.65%)
66 (0.50%)
17 (0.13%)
40 (0.30%)
66 (0.50%)
499 (3.75%)
39 (0.29%)
30 (0.23%)
174 (1.31%)
20 (0.15%)
38 (0.29%)
28 (0.21%)
5.05
3.44
6.69
4.65
4.70
2.36
5.03
5.17
2.38
5.68
3.81
4.43
⬍105
⬍105
⬍105
⬍105
⬍105
⬍105
⬍105
⬍105
⬍105
1 ⫻ 105
3 ⫻ 105
3 ⫻ 105
dent proton pumping, mainly because of phagosome acidification by the V-ATPase complex, and killing of the engulfed
bacteria (Table I). KEGG pathways stress that phagocytosis is
an endocytic process and plays a central role in nutrition and
cell metabolism. Unexpected is the significant enrichment of
ER-associated protein processing, protein export machineries and the proteasome in the phagosome (Table I). Indeed,
the ER-associated degradation (ERAD) machinery (sec61,
p97, Ufd1, Dsk2) and ER-Golgi intermediate compartments
(ERGIC; ERGIC53, COPII) represent a majority of the proteins
contributing to the “protein processing in the ER” pathway
(supplemental Fig. S2). In addition, ubiquitin mediated proteolysis, a pathway closely associated with the proteasome, is
also highly represented on the phagosome (p ⫽ 0.58).
The comparison of our phagosome proteome with the recently published Dictyostelium macropinosome proteome
(36) allowed us to identify pathways and proteins unique to
the phagosome. Both proteomes share 62% of identified
proteins (supplemental List S1) (36). General metabolism
(81%), lipid metabolism (75%), protein degradation (75%) and
biosynthesis (85%) and membrane trafficking (68%) are
mostly shared between the two organelles whereas receptors
(47%) and signaling (47%) are less shared than the average.
Indeed, both macropinosome and phagosome are used to
assimilate endocytosed nutrients but phagosome formation is
receptor-mediated, confirming our classification. Interestingly, the KEGG pathways “ER-associated protein processing, protein export machineries” and “proteasome” are also
significantly enriched on the macropinosome (p value ⬍ 10⫺5,
data not shown).
MyoK and Abp1 have Distinct Roles in Phagosome Maturation—The characterization of the phagosome proteome in
Molecular & Cellular Proteomics 11.10
wild type Dictyostelium was a prerequisite to analyze phagocytosis mutants at the molecular level. The phagocytic phenotypes of myoK- and abp1-null mutants have been characterized previously using rather big particles (yeasts or 4.5 ␮m
beads) compared with the size of Dictyostelium (10 ␮m diameter in suspension) (12, 13), but usually, smaller beads have to
be used to isolate phagosomes in sufficient quantities for
extensive immunoblot and 2D-DIGE analyses. Thus, the ability of both mutants to ingest beads of different sizes was
tested (Fig. 1C). In abp1-null cells, uptake of fluorescent latex
beads was increased, independently of bead size. In myoKnull cells, uptake was generally decreased for all bead sizes,
but only significantly for the 4.5 ␮m beads. Therefore, it confirms that Abp1 is a negative regulator of phagocytosis
whereas MyoK is a positive regulator essential for efficient
phagocytosis of large particles (12). Although the absence of
MyoK is not limiting for the uptake of smaller beads, our
experimental set up might still allow us to observe the signature of a defect in mutant cells. Indeed, the slow uptake of
single large particles is likely mimicked by the simultaneous
ingestion of a large number of medium-sized beads as is the
case during the initial pulse of our phagosome isolation
procedure (9).
To delineate the time frame of action of MyoK and its
binding partner Abp1 during maturation, their presence was
monitored by quantitative immunoblotting of phagosomes
isolated from wild type cells following our pulse/chase feeding
protocol. To further assess their functional inter-relationships,
their relative abundance was measured in phagosomes from
myoK-null and abp1-null mutants (Fig. 1D). The maturation
profiles of MyoK and Abp1 in phagosomes isolated from wild
type cells were similar but distinct. Both proteins were more
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MyoK Modulates the Early Delivery of ER to the Phagosome
FIG. 2. Absence of Abp1 alters the maturation profile of its direct binding partners. A, The MyoK-Abp1 interaction network as
characterized in (12) B–E, Maturation profiles of the indicated direct Abp1 binding-partners are affected in the abp1 null (red) cells but not in
myoK null (green) cells compared with wild type (black). Phagosomes were purified from the respective genetic background at the indicated
times. Immunoblotting signals were quantified and normalized to 100% according to the peak value in wild type. Curves are the average of at
least triplicate experiments. Error bars represent ⫾ S.E.
abundant on early than on later phagosomes but Abp1 remained associated longer with phagosomes. MyoK was significantly more abundant on early phagosomes from abp1null cells although its overall maturation profile was not
drastically altered. Abp1 showed both an increased abundance on early phagosomes from myoK-null cells and a decreased relative abundance at later time points. In conclusion,
Abp1 and MyoK are not necessary for their mutual recruitment on the phagosome, but the absence of one increases
the relative abundance of the other at early time points, confirming their distinct but mutually dependent roles in phagosome maturation.
Absence of Abp1 Alters the Maturation Profile of its Direct
Binding Partners—To delineate the impact of myoK- or abp1null mutations on maturation, the relative abundance profiles
in wild type and mutants of the direct binding partners of
MyoK and Abp1 (schematically presented in Fig. 2A) were
compared by quantitative immunoblotting. Absence of Abp1
but not MyoK markedly altered the maturation profile of its
direct binding partners. Wild type phagosomes were highly
enriched in actin at early stages only. In contrast, the relative
abundance of actin did not vary significantly during maturation in abp1-null cells (Fig. 2B). However, it is not clear if this
signal corresponds to actin monomers or polymers, because
the signal was under the detection threshold for staining with
the fluorescent probes DNase I and phalloidin, binding respectively G- and F-actin, or indirect immunofluorescence
against the G-actin binding protein, profilin II. Furthermore,
abp1 mutation markedly delayed the maximum peak of rela-
892
tive abundance of profilin II (Fig. 2C) or Arp3 (Fig. 2D). Dynamin A also remained associated longer with abp1-null phagosomes (Fig. 2E). Therefore, the absence of Abp1 directly
impacts on the abundance of actin and proteins involved in
actin dynamics on the phagosome whereas this is not the
case in absence of MyoK.
2D-DIGE Reveals An Early Maturation Defect in myoK Null
Cells and an Overall Maturation Defect in abp1-null Cells—
Defects in the maturation profiles of individual proteins were
readily apparent using targeted immunoblot profiling. To expand the analysis, in depth characterization of the phagocytic
phenotypes of myoK- and abp1-null mutants was carried out
with an unbiased 2D-DIGE cross-comparison. Differences in
the relative abundance of proteins were visible between phagosomes of wild type and mutant cells. These differences
were not only seen early in maturation but were also consistent throughout maturation as, for example, for the individual
subunits (VatA, VatG and VatB) of the V-ATPase complex
(Figs. 3A, 3B). Phagosomes from each maturation stage were
isolated in quadruplicate from wild type cells and in duplicate
from each mutant cell line. Thus, each of the six time-points in
the wild type was compared in duplicate to its corresponding
time point in the mutant, with dye inversion, corresponding to
24 gels in total (Fig. 3C and supplemental Fig. S3). Gels were
first matched between biological replicates of identical timepoints to obtain a reliable quantitative mutant-to-wild type (wt)
spot ratio. Such ratios were determined for about 200 to 600
spots pairs per time point (Fig. 3D). The analysis of myoK-null
phagosomes revealed a higher proportion (17%) of differential
Molecular & Cellular Proteomics 11.10
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MyoK Modulates the Early Delivery of ER to the Phagosome
FIG. 3. Characterization of the phagocytosis defect of Dictyostelium myoK null and abp1 null mutants by quantitative 2D-DIGE
analysis of isolated phagosomes. A, Differences were visible in phagosomes isolated at early times (5⬘/0) from wild-type compared with
myoK null cells . Differential spots with increased or decreased intensity in the mutant are highlighted in green (increased) or blue (decreased).
The spots containing the VatA, VatB, and VatG subunits of the V-ATPase are circled in red. B, A representative gallery of these spots highlights
variations in spot intensity during maturation in wild type compared with myoK null. C, Workflow of the approach used to quantify
Molecular & Cellular Proteomics 11.10
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MyoK Modulates the Early Delivery of ER to the Phagosome
spot pairs early (5⬘/0) and fewer differences later during maturation (4 –12%, average ⬃7%) (Figs. 3B–3D). In contrast, the
number of differential spots in abp1-null phagosomes was
higher and rather constant (11–18%, average ⬃13%) (Fig.
3E). Therefore, 2D-DIGE analyses identify an early maturation
defect in myoK-null cells but an overall perturbation of maturation in abp1-null cells.
In order to identify general cellular defects caused by gene
knockout and evaluate their impact on phagosome maturation, total cell lysates of both mutants were compared with
wild type by 2D-DIGE. Two spots were differential in the
myoK-null mutant, representing less than 0.5% of all spots,
and contained two proteins of unknown function (supplemental Table S1). Eight spots were differential in the lysate of
abp1-null cells, representing 1% of all spots, and spanned
various functions at all levels of cell metabolism (supplemental
Table S2). The impact of the mutations on the whole cell is
very limited (ⱕ 1% differential spots) in comparison to the
impact on phagosome maturation (4 –18% differential spots).
Nevertheless, the mutation of abp1 affects more profoundly
cell metabolism than does the mutation of myoK, confirming
the phagosome analysis.
To monitor the variation of the mutant-to-wt ratio during
maturation, spots pairs were then matched across the whole
time-course resulting in time-resolved ratio profiles (Fig. 3C).
70% of all spot pairs (ⱖ 1000) matched between biological
replicates were matched to at least another pair across the
time-course. The 150 spot pairs were matched to at least five
time-points resulting in a complete ratio profile. Between 200
(myoK-null) and 300 additional (abp1-null) pairs were matched
across at least three time-points. Gel references were chosen
in the early time-points (5⬘/0) to match myoK-null to abp1-null
gel series so that mutant-to-wt ratio profiles could be compared across mutants. Abundant spots and most differential
spots were picked from the gel and identified by mass spectrometry-based protein sequencing. The 104 proteins were
identified, representing an average of 220 spot pairs with
known ratio profiles over the two mutant-to-wt series.
myoK-null Phagosomes Display an Early Imbalance in the
Abundance of ER Components and the V-ATPase Complex—
2D-DIGE intensity profiles were first confirmed by immunoblot
profiling. Using both detection techniques, the wild type profiles of the ER proteins, calreticulin and protein disulfide
isomerase, showed a maximum enrichment in early phagosomes and then a progressive decrease during maturation
(Figs. 4A, 4B). These profiles implied that ER-derived membranes fuse with the phagosome just after cup closure and
ER-associated proteins are then removed later during
maturation.
When comparing phagosomes from wild type and myoKnull cells, prominent differences were seen in the abundance
profiles of both ER proteins and subunits of the V-ATPase
complex (Figs. 3B and 4B). Measured ratios of the quantified
spot intensities confirmed these observations. Mutant-to-wt
ratio profiles of all identified ER proteins (Fig. 4C) were sufficiently homogenous to unequivocally group together. Similar
grouping was also obvious for the ratio profiles of the soluble
subunits of the V-ATPase complex (Fig. 4D). Ratio profiles of
individual proteins fitted so well that a representative average
ratio profile could be drawn for both functional groups (black
line). A significant increase (1.6-fold, log2 ⬎ 0.68) in the abundance of ER proteins and a significant decrease (1.6-fold,
log2 ⬍ 0.68) in the abundance of the V-ATPase subunits were
observed in early phagosomes (5⬘/0 min.) from myoK-null
cells. Strengthening this early deviation from the wild type
maturation profile, the ratio profile of both groups showed
reciprocal variations during maturation. This revealed that
early imbalance of ER versus V-ATPase proteins was then
progressively compensated during maturation. Other proteins
were also functionally grouped according to their ratio profile,
like the phagosomal chaperones and the different actin isoforms (Figs. 5A, 5B, details in supplemental Fig. S4). Unlike
the ER and V-ATPase functional groups, the actin and chaperones groups were not significantly differential in early phagosomes and their profile was not linked either to the ER or
V-ATPase group in the myoK-null background (Fig. 5A). Neither ER nor V-ATPase proteins were differential in abp1-null
phagosomes. Therefore, significant differences in early abundance of ER and V-ATPase proteins on the phagosome and
their reciprocal ratio profile variation were specific to the
myoK-null to wild type comparison.
Strikingly, changes in the myoK-null to wild type ratio profile
of the small GTPases Rab11A and Rab11C anticipate
changes in the profile of the V-ATPase complex, in both time
and amplitude changes (Fig. 5). Furthermore, the regulation or
the kinetics of these small GTPases on myoK-null phagosomes might depend on their post-translational modifications
(supplemental Fig. S5).
The Relative Abundance of Lysosomal Enzymes is Altered in
Early abp1-null Phagosomes—Relative abundance of lysosomal enzymes was specifically increased in early (15⬘/0)
abp1-null phagosomes (2.5–3.9-fold, Fig. 6). This increase
was transient as minimal difference was observed at later
time-points. Chaperones of the hsp70 family (HspE) and
mutant-to-wild type spot intensity ratios during the maturation process. D, The relatedness of the phagosome maturation program in wild type
and mutants is represented by the number of differential versus nondifferential phagosomal spots at each time point in myoK null (left) and abp1
null mutants (right). Differential spots are in color (detailed view in Fig. 3E) whereas non-differential spots are in gray (myoK null) and black (abp1
null). E, The variation of spots differential during maturation in mutants is expressed as a percentage of total matched spots. The left panel
shows spots decreased (green) or increased (orange) in myoK null phagosomes. The right panel shows spots decreased (red) or increased
(violet) in abp1 null phagosomes.
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Molecular & Cellular Proteomics 11.10
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MyoK Modulates the Early Delivery of ER to the Phagosome
FIG. 4. The ratio profiles of ER components and V-ATPase subunits are inversely correlated when comparing phagosomes from
myoK null and wild type cells. A, Curves from both quantitative immunoblotting and 2D-DIGE intensity profiling of calreticulin (crtA) and
protein disulfide isomerase (pdi) isoforms in phagosomes from wild-type cells were highly similar, with a maximum early in maturation (5⬘/0).
For immunoblot and 2D-DIGE, the peak intensity value of the profile was set to 100% and the rest of the curve was normalized accordingly.
In immunoblots, curves are based on triplicates and error bars represent ⫾ S.E. 2D-DIGE profiles represent duplicates. B, Immunoblot profiles
and 2D-DIGE spot images of the maturation sequence of calreticulin in phagosomes from wild-type and myoK null cells. C, D, myoK
null/wild-type ratio profiles in log2 scale of normalized spot intensities of identified ER proteins (C) and of subunits of the v-ATPase complex
(D). The profiles of all ER proteins and V-ATPase subunits identified in myoK-null phagosomes are shown here. The profiles were so similar that
an averaged profile of the corresponding functional groups was computed and drawn in black. Only gene names (www.dictybase.org) are
indicated in the legend. ER proteins: crtA, calreticulin A; cnxA, calnexin A; grp78 and grp94, glucose regulated protein of 78 kDa and 94 kDa;
pdi1 and 2, protein disulfide isomerase isoform 1 and 2; soluble v-ATPase complex subunits vatA, B, E, G, H.
hsp40 family (ddj1, DnaJC7) were also clearly differential (1.7–
3.1-fold) from the 15⬘/15⬘ time point across the whole phagosomes maturation process in abp1-null cells (Fig. 5B, details
in supplemental Fig. S4).
Functional Impact of the Trafficking Defects Detected by
2D-DIGE on Phagosomal pH and Proteolytic Activity—In order
to monitor in vivo the functional impact of the severe trafficking defects revealed by ratio profiling, we adapted to Dictyostelium the use of intra-phagosomal biosensors developed by
Russell and colleagues (57, 58). Such approaches based on
dual wavelength fluorescence intensity ratio measurements
allow to quantitatively measuring environmental parameters
such as pH and proteolysis. Compared with the acidification
profile obtained in wild type cells, in which the pH minimum of
⬃4.5 was reached after about 40 min and the re-neutralization
plateau reached after about 70 min, the acidification of phagosomes in myoK-null cells was significantly delayed. (Fig.
7A, 7D). It also reached a pH of ⬃4.5, but after about 50 min
and the neutralization plateau not before about 100 min. As a
likely consequence, the onset of proteolytic activity was
slightly delayed but it also progressed at a severely reduced
but almost linear rate (Figs. 7B, 7D). Note that the proteolytic
Molecular & Cellular Proteomics 11.10
activity in wild type phagosomes was also subjected to a
decrease of rate after about 70 min, corresponding to the time
taken to reach the neutralization plateau (Fig. 7C). Strikingly,
in abp1-null cells, phagosomes reached a more acidic value
of 4.0 earlier, at about 30 min, the re-neutralization phase
started earlier but led to a plateau at a time (70 min) similar as
in wild type cells (Figs. 7A, 7C). The proteolytic activity monitored in abp1-null cells was strongly affected (Figs. 7B, 7E).
The onset of activity occurred slightly earlier than in the two
other cell lines, and initially progressed at a higher rate, but
after about 30 min, this rate decreased significantly, corresponding to the start of the neutralization phase.
Overall, these functional results are in agreement with the
2D-DIGE ratio profiling data documenting the transient initial
deficit in V-ATPase delivery in myoK-null cells and the transient initial overabundance of lysosomal enzymes in abp1-null
cells.
DISCUSSION
Before engaging on an exhaustive comparative and quantitative analysis of phagosome maturation in Dictyostelium
myoK-null and abp1-null mutants, we characterized the pu-
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MyoK Modulates the Early Delivery of ER to the Phagosome
FIG. 5. The early trafficking imbalance of ER components versus V-ATPase subunits is specific to myoK-null phagosomes and
might be regulated by Rab11 small GTPases. A, B, myoK null/wild
type (A) or abp1 null/wild type (B) averaged ratio profiles in log2 scale
for ER proteins, V-ATPase, actin and hsp40/hsp70 chaperones show
that the profiles of ER versus V-ATPase are only inversely correlated
when comparing myoK null to wild type phagosomes. Proteins included in the ER and V-ATPase groups are listed in Figs. 4C and 4D,
respectively. The chaperone group includes all hsp40/hsp70 chaperones identified by 2D DIGE (hsp40-like chaperones, ddj1 & dnaJC7;
hsc70-like chaperone, hspE). The actin group includes all the spots
where actin isoforms were identified, excluding degradation products. C, Averaged ratio profiles for Rab11A and Rab11C small
GTPases isoforms anticipate changes in the averaged profile of the
V-ATPase complex, in both time and amplitude on myoK null phagosomes. The profiles of all Rab11A and Rab11C isoforms have been
averaged (details in supplemental Fig. S5).
rity, the morphology and the composition of phagosomes
isolated from wild type cells (Fig. 1). Observation of intraphagosomal vesicles and formation of large multi-bead phagosomes during maturation highlight that phagosome plasticity
is achieved not only through classical vesicle fission from and
fusion to the phagosome, but also via the formation of intralumenal vesicles and phagosome-phagosome fusions. Phagosomes not only shares 62% of their protein composition with
macropinosomes (36) but also morphological features with
896
the pinocytic endosomal pathway as changes of intraphagosomal vesicles during maturation are similar to those observed in fluid-phase endosomes (37). Intralumenal vesicles
have already been observed in macrophage phagosomes
containing latex beads or red blood cells (38). These vesicles
might result from the action of the ESCRT pathways (39) or
from fusion with autophagosomes containing cytoplasmic
material (40).
The analysis of phagosome composition revealed several
proteins involved in lipid metabolism, indicating that the phagosome lipid composition might be dynamically modified to
participate in membrane remodeling and vesicle trafficking
(Fig. 1B). Comparison with the macropinosome proteome (36)
emphasized the fact that the pathways for ER-associated
protein processing, protein export machineries and the proteasome are significantly enriched in both proteomes (Table I).
Given the differences in the purification procedures of both
organelles and their significant enrichment, it strongly suggests that ER components and the proteasome are not
mere contaminants in the biogenesis of both phagosome and
macropinosome.
Detailed analysis of the phagocytic phenotypes of myoKand abp1-null cells confirmed that Abp1 is a negative regulator of phagocytosis whereas MyoK is a positive regulator
determinant for phagocytosis efficiency of large particles only
(Fig. 1C). The analysis of maturation profiles shows that Abp1
and MyoK are not necessary for their mutual recruitment on
the phagosome and, in addition, that the absence of one
increases the relative abundance of the other (Fig. 1D). These
observations confirm that MyoK and Abp1 have opposite
regulatory roles in phagocytosis (12).
2D-DIGE analysis of phagosomes showed that myoK-null
cells display an early maturation defect whereas an overall
perturbation of maturation is observed in abp1-null cells. The
amplitude of the defects correlates well with their respective
abundance during phagosome maturation in wild type cells
(Figs. 3D, 3E). These defects also correlated with differences
in the cellular transcriptome (data not shown) and proteome
(supplemental Tables S1, S2). The impact of the mutations is
on average ten times bigger on phagosome maturation than
on general cell metabolism, as judged by the number of
differential spots in phagosomes compared with whole cell
lysates.
We had previously shown that quantitative analysis of maturation profiles, by Western blotting and two-dimensional gels,
allowed hierarchical clustering of functional groups based on
profile similarities (10). Here, we show that comparative 2DDIGE analysis of phagosome maturation in mutant cells resulted
in the quantitative monitoring of the mutant to wild type ratio
profiles of more than one hundred identified proteins. Strikingly,
ratio profiles of functionally related proteins showed coordinated changes during maturation. These changes were specific
for each of the two mutants examined, thus validating the functional classification of phagosomal proteins.
Molecular & Cellular Proteomics 11.10
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MyoK Modulates the Early Delivery of ER to the Phagosome
FIG. 6. Ratio profiles of the lysosomal enzymes in abp1-null phagosomes highlight a trafficking defect in early maturation. Ratio
profiles of the lysosomal enzymes (A, B) from phagosomes of myoK-null cells (A) or abp1-null cells (B). Lysosomal enzymes: cathepsin D (ctsD,
DDB_G0279411), cysteine protease 1-like (CP1-like, DDB_G0291191), lytic enzyme (DDB_G0293566), a/b hydrolase (DDB_G0287609), mix of
putative physaropepsin, acyloxyacyl hydrolase, peptidase V4 –7 (DDB_G0290333, DDB_G0272785, DDB_G0281823). Cathepsin D is not
present in the gels of one of the time-series of abp1-null phagosomes. Therefore, it does not fulfill our criteria of significance and is not
represented in the left panel. CinB is an esterase/lipase with its closest mammalian homolog (UniProt ID: NCEH1_MOUSE) being a cholesterol
esterase present in the ER. It is represented here as a counterexample.
FIG. 7. myoK-null cells are defective in phagosomal acidification and proteolytic digestion whereas abp1-null cells display a
sudden decrease in phagosomal proteolytic digestion during
maturation. Measure of pH variation (A) and proteolytic digestion (B)
during maturation in phagosomes in vivo in AX-2 (black), myoK-null
cells (green), and abp1-null cells (red). Curves are averaged from at
least three experiments in duplicates and error bars represent ⫾ S.E.
Our study shows that Abp1 has pleiotropic effects on phagosome maturation. Its absence transiently increases phagosomal proteolytic activity paralleling an early decrease in pH
(Fig. 7E) and the transient early accumulation of lysosomal
enzymes (Fig. 6B). This increased abundance of lysosomal
enzymes in the phagosome is either due to increased delivery
or to impaired retrieval. The causes for this early defect in
trafficking efficiency could originate from an accumulation of
actin and other direct binding partner of Abp1 on the phagosome as shown by immunoblot profiling (Fig. 2). Other data
Molecular & Cellular Proteomics 11.10
support this hypothesis. Abp1 is the major F-actin binding
player on phagosomes isolated from Dictyostelium in vitro
and its absence might disorganize the dynamics of the actin
network on the phagosome (18). Indeed, an increase in phagocytosis rates in knockdown of human Rab27a correlates with
a quicker turnover of F-actin coating and degradation at the
phagocytic cup (41). Furthermore, persistence of transient
actin flashes on the phagosome prevents fusion with lysosomes (22). Unexpectedly, the early trafficking defect of lysosomal enzymes in abp1-null phagosomes is followed by a
decrease in proteolytic activity. Therefore, not only a decrease
in pH but also a precisely coordinated trafficking of lysosomal
enzymes regulates phagosomal proteolysis.
In the myoK-null mutant, 2D-DIGE temporal ratio profiling
clearly shows an early imbalance of ER components versus
V-ATPase subunits (Figs. 4C, 4D, Fig. 5A). In addition, ratio
profiles of ER proteins and the V-ATPase complex are reciprocal. According to wild type intensity profiles, ER proteins are
delivered early to the phagosome to be recycled later suggesting that ER fusion occurs just after phagosome closure (Figs.
4A, 4B). ER proteins and the V-ATPase complex are not delivered from the same organelle, because the largest amount of
V-ATPase complexes is localized on the contractile vacuole and
V-ATPase complexes are delivered from endosomes and uncharacterized vesicles in Dictyostelium (4). Thus, we interpret
this imbalance in protein abundance as an early trafficking
imbalance to the phagosome in myoK-null cells.
Our data raise the following essential questions. Which
proteins regulate the delivery of the V-ATPase complex to
the phagosome? Why are ER proteins delivered early to the
phagosome in Dictyostelium and do our data correlate with
existing hypotheses? Is an imbalance in the delivery of ER
and V-ATPase to the phagosome a conserved phenomenon
and how is MyoK implicated in this trafficking event?
Potential candidates regulating the trafficking of the VATPase complex are the canonical endosomal GTPases
897
144
MyoK Modulates the Early Delivery of ER to the Phagosome
Rab7A and Rab5A. Both proteins have been identified in
phagosomal samples (this study) and could participate in the
delivery of the V-ATPase from endolysosomes. On the other
hand, our 2D-DIGE ratio profiling study suggests that the
small GTPases Rab11A, Rab11C and RabC could participate
in the delivery of the V-ATPase from an uncharacterized vesicle pool. Changes in the myoK-null to wild type ratio profiles
of the Rab11A, Rab11C and RabC small GTPases as well as
calmodulin precede, in time and amplitude, the changes in the
ratio profile of the V-ATPase complex (supplemental Fig. S5).
Like the V-ATPase complex and calmodulin, Rab11A and
Rab11C are mainly localized to the contractile vacuole (4,
42– 44), and Rab11A is important for the maintenance of the
contractile vacuole function and morphology (43). Rab11C
and RabC have not been functionally characterized and RabC
has no close homolog in any other organism (45). The small
GTPase Rab14 was also identified on phagosomes (this
study) and might be additionally involved in this process.
Indeed, Rab14 localizes both to the contractile vacuole and
the endosomal system, where it controls lysosome and phagosomes homotypic fusion (46).
The fusion with ER-derived membranes is an early event of
phagosome maturation, and these components are then either recycled or degraded during maturation (Fig. 4, (47, 48)).
However, the contribution of ER-derived proteins to phagosome function and the mechanisms driving their delivery are
still unclear both in Dictyostelium and animal phagocytes (5,
49). Three working hypotheses might be invoked. First, focal
exocytosis of endomembranes including the ER might be
crucial to compensate for the loss of cell surface area during
particle uptake (7, 50). Our data do not fit with a contribution
of ER membranes to enhance efficiency of uptake, because
the myoK-null mutant which displays an increase in the delivery of ER components does not exhibit increased uptake of
any big-sized particles (Fig. 1C) (13). Second, cortical or phagosomal ER patches could trigger a periphagosomal increase
in free calcium concentration, regulating focal exocytosis or
local signaling cascades (6, 51, 52). The decreased phagocytic uptake of calnexin/calreticulin double knock out mutants
was interpreted as the involvement of calcium signaling in
efficient uptake (53). Nevertheless, enhancing contact of the
ER with the phagosome does not enhance uptake in myoKnull mutants. Therefore, the extent and duration of the contact
between the ER and the phagosome and how long this contact takes place has to be tightly regulated to optimize the
efficiency of uptake. Finally, recruitment of ER proteins to the
phagosome in macrophages and dendritic cells parallels an
increase in cross-presentation of exogenous antigens, a process controlled by Sec22b, a SNARE present on ERGICs (5,
48, 54). This increase in cross-presentation is linked to a slow
initial decrease in pH in dendritic cells (55). Although amoebae
use phagocytosis to feed and do not express antigen-presenting molecules, molecular machines essential for crosspresentation like ER chaperones, ERGIC and ERAD compo-
898
nents are enriched in the phagosome of Dictyostelium (Table
I) and have been identified in a recent comparative proteomic
analysis of the phagosome in mouse, Drosophila and Dictyostelium, showing that these components have been conserved throughout evolution as parts of the phagosome proteome core (1). It was recently proposed that, during evolution
from prokaryotes to eukaryotes, the ER, the ERAD and the
proteasome machineries primarily served a function linked to
feeding, i.e. import and cleavage of polypeptides resulting
from extracellular digestion (56). Thus, ER components could
have been first used to assist feeding and then co-opted to
maximize efficiency of antigen cross-presentation.
The balance in the delivery of ER components and of the
V-ATPase complex to the phagosome can be genetically
altered in Dictyostelium by knocking out myoK. This balance
can be also altered pharmacologically in macrophages. Specific inhibition of the V-ATPase complex by bafilomycin or
inhibition of early membrane trafficking by 3-methyladenine
and wortmannin increases interaction of the ER with the phagosome (5). Thus, reducing V-ATPase activity or blocking
early membrane delivery might increase ER fusion. In dendritic cells, the balance between ER fusion and lysosome
delivery can be altered genetically. Absence of Sec22b delays
fusion of lysosomes to latex bead containing phagosomes
and reduces phagosomal proteolysis (48). In the myoK-null
mutant, the imbalance is likely because of a slowed delivery of
V-ATPase versus ER membranes as the abundance of the
V-ATPase in the mutant reaches wild type levels at 15⬘/15⬘
and even exceeds it beyond this time point (Fig. 4D). Because
MyoK has been localized at the phagocytic cup and on early
phagosomes [(12) and Fig. 1D], we suggest that it might
directly control retrieval of ER components rather than VATPase delivery. In analogy, another Dictyostelium class I
myosin, MyoB, controls the retrieval of plasma membrane
components from the endosomes (14). Alternatively, MyoK
could indirectly alter membrane trafficking. MyoK is enriched
at sites of cup closure and its absence lowers phagocytic
rates (12) (Fig. 1C). Therefore, its absence potentially slows
down cup closure, which would increase contact of ER membranes with the phagocytic cup. This might in turn hinder
contact and delay fusion with a Rab11-positive endosomal
compartment delivering the V-ATPase. In the myoK-null mutant, the reduced delivery of V-ATPase correlates with a reduced phagosomal acidification in vivo (Fig. 7A). As a likely
consequence, the intra-phagosomal proteolytic activity is severely decreased (Fig. 7B). Thus, we propose that the efficiency of particle digestion and killing reflects this balance
between fusion with the ER and delivery of the V-ATPase
complex.
In conclusion, quantitative ratio profiling by immunoblots
and 2D-DIGE is a powerful tool to measure mutant to wild
type protein abundance ratios over the complete course of
the maturation process. Targeted immunoblot profiling pinpointed the central role of Abp1 in organizing actin dynamics
Molecular & Cellular Proteomics 11.10
145
MyoK Modulates the Early Delivery of ER to the Phagosome
on the phagosome. Quantitative and comparative 2D-DIGE
profiling was a reliable discovery tool to highlight the proteins
that deviate from the wild type maturation profiles induced by
genetic ablations. Clustering of such deviant profiles revealed
concerted alterations in the abundance of functional protein
groups. As a proof of concept, this approach exposed for the
first time that the participation of ER components is part of the
primordial process of maturation and that a reciprocal imbalance in the delivery of ER and V-ATPase components is
induced genetically by knocking out the myosin IK and, consequently, impacts on the kinetic of acidification and proteolysis. Our study brings a genetic and proteomic demonstration
that the ER participation in phagocytosis is an evolutionary
conserved process regulated by proteins at the interface between cytoskeleton and membrane traffic.
Acknowledgments—We thank A. Bosserhof and M. Ellis for help
with mass spectrometry, N. Gopaldass and N. Sattler with the pH and
proteolysis measurement assays, and M. Hagedorn with electron
microscopy. A special thank you goes to P. Gaudet, M.-C. Blatter,
and E. de Castro from the SIB Swiss Institute of Bioinformatics for
their help with bioinformatic analyses. We are also grateful to the
whole team of DictyBase for their support. We thank Matthias Trost,
Jonathan Boulais, and Florence Niedergang for careful reading of the
manuscript and thoughtful suggestions.
* This work was supported by the Max-Planck Society, the Deutsche Forschungsgemeinschaft, the UK BBSRC, and the Swiss National Science Foundation.
□
S This article contains supplemental Figs. S1 to S5 and Tables S1
to S4.
储 To whom correspondence should be addressed: Départment de
Biochimie, Faculté des Sciences, University de Genève, Sciences II,
30 quay Ernest Ansermet, CH-1211 Genève-4, Switzerland. Tel.:
⫹41-22-379-6496;
Fax:
⫹41-22-379-3499;
E-mail:
[email protected].
** Present address: Klinisches Institut für Pathologie, 1090 Wien,
Austria.
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Molecular & Cellular Proteomics 11.10
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5.3-WASH is required for lysosomal recycling and efficient autophagic and phagocytic digestion
Contributed Figure 3, supplementary Figure 1A and supplementary table 1
148 MBoC | ARTICLE
WASH is required for lysosomal recycling and
efficient autophagic and phagocytic digestion
Jason S. Kinga,*, Aurélie Guehob, Monica Hagedornc, Navin Gopaldassb,†, Florence Leubab,
Thierry Soldatib, and Robert H. Insalla
a
Beatson Institute for Cancer Research, Bearsden, Glasgow G61 1BD, United Kingdom; bDepartment of Biochemistry,
University of Geneva, CH-1211 Geneva, Switzerland; cBernhard Nocht Institute for Tropical Medicine, 20359 Hamburg,
Germany
ABSTRACT Wiskott-Aldrich syndrome protein and SCAR homologue (WASH) is an important
regulator of vesicle trafficking. By generating actin on the surface of intracellular vesicles,
WASH is able to directly regulate endosomal sorting and maturation. We report that, in
Dictyostelium, WASH is also required for the lysosomal digestion of both phagocytic and
autophagic cargo. Consequently, Dictyostelium cells lacking WASH are unable to grow on
many bacteria or to digest their own cytoplasm to survive starvation. WASH is required for
efficient phagosomal proteolysis, and proteomic analysis demonstrates that this is due to
reduced delivery of lysosomal hydrolases. Both protease and lipase delivery are disrupted,
and lipid catabolism is also perturbed. Starvation-induced autophagy therefore leads to
phospholipid accumulation within WASH-null lysosomes. This causes the formation of multilamellar bodies typical of many lysosomal storage diseases. Mechanistically, we show that, in
cells lacking WASH, cathepsin D becomes trapped in a late endosomal compartment, unable
to be recycled to nascent phagosomes and autophagosomes. WASH is therefore required for
the maturation of lysosomes to a stage at which hydrolases can be retrieved and reused.
Monitoring Editor
Carole Parent
National Institutes of Health
Received: Feb 14, 2013
Revised: Jun 5, 2013
Accepted: Jul 1, 2013
INTRODUCTION
The Wiskott-Aldrich syndrome protein and SCAR homologue (WASH)
is an evolutionarily conserved regulator of the Arp2/3 complex. Like
other members of the Wiskott-Aldrich syndrome protein (WASP)
This article was published online ahead of print in MBoC in Press (http://www
.molbiolcell.org/cgi/doi/10.1091/mbc.E13-02-0092) on July 24, 2013.
Present addresses: *Department of Biomedical Sciences, Firth Court, Sheffield
University, Sheffield S10 2TN, United Kingdom; †Département de Biochimie,
Université de Lausanne, Chemin des boveresses 155, CH-1066 Epalinges,
Switzerland.
Address correspondence to: Jason S. King ([email protected]).
Abbreviations used: BSA, bovine serum albumin; catD, cathepsin D; CD, cation
dependent; CI, cation independent; CID, collision-induced dissociation; EGF,
epidermal growth factor; EGTA, ethylene glycol tetraacetic acid; FITC, fluorescein
isothiocyanate; GFP, green fluorescent protein; HCD, high-energy C-trap dissociation; M6PR, mannose-6-phosphate receptor; MLB, multilamellar body; MS,
mass spectrometry; NA, numerical aperture; PBS, phosphate-buffered saline;
PDI, protein disulfide isomerase; siRNA, small interfering RNA; TCEP, Tris(2-carboxyethyl) phosphine hydrochloride; TEAB, triethylammonium hydrogen
carbonate buffer; TEM, transmission electron microscopy; TMT, tandem mass
tag; v-ATPase, vacuolar ATPase; WASH, Wiskott-Aldrich syndrome protein and
scar homologue; WASP, Wiskott-Aldrich syndrome protein.
© 2013 King et al. This article is distributed by The American Society for Cell Biology under license from the author(s). Two months after publication it is available
to the public under an Attribution–Noncommercial–Share Alike 3.0 Unported
Creative Commons License (http://creativecommons.org/licenses/by-nc-sa/3.0).
“ASCB®,” “The American Society for Cell Biology®,” and “Molecular Biology of
the Cell®” are registered trademarks of The American Society of Cell Biology.
2714 | J. S. King et al.
family, the primary role of WASH is to activate Arp2/3 and regulate
the spatial and temporal formation of actin networks (Linardopoulou
et al., 2007). Specifically, WASH generates actin subdomains on endocytic vesicles and is important for several sorting and maturation
steps. These include retrograde transport from endosomes to the
trans-Golgi via direct regulation of the retromer (Derivery et al.,
2009; Gomez and Billadeau, 2009; Duleh and Welch, 2010; Harbour
et al., 2010) and the sorting and trafficking of other signaling molecules, such as integrins, the epidermal growth factor (EGF), and
transferrin receptors (Derivery et al., 2009, Zech et al., 2011).
A comparable role has also been described in Dictyostelium,
whereby WASH-generated actin sequesters the vacuolar (v)-ATPase
to late endosomal subdomains, leading to v-ATPase removal via
recycling vesicles (Carnell et al., 2011). The v-ATPase is the proton
pump responsible for establishing and maintaining an acidic pH in
the lumen of lysosomes. Therefore its WASH-mediated removal
leads to the neutralization and maturation of lysosomes into
postlysosomes prior to exocytosis (Neuhaus et al., 2002). In
Dictyostelium, the loss of WASH therefore arrests lysosome maturation before v-ATPase removal, blocking both neutralization and
exocytosis (Carnell et al., 2011).
Like other WASP-family proteins, WASH acts as part of a complex (Jia et al., 2010; Veltman and Insall, 2010). Our recent work
Supplemental Material can be found at:
http://www.molbiolcell.org/content/suppl/2013/07/23/mbc.E13-02-0092v1.DC1.html
Molecular Biology of the Cell
149
showed that, while disruption of the Strumpellin, ccdc53, or SWIP
subunits leads to total loss of WASH activity, the FAM21 subunit has
a unique role, driving the recycling of WASH (Park et al., 2013).
Therefore loss of FAM21 gives hyperactive WASH, and, although
cells retain the ability to recycle v-ATPase, endocytic maturation is
blocked at a later stage, at which WASH is itself recycled. Importantly, these contrasting phenotypes of WASH and FAM21 mutants
now allow us to dissect the pathway in more detail and discriminate
direct and indirect effectors of WASH.
In addition to its specific trafficking roles, WASH appears to be
more generally important in maintaining endosomal and lysosomal
integrity. In mammalian cells, small interfering RNA (siRNA) leads to
altered endosome morphology (Derivery et al., 2009; Duleh and
Welch, 2010), while full deletion causes lysosomal collapse (Gomez
et al., 2012; Piotrowski et al., 2012). We therefore hypothesized that
WASH may also be important for normal lysosomal function.
Lysosomes are responsible for the degradation of endosomal
contents. They are therefore essential for breaking down endocytic
cargo, such as membrane receptors, as well as the contents of
autophagosomes and bacteria captured within phagosomes.
Autophagy is the name given to a family of pathways leading to the
capture of cytoplasmic constituents and their delivery to lysosomes.
This includes both microautophagy, wherein selected proteins are
directly transported across the lysosomal membrane, and macroautophagy, wherein double-membrane vesicles are formed de novo,
capturing parts of the cytoplasm and fusing with lysosomes (Xie and
Klionsky, 2007). In this study, we have specifically studied macroautophagy, which we will simply refer to as autophagy for clarity.
Cells use autophagy for a number of purposes, and defective
autophagy is associated with a number of medical conditions, including aging, cancer, infection, and neurodegeneration (Mizushima
et al., 2008). The best-understood function of autophagy is during
starvation, when degradation of the cytosol provides nutrients and
energy to the cell, enabling survival. In professional phagocytes
such as Dictyostelium, this is similar to the phagocytic pathway, in
which the cell uses the same digestive pathway to liberate nutrients
from captured bacteria (Deretic, 2008). Autophagy and phagocytosis therefore both require lysosomal delivery and digestion in order
to feed the cell.
The WASH complex has been assigned a number of roles in vesicular trafficking. In this study, we assess the importance of WASH
in lysosomal function within the context of both the autophagic and
phagocytic pathways. We show that WASH is required for both processes in Dictyostelium and describe a new role for WASH in maintaining lysosomal flux.
RESULTS
WASH is not required for autophagosome formation
Previous work has shown that the WASH complex is involved in multiple stages of the endocytic pathway. Autophagy intersects with the
endocytic pathway and relies on many of the same processes, such
as v-ATPase and lysosomal hydrolase trafficking. We therefore directly assessed the role of WASH in autophagy, using the model
system Dictyostelium discoideum, for which there are well-defined
assays for autophagosome induction and formation (Calvo-Garrido
et al., 2010, King et al., 2011).
To directly observe autophagosome formation, we used a green
fluorescent protein (GFP)-Atg8 fusion probe. During autophagosome formation, GFP-Atg8 is lipidated and incorporated into the
expanding phagophore membrane (Ichimura et al., 2000) and can
therefore be used to label autophagosomes until they are fully
formed. When the autophagosomes are completed, the cytosolically
Volume 24 September 1, 2013
exposed GFP-atg8 is cleaved by Atg4, the pH-sensitive GFP fluorescence on the interior is quenched by acidification, and the GFP
signal is lost (Kirisako et al., 2000). Using cellular compression to
both induce autophagy and improve imaging (King et al., 2011),
we observed no gross defects in autophagosome formation or
morphology in WASH-null cells compared with wild-type (Ax2)
controls (Figure 1A and Supplemental Movies S1 and S2).
Previous work in Dictyostelium has demonstrated an important
role for the WASH complex in v-ATPase trafficking and, consequently, the regulation of vesicular pH (Carnell et al., 2011). During
maturation, autophagosomes are also acidified, due to delivery of
the v-ATPase from the lysosomal compartment. The rate of acidification and maturation is represented by the loss of GFP-Atg8 fluorescence after phagophore completion, allowing quantification (see
Movies S1 and S2). When we compared the time taken for the fluorescent signal to be removed in Ax2 and WASH mutants (n > 50,
across three independent experiments), no difference was observed,
which indicated there were no defects in acidification (Figure 1B).
In the above assays, autophagy was induced by mechanical
compression, via pathways that are poorly understood (King, 2012).
We therefore also tested the role of WASH under amino acid starvation. When WASH-null cells were placed in the defined SIH medium
lacking arginine and lysine (SIH-Arg/Lys, which induces autophagy,
but not the development of Dictyostelium; King et al., 2011) we
found no differences in either the rate of induction or steady-state
levels of autophagosomes after 24 h (Figure 1, C and D). We therefore conclude that WASH is not required for the induction, formation, or maturation of autophagosomes.
WASH is required for cytoplasmic degradation and surviving
starvation
Although not required for autophagosome formation, we hypothesized that WASH may be necessary for subsequent processing of
autophagosomes. The canonical role for autophagy is during starvation, when the degradation and recycling of cytoplasmic components is essential for cells to maintain nutrient levels and survive. To
test whether the autophagic pathway was fully functional in WASHnull cells, we determined their ability to survive Arg/Lys starvation.
While Ax2 cells were able to survive for almost 7 d before their viability decreased, WASH mutants died significantly faster, losing
almost 50% viability after only 4 d (Figure 1E). Their survival was,
however, better than that of Atg1 mutants, which are completely
deficient in autophagy and started dying after just 24 h. WASH is
therefore required for effective survival of Arg/Lys starvation.
WASH exists in a complex with four other proteins. We recently
determined that one of the complex subunits, FAM21, is required to
recycle WASH from endosomes. Loss of FAM21 therefore leads to
persistent WASH activity, blocking the endocytic pathway at a later
stage, after v-ATPase removal (Park et al., 2013). FAM21 mutants
were also unable to survive starvation, to the same degree as WASH
mutants (Figure 1E), indicating that the defect in survival is due to a
trafficking event downstream of WASH and v-ATPase recycling.
The function of autophagy under starvation is to degrade the
cytosol, which results in reduction of both cell size and mass (Otto
et al., 2003). Total protein levels were decreased by 15% in Ax2
cells upon 24 h Arg/Lys starvation, but remained constant in
Atg1-null cells (Figure 1G). Consistent with their inability to survive starvation, both WASH and FAM21 mutants also had no reduction in total cellular protein and retained their size over several days (Figure 1, F and G). Therefore, while autophagosomes
form and acidify normally in WASH mutants, total protein is not
reduced, and the mutant cells are unable to survive starvation.
WASH and lysosomal recycling | 2715 150
A
B
WASH
Acidification time (s)
Ax2
50
40
30
20
10
0
2
1
0
F
0
10
40
WASH
-Arg/lys 72h
Full Medium
Ax2
20
30
Time (min)
Ax2
WASH
2
1
0
0h
FAM21
Ax2
WASH
FAM21
Atg1
125
Viability (%)
3
WASH
E
3
Puncta per cell
Puncta per cell
D
Ax2
WASH
4
100
75
Atg1
50
*
25
***
*
*
0
24h
0
2
4
6
8 10
Days starvation
G
120
% Total protein
C
Ax2
100
80
0h *
24h
**
using beads coated with self-quenching
fluorescently labeled DQ-BSA (bovine serum albumin), which becomes dequenched
upon proteolysis (Gopaldass et al., 2012).
With this assay, the rate of proteolysis was
reduced by ∼50% in both WASH and FAM21
mutants compared with Ax2, and rescued
by reexpression of GFP-WASH in WASHnull cells (Figure 2A). WASH is therefore required for both phagocytic and autophagic
digestion, indicating a general role for
WASH in vesicular degradation. Importantly,
this cannot be due to a defect in acidification, as this is unaffected in both WASH and
FAM21 mutants (Figure 4E; Carnell et al.,
2011; Park et al., 2013).
Efficient phagocytic digestion is essential for Dictyostelium to survive on bacteria
as a food source. We therefore tested the
ability of WASH-null Dictyostelium to grow
on several bacterial strains. While WASHand FAM21-null cells grew normally on our
laboratory strains of Klebsiella aerogenes
and Bacillus subtilis, both mutants had dramatically reduced growth on the majority of
the other strains tested (Figure 2, B and C),
demonstrating a physiological requirement
for WASH in growth on bacteria.
Lysosomal components are reduced
in the phagocytic compartment
Phagosome maturation is a highly regulated and organized process during which
Ax2 WASH FAM21 Atg1
lysosomal components are sequentially deFIGURE 1: Autophagy in WASH mutants. (A) High-resolution imaging of autophagosome
livered and retrieved (Clarke et al., 2002;
formation in GFP-atg8–expressing cells compressed under agar. Stills are taken from Movies S1
Gotthardt et al., 2002, 2006a). To establish
and S2. (B) Quantification of the time taken for the GFP signal of individual autophagosomes in
the mechanism underlying the defects in
cells treated as in (A) to fade after autophagosome formation was complete. (n > 50
autophagosomes in total for each strain). (C) Autophagy was induced by placing cells expressing degradation, we used quantitative comparative proteomics to identify differentially
GFP-atg8 in SIH-Arg/Lys. The number of autophagosomes was counted at each time point.
represented proteins in the isolated pha(D) The steady-state levels of autophagosomes after 24 h of starvation. (E) Survival of mutants
gosomes of Ax2, WASH, and FAM21 muunder Arg/Lys starvation. At each time point, viability was measured by the ability to reform
colonies on bacterial plates. (F) Morphology of cells under starvation. DIC images of cells after
tants (Dieckmann et al., 2012; Gotthardt
3 d in full medium or SIH-Arg/Lys. (G) Total protein of cells after 24 h of Arg/Lys starvation. All
et al., 2006a). To confirm this approach,
values plotted are the means ± SD of three independent experiments. *, p < 0.05; **, p < 0.01;
we first looked at the levels of v-ATPase. In
***, p < 0.005 (Student’s t-test). Scale bars: 5 μm.
Dictyostelium, WASH is required for
v-ATPase recycling, and WASH mutants retain v-ATPase on their endosomes (Carnell
This indicates a defect in autophagic degradation and nutrient
et al., 2011). Consistent with this, all nine subunits of the v-ATPase
recycling.
complex were elevated in the WASH-null phagocytic compartment
compared with wild-type (Figure 3B). In contrast, FAM21­ mutants
retain WASH activity (Park et al., 2013), and several of the v-ATPase
Phagocytic proteolysis is reduced in WASH mutants
subunits were identified as decreased in FAM21-null phagosomes
Autophagy is not the only pathway that breaks down vesicular
relative to Ax2. This validates the data set and confirms that we are
cargo. In soil, Dictyostelium is a professional phagocyte, relying on
able to identify differentially trafficked proteins with this method.
the capture and digestion of bacteria within phagosomes to provide
Although the loss of WASH or FAM21 blocks the endocytic pathnutrients. After formation, phagosomes and autophagosomes are
way at different points, both mutants have comparable defects in
processed in a similar manner, with the same requirement for acidiphagocytosis and autophagy. We therefore looked for differences in
fication and hydrolytic enzymes in order to release nutrients (Deretic,
the phagocytic pathway common to both mutants. Of 85 proteins
2008). We therefore tested whether WASH is also required for
that were decreased by at least 30% in both mutants, 11 (13%) were
phagocytic digestion.
lysosomal proteins (Figure 3A and Supplemental Table S1). Several
Like autophagosome formation, loss of either WASH or FAM21
additional lysosomal proteins were also decreased in either WASH or
did not affect the rate of phagocytosis (Supplemental Figure S1A).
FAM21 mutants, indicating a general loss of lysosomal enzymes from
We were therefore able to directly measure phagocytic proteolysis
2716 | J. S. King et al.
Molecular Biology of the Cell
151
A
A
Ratio A488/594
6
Transmembrane
Ax2
WASH
FAM21
WASH::
GFP-WASH
4
Endosomal
membrane
Cytoskeleton
Unknown
Nucleus
ER
2
Mitochondria
Lysosome
0
B
Cells
0
25
50
75
Time (min)
K. aerogenes
Ax2 WASH FAM21
100
K. pneumoniae 52145
Ax2 WASH FAM21
10K
1K
Cytoplasm
Secreted
B
Gene
Protein
Gene ID
ctsD
Cathepsin D
DDB_G0279411
0.46***
ctsZ
Cathepsin Z;
DDB_G0283401
0.44***
V4-7
peptidase S8 and S53
domain-containing protein
Cysteine protease;
DDB_G0281823
0.40**
0.36*
DDB_G0291191
0.57***
0.45**
Beta-hexosaminidase
subunit A1
Beta-hexosaminidase
subunit A2
DDB_G0287033
0.60*
DDB_G0282539
0.45*
0.23**
GH-Family 25 lysozyme 2
DDB_G0274181
0.65***
0.47***
manA
Alpha-mannosidase
DDB_G0292206
0.72***
0.52***
m6pr
Mannose-6-phosphate
receptor
Lysosome membrane
protein 2-A
DDB_G0279059
0.62***
0.62**
DDB_G0267406
0.75**
0.40***
lmpB
Lysosome membrane
protein 2-B
DDB_G0287035
aprA
Autocrine Proliferation
Repressor
Sphingomyelinase A
DDB_G0281663
0.56***
0.39***
DDB_G0270834
0.64*
0.66**
Carboxylesterase, type B
DDB_G0279717
0.57***
0.38***
V-type ATPase subunit
DDB_G0287127
DDB_G0277401
DDB_G0284473
DDB_G0273071
DDB_G0275701
DDB_G0271882
DDB_G0277971
DDB_G0274553
DDB_G0291858
1.52***
1.56***
1.25***
1.68***
1.42***
1.61***
1.24*
1.41*
1.97***
0.89**
1.36***
100
nagA
nagB
P. aeruginosas
PT5
P.aeruginosas
PT531
S. aureus
E.coli DH5
E.coli B/r
B. subtilis
M. leutus
K. pneumoniae
52145
E. aerogenes
C
K. aerogenes
10
Ax2
WASH
FAM21
lmpA
sgmA
4
3
2
1
0
FIGURE 2: Phagocytic defects in WASH mutant cells (A) Proteolytic
activity is decreased in WASH mutants, as measured by feeding cells
DQ-BSA–labeled beads and measuring unquenching of the
fluorophore over time. (B) WASH- and FAM21-null cells are unable to
grow on certain bacteria. The indicated number of cells were seeded
on the bacterial lawn, and Dictyostelium growth is indicated by the
formation of dark plaques where the bacteria have been consumed.
(C) Summary of mutant growth on various bacteria. The color code
refers to the number of wells in (B) in which Dictyostelium were able
to grow, indicating the severity of the growth defect.
the phagocytic compartment (Figure 3B). These include several proteases, such as cathepsins D and Z, as well as a cysteine protease,
explaining the measured reduction in proteolytic activity. A form of
lysozyme (DDB_G0274181) was also reduced in both mutant
Volume 24 September 1, 2013
Ribosome
vatA
vatB
vatC
vatD-1
vatE
vatF
vatG
vatH
vatM
WASH/Ax2
Fam21/Ax2
0.37**
0.64***
0.54***
0.78*
0.64***
FIGURE 3: Proteomic analysis of mutant phagosomes. (A) The relative
proportion of proteins from different compartments is reduced by at
least 30% in both WASH and FAM21 mutant phagosomes. Eighty-five
proteins in all were identified by this criteria. (B) Identity and relative
abundance of lysosomal and secreted (below dashed line) proteins
identified in this screen. Lysosomal proteins only identified in one
mutant are also listed. The relative abundance of v-ATPase subunits is
included in the lower section. *, p < 0.05; **, p < 0.005; ***, p < 0.001
(Student’s t-test).
WASH and lysosomal recycling | 2717 152
A
WASH-
Full Medium
Ax2
B
70
-Arg/Lys
% positive cells
60
Full Medium
-Arg/Lys
***
50
40
30
20
10
0
Ax2
C
Lipidtox Red
FITC-dextran
D
Lipidtox Red
Lysosensor
Cascade blue Dextran
Merge
WASH
Merge
pH
phagosomes. As lysozymes are necessary to
break down bacterial cell walls, this reduction
would also contribute to the defective growth
of the mutants on bacteria.
In addition to known endo/lysosomal
proteins, our proteomic analysis also identified differences in a number of mitochondrial and ribosomal components that would
not normally be expected to be present in
phagosomes. As the data show relative
changes, these components are unlikely to
be contaminants, and their absolute abundance is not known. Instead, we speculate
that these proteins indicate the convergence of the autophagic and phagocytic
pathways and represent either changes in
autophagosome trafficking or global transcriptional changes.
Interestingly, the reduction in hydrolytic
enzymes was accompanied by a decreased
mannose-6-phosphate receptor (M6PR) in
both mutants. It should be noted, however,
that this M6PR is most similar to the cationdependent (CD)-M6PR family. Unlike the
cation-independent (CI)-M6PRs, CD-M6PRs
do not interact with the retromer (Arighi
et al., 2004; Seaman, 2004) and are therefore unlikely to be directly mediated by
WASH, as recently described (Gomez and
Billadeau, 2009).
WASH is also required for autophagic
lipid catabolism
E
7
pH
6
5
4
3
2
Ax3
WASHLipidtox -
WASHLipidtox +
FIGURE 4: Phospholipid accumulation in WASH-null cells. (A) Cells grown for 24 h in either full
SIH or SIH-Arg/Lys in the presence of LipidTOX Red. Arrows indicate large phospholipid
accumulations. Images are confocal maximum intensity projections. Scale bar: 10 μm. The
proportion of cells containing these large accumulations (>1.5-μm diameter) is quantified in (B).
More than 50 cells were scored for each sample, and the values plotted are the mean ± SD of
three independent experiments. ***, p < 0.005 (Student’s t-test). (C) Colocalization of LipidTOX
Red accumulations with both FITC (pH-sensitive) and Cascade Blue (pH-insensitive) dextran.
WASH-null cells were grown overnight in SIH-Arg/Lys supplemented with LipidTOX Red and
both dextrans. Arrows indicate subdomains of brighter FITC-dextran fluorescence. Image
shown is a single confocal plane. Scale bar: 5 μm. (D) Endocytic pH of cells starved overnight in
medium containing LysoSensor yellow/blue dextran and LipidTOX Red. The LysoSensor
emission at 450 nm and 510 nm is colored blue and yellow, respectively, in the LysoSensor panel
and was used to calculate the pH in the false-colored final image. The distribution of pH in
individual vesicles is shown in (E). All the LipidTOX Red–negative vesicles in 10 cells of each
strain were measured and are compared with the pH within the LipidTOX Red accumulations in
>20 WASH-null cells.
2718 | J. S. King et al.
In addition to proteolytic enzymes, several
enzymes involved in lipid catabolism were
also reduced in WASH mutant phagosomes.
These include both subunits of βhexosaminidase, as well as sphingomyeli­
nase A. In humans, loss of these enzymes
causes the accumulation of large quantities of lipid within lysosomes, leading to
Tay-Sachs and Sandhoff syndromes (βhexoaminidase deficiency) and Niemann-Pick
disease types A and B (sphingomyelinase A
deficiency). Inhibition of sphin­gomyelinase
has also been proposed to be a major cause
of drug-induced phospholipidosis (Yoshida
et al., 1985). Therefore, to test whether lipid
catabolism was disrupted in WASH-null cells,
we used a fluorescently labeled phospholipid
(LipidTOX Red) to label phospholipid accumulations in live cells.
In full medium, both Ax2 and WASH mutant cells were indistinguishable (Figure 4A).
However, after 16 h of Arg/Lys starvation,
WASH-null cells contained large accumulations of phospholipid, with 53% of cells containing a structure >1.5 μm in diameter
(Figure 4, A and B). To determine whether
these structures were endocytic in origin, we
also incubated the cells with fluorescently
labeled dextrans. These colocalized with the
Molecular Biology of the Cell
153
-
Ax2
WASH
Atg1
-
-
-
WASH /Atg1
Full Medium
A
1µm
-Arg/Lys
*
C
*
*
D
MLB's per cell
B
*
500nm
Full medium
***
- Arg/lys
***
4
3
2
1
0
Ax2 WASH Atg1 WASH/
Atg1
FIGURE 5: Ultrastructural analysis of starved WASH mutants. (A) Cells were placed in either full
or Arg/Lys-deficient medium for 36 h before processing for TEM. Micrographs show the
formation of large MLBs in starved WASH mutant cells, as indicated by asterisks. (B) and (C)
Enlarged images of MLBs, (B) is the boxed region in (A). (D) Quantification of MLB frequency.
At cells were grown as indicated, and the number of MLBs was scored per section, per cell. At
least 20 cells were scored for each sample, and values are the mean ± SD of three experiments.
***, p < 0.005 (Student’s t-test).
large phospholipid structures, indicating the lipid was accumulating
within the endocytic system (Figure 4C).
In WASH-null cells, the largest phospholipid-rich structures colocalized with pH-sensitive fluorescein isothiocyanate (FITC)-dextran,
indicating they were neutralized. To confirm this, we directly
measured the pH of these vesicles using LysoSensor yellow/blue
dextran, which has both pH-sensitive and pH-insensitive emission
peaks (Diwu et al., 1999). This again accumulated in the phospholipid-rich structures and indicated they were pH 5, in contrast to
pH 3.1 found in normal endocytic vesicles (Figure 4, D and E). This
was surprising, because WASH is essential for v-ATPase recycling in
Dictyostelium, and neutral compartments are normally completely
absent in WASH-null cells (Carnell et al., 2011). Elevated lysosomal
pH has, however, been reported in macrophages forced to accumulate cholesterol and in several lipid storage disorders (Bach et al.,
1999; Holopainen et al., 2001; Cox et al., 2007). It is therefore likely
that the excessive accumulation of lipids somehow disrupts acidification. These compartments also frequently contained a subdomain
of brighter FITC-fluorescence (Figure 4C, arrows), indicating they
must contain some intravesicular structures.
To look at these structures more closely, we analyzed mutant
cells by transmission electron microscopy (TEM). Confirming our
live-cell experiments, starved WASH-null cells contained many large
multilamellar structures that were almost completely absent in both
wild-type and full medium controls (Figure 5). Lysosomal multilamelVolume 24 September 1, 2013
lar bodies (MLBs) are the classical hallmarks
of many lysosomal storage disorders, including Tay-Sachs and Niemann-Pick disease, consistent with a reduction in βhexosaminidase and sphingomyelinase
activity in WASH mutant cells (Schmitz and
Muller, 1991). WASH is therefore required
for lysosomal lipid catabolism during starvation. Note also that, while the cytosol of Ax2
cells showed signs of digestion, becoming
patchy upon starvation, WASH mutants retained a dense cytoplasm (Figure 5A). This
is comparable with previous observations of
autophagy mutants, confirming that WASHnull cells do not lose mass upon starvation
(Otto et al., 2003).
MLBs are induced in WASH-null cells by
starvation. As autophagy is highly active under these conditions, we tested whether
autophagy directly contributed to this phenotype. Examination of WASH/Atg1 double
mutants by TEM indicated the accumulation
of MLBs was blocked (Figure 5, A and D).
The extensive phospholipid accumulation in
WASH mutant cells is therefore dependent
on autophagy. We interpret this to mean
WASH is required for the membranes and
contents of autophagosomes to be efficiently degraded, causing collapse of the
perturbed lysosomal system in WASH-null
cells.
Aberrant cathepsin D and βhexosaminidase trafficking
in WASH mutants
To establish why the degradative capacity
was reduced in WASH mutants, we examined the activity and localization of lysosomal enzymes. Despite a
50% reduction in the phagosomal compartment, when we measured the total protein level of cathepsin D (catD) by Western blot,
there was no difference between WASH-null and Ax2 cells, nor any
accumulation of partly processed enzyme (Figure 6, A and B). We
then measured the total enzymatic activity of lysosomal enzymes,
using fluorogenic substrates for cathepsin D/E and β-hexosaminidase.
Again, despite reduced protein levels in the phagocytic compartment, WASH-null cells contained 50% more cathepsin D/E and
more than threefold more β-hexosaminidase activity than wild-type
(Figure 6, C–E).
As Dictyostelium cells constitutively secrete lysosomal components, and WASH is essential for the secretion of insoluble material
such as dextran (Dimond et al., 1981; Carnell et al., 2011), we asked
whether lysosomal enzyme secretion was also blocked. In WASHnull cells, the secretion of both cathepsin D/E and β-hexosaminidase
activity was reduced by 50% (Figure 6, F–H). This explains the increase in total activities in the mutant but also demonstrates that,
unlike dextran, lysosomal enzymes are partly secreted by a WASHindependent pathway.
These experiments indicate that, despite the presence of elevated total levels of lysosomal hydrolases, their delivery to both
phagosomes and autophagosomes is blocked. WASH is therefore
needed for the correct trafficking, rather than the synthesis or processing, of the digestive enzymes.
WASH and lysosomal recycling | 2719 154
A
Ax2 WASH
Cathepsin D protein
kDa
B
100
70
50
40
35
1.0
0.5
0
Ax2 WASH
D
Cathepsin
600
Relative activity
400
1200
800
400
0
F
20
40
Time (min)
0
60
G
Cathepsin
Secreted activity (%)
100
75
50
25
0
1
2
3
Time (Hrs)
4
0
20
40
Time (min)
Hexosaminidase
Ax2
WASH
**
3
2
*
1
DISCUSSION
Cathepsin
Hex.
H
1
100
75
50
25
0
4
0
60
Relative activity
200
0
Secreted activity (%)
E
5
1600
WASH
0
Hexosaminidase
Ax2
µmol MU / 103 cells
Fluorescence (A.U.)
C
0.8
1
2
3
Time (Hrs)
4
*
0.6
***
0.4
0.2
0
0
Cathepsin
Hex.
FIGURE 6: Hydrolytic activities in WASH mutants. (A) Western blot of
catD (green) with 3-methylcrotonyl-CoA carboxylase α (MCCC1) as
loading control in red. Quantitation of three independent blots shown
in (B). (C–E) hydrolytic activity in Ax2 (circles) and WASH-null (crosses)
whole-cell lysates using fluorogenic substrates for (C) cathepsin D/E
and (D) β-hexosaminidase. Activity is indicated by the increase in
fluorescence over time. (E) The relative activity of WASH-null cells
(dark gray) compared with Ax2 (light gray). (F–H) Secreted activity of
(F) catD and (G) β-hexosaminidase. Cells were placed in fresh media
for the times indicated, and the activity in the media was measured as
above. (H) Relative rates of secretion are plotted in (H). All values are
the means ± SD of three independent experiments. *, p < 0.05; **,
p < 0.01; ***, p < 0.005 (Student’s t-test).
WASH is required for lysosomal enzyme recycling
To determine where lysosomal trafficking was disrupted, we examined the localization of catD in mutant cells by immunofluorescence.
As previously reported, in Ax2 cells fixed with ultracold methanol,
the catD antibody labeled spots on several large ring structures
(endocytic vesicles) and numerous smaller puncta (Figure 7A; Neuhaus et al., 2002). In both WASH- and FAM21-null cells however,
catD frequently accumulated in a single large structure, with a significant reduction in other small puncta (Figure 7, A and C). This was
confirmed using an alternative probe, fluorescently labeled pepstatin A, which specifically labels active catD and gave identical results
(Figure 7B; Chen et al., 2000).
These observations indicate that catD is retained within a specific cellular compartment in WASH mutants. These concentrated
catD structures, however, neither colocalized with the Golgi marker
golvesin-GFP (Schneider et al., 2000) nor with the endoplasmic reticulum marker protein disulfide isomerase (PDI; Figure 7, D and E).
Recently we showed that FAM21-GFP is able to localize to maturing
endosomes independently of the other WASH complex members
2720 | J. S. King et al.
(Park et al., 2013). When we expressed FAM21-GFP in WASH-null
cells, it clearly localized to a vesicular membrane surrounding the
catD structure, indicating that the catD accumulates where WASH
would normally be active.
Previous analysis of Dictyostelium phagosome maturation has
shown that the lysosomal hydrolases, including catD, are recovered
from the phagosome at a late stage, just prior to exocytosis of the
indigestible material (Gotthardt et al., 2002). Loss of WASH must
therefore block the pathway before this recycling can take place.
Lysosomal proteins therefore accumulate in a terminal compartment, unable to fuse with or be delivered to nascent phagosomes
and autophagosomes. Furthermore, the identical results obtained in
FAM21­-null cells (in which the pathway proceeds past v-ATPase recycling and neutralization before stalling) indicate that hydrolase recycling occurs at the same time as or after FAM21-mediated removal
of WASH and is independent of v-ATPase retrieval (Figure 8).
The WASH complex, and the actin it polymerizes on vesicles, has
been associated with an increasing number of trafficking roles. In
this paper, we demonstrate that disruption of WASH causes the accumulation of lysosomal hydrolases in a late endocytic compartment, where they get trapped and are thus unavailable for recycling
to nascent degradative compartments.
The reduced delivery of lysosomal hydrolases in WASH mutants
has several physiological implications. The primary defect in these
mutants is a decreased capacity to degrade lipids and proteins. This
means that both phagocytosis and autophagy are compromised,
and the cell is unable to use either bacteria or its own cytoplasm to
provide nutrients. Therefore, while WASH mutants grow normally
on our standard laboratory strain of K. aerogenes, they have much
more difficulty growing on other bacteria. It should be noted that
both the Dictyostelium and laboratory Klebsiella have been strongly
selected for optimum growth over decades of coculture in the
laboratory, and, although K. aerogenes, Klebsiella pneumoniae, and
Enterobacter aerogenes are now classified as the same bacteria
(Brisse et al., 2006), WASH mutants could grow only on our standard
laboratory strain. In the wild, amoebae need to consume a wide
range of bacteria, and so WASH function will be crucial for survival.
While there is no obvious pattern in the types of bacteria that WASH
mutants cannot use, it is likely that they have particularly high levels
of fat or lipid or are simply more difficult to digest. In addition to
enzymes required for protein and lipid catabolism, we also identified lysozyme as reduced in WASH mutant phagosomes. As
lysozyme is required to break down the bacterial cell wall, this reduction will also disrupt the digestive process and explain this
phenotype.
Consistent with a general lysosomal defect, WASH is also critical
for autophagic degradation. While there are no overt defects in autophagosome formation or acidification, WASH-null cells are unable
to digest their cytoplasm and survive starvation. In addition to
known endocytic proteins, our proteomics assay also identified a
number of components, such as mitochondrial and ribosomal proteins, that would not normally be expected to be present in phagosomes. As these data are ratiometric, it is unlikely that these are
contaminants, and they instead may be the products of autophagy,
indicative of altered autophagic trafficking and fusion with the endo/
phagocytic pathway.
Interestingly, we also show that induction of autophagy leads to
the accumulation of large quantities of phospholipid within lysosomal multilamellar bodies. This is characteristic of many lysosomal
storage disorders and implies that autophagy is a major source of
Molecular Biology of the Cell
155
(Carnell et al., 2011), so we were surprised to
find the largest accumulations of lipid within
neutral vesicles (Figure 4C). However, lyso60
somal pH is elevated in several lipid storage
***
diseases (Bach et al., 1999; Holopainen
et al., 2001), and the autophagy-mediated
40
accumulation of lysosomal cholesterol in
***
Niemann-Pick type C disease also inhibits
proteolytic activity (Elrick et al., 2012). It is
20
therefore likely that the excessive lipid accumulation in WASH-null lysosomes disrupts lysosomal function in multiple ways,
0
exacerbating the underlying defect in
Ax2 WASH FAM21
enzyme delivery.
In a number of respects, our observations contrast with the work of others on
Cat D
golvesin-GFP
Merge
mammalian epithelial cells, indicating a
novel mechanism by which WASH influences lysosomal trafficking. In HeLa cells,
WASH was shown to directly regulate retromer-mediated retrograde transport of the
CI-M6PR from endosomes to the Golgi.
WASH knockdown therefore leads to CIM6PR accumulation on endosomes (Gomez
and Billadeau, 2009). In contrast, our proteomic data show a putative CD-M6PR deCat D
α-PDI
Merge
creased on both WASH- and FAM21-null
phagosomes. Loss of FAM21 causes the
constitutive activation of WASH in both
Dictyostelium and mammalian cells (Park
et al., 2013), and the FAM21-free complex
remains able to mediate v-ATPase recycling
from late endosomes. In this respect, WASH
and FAM21 mutants have opposing phenotypes. Despite the contrasting defects in
v-ATPase trafficking, the putative CD-M6PR
Cat D
FAM21-GFP
Merge
is reduced in both mutant phagosomes.
This implies either indirect regulation by
WASH or that the hydrolases are recycled
with the WASH complex independently of
its actin polymerization activity.
Our data indicate that lysosomal hydrolases are retrieved after WASH-mediated
neutralization. This is in good agreement
with previous analysis of Dictyostelium phagosome maturation, which showed that difFIGURE 7: Localization of catD in WASH mutant cells. (A) Ultracold methanol fixed cells stained
ferent hydrolases are delivered to and rewith anti-catD antibody. (B) Paraformaldehyde-fixed cells stained with Bodipy-pepstatin A.
(C) Quantification of large aggregate frequency in cells stained as in (A); ***, p < 0.005
covered from phagosomes at distinct stages
(Student’s t-test). For further characterization of these structures, colocalization of catD with (D)
in their maturation (Gotthardt et al., 2002). It
golvesin-GFP (Golgi), (E) anti-PDI antibody staining, and (F) FAM21-GFP was examined in fixed,
is not known how the soluble enzymes are
WASH-null cells. Deconvolved wide-field images shown in all panels are all maximum intensity
selectively retrieved, or how different lysoprojections, except (E) and (F), which are single planes. All scale bars: 5 μm.
somal populations are maintained, but
WASH is clearly important to define distinct
lipid delivery to the lysosome. Autophagy is also required for the
phases of phagosome maturation. Binding of M6PRs to lysosomal
biogenesis of physiological MLBs, such as those secreted by lung
hydrolases is pH dependent, resulting in their release when they
alveolar cells as a surfactant, indicating a general role for autophagy
enter acidified lysosomes (Seaman, 2004). It is therefore plausible
in the transport of lipids (Hariri et al., 2000). Autophagy is therefore a
that a similar mechanism occurs during hydrolase recycling, requiring WASH-mediated neutralization before enzyme retrieval.
conserved source of lysosomal phospholipid and may therefore be a
While the general mechanisms are conserved, there are clearly
viable therapeutic target for the treatment of lysosomal storage
substantial differences in the trafficking pathways of Dictyostelium
disorders.
and mammalian epithelial and fibroblast cell lines. As a professional
Previous work has shown that WASH is absolutely required
phagocyte, Dictyostelium is more representative of other phagocytic
for v-ATPase recycling and vesicle neutralization in Dictyostelium
Bodipy-PepA
B
WASH
FAM21
C
% cells with large structures
Ax2
anti-catD
A
D
E
F
Volume 24 September 1, 2013
WASH and lysosomal recycling | 2721 156
A
Lysosomal hydrolases
V
Phagosome
V-ATPase
Post
lysosome
W W
V W VW
W
W
V
W
W
V
V Neutralisation W
W
VW VW
W
W
Actin coating and
FAM21-mediated
V-ATPase recycling
WASH recycing
Recycling
V
V
V V
Acidification and
degradation
V
V
V
V V
Autophagosome
W
Recycling
V
V
Recycling
V V
Exocytic
vesicle
While there is good evidence that WASH
specifically regulates the trafficking of several transmembrane proteins, such as the
v-ATPase, retromer, growth factor receptors,
and integrins (Derivery et al., 2009; Gomez
and Billadeau, 2009; Carnell et al., 2011;
Zech et al., 2011), such severe disruption of
the lysosomal system will have global physiological significance. In this study, we have
shown that, in Dictyostelium at least, WASH
has a general role in lysosomal maintenance
and the efficient digestion of phagocytic
and autophagic cargo.
MATERIALS AND METHODS
Cell strains and culture
WASH complex
For all experiments, except the proteomics
analysis, the Dictyostelium Ax2 strain was
the wild-type control and was the parent of
V
all mutants. Ax3 parent and isogenic knockV-ATPase
out strains were used for the phagosome
V
proteomics. All WASH and FAM21 knockout
V V
V
Hydrolase
Post
strains were described recently (Park et al.,
V
lysosome
accumulation
V
2013). Cells were routinely cultured in HL-5
V V
W W
V V
W
axenic medium, except in starvation experiV
Exocytic
W
V
ments, when they were grown in defined SIH
vesicle
V Neutralisation W
V
V V
V
W W
medium (Han et al., 2004) and then washed
V
Terminal
V V
three times in SIH-Arg/Lys. All media were
Compartment
supplied by Formedium (Hunstanton, UK).
W
Cells were transformed by electroporation
WASH complex
W
using standard methods.
The procedure to test growth on bacteFIGURE 8: (A) Schematic model of WASH and the endocytic pathway in Dictyostelium. After
ria has been extensively described previthe formation of phagosomes and autophagosomes, v-ATPase and lysosomal hydrolases are
ously (Froquet et al., 2009). Briefly, 10–104
delivered. These digest the cargo until WASH is recruited, sorting the v-ATPase into small
recycling vesicles and neutralizing the compartment. Subsequently the FAM21-mediated
Dictyostelium cells are deposited on a bacrecycling of WASH removes the actin coat, prior to exocytosis. At the same late stage,
terial lawn (50 μl of an overnight culture) and
lysosomal hydrolases are also recovered for delivery to new (auto)phagosomes. (B) In the
allowed to grow until the colonies become
absence of WASH, v-ATPase recycling is blocked, and lysosomes remain acidic. The pathway
visible. The bacterial strains used for
can proceed no further, which traps the hydrolases in this terminal compartment and blocks
the growth tests were kindly provided by
their recycling and delivery to nascent autophagosomes and phagosomes.
Pierre Cosson (Centre Médical Universitaire,
Geneva) and were K. pneumoniae laboratory strain and 52145 isogenic mutant (Benghezal et al., 2006), the
cells, such as leukocytes, macrophages, and dendritic cells, which
isogenic Pseudomonas aeruginosa strains PT5 and PT531 (rhlR-lasR
have similar requirements for the delivery of lysosomal enzymes to
avirulent mutant; Cosson et al., 2002), Escherichia coli DH5α (Life
phagosomes (Neuhaus and Soldati, 1999). A role of WASH in preparTechnologies, Paisley, UK), E. coli B/r (Gerisch, 1959), nonsporulating phagosomes for exocytosis has also been shown in macrophages
ing B. subtilis 36.1 (Ratner and Newell, 1978), and Micrococcus
expelling the pathogenic yeast Cryptococcus neoformans, indicatluteus (Wilczynska and Fisher, 1994). E. aerogenes was obtained
ing conserved functionality (Carnell et al., 2011). Macrophages seem
from the American Type Culture Collection (strain no. 51697;
particularly susceptible to drug-induced phospholipidosis (Schmitz
Manassas, VA).
and Grandl, 2009), so a similar WASH-dependent pathway may also
be important to protect cells against drug toxicity.
Dictyostelium viability and total protein assays
The secretion of lysosomal hydrolases is not unique to DictyosteCells were washed three times in SIH–Arg/Lys before being resuslium. In specialized mammalian cells, lysosome-related organelles
pending at 5 × 106 cells/ml in shaking flasks. Every 2 d, samples
also undergo exocytosis (Raposo et al., 2002), and calcium can stimwere removed and plated onto SM agar plates (Formedium,
ulate the secretion of lysosomes in fibroblasts (Andrews, 2000;
Hunstanton, UK) in conjunction with a lawn of K. aerogenes, and
Laulagnier et al., 2011). Whether the release of lysosomal enzymes
viability was assessed by the proportion of cells able to form new
is the primary function of such secretory events is doubtful, considcolonies, as previously described (Otto et al., 2003). For determiering that most will be inactive at the extracellular pH. It is therefore
nation of total protein upon starvation, cells were washed as above,
possible that mechanisms exist to recover the hydrolases prior to
and 1 × 106 cells were seeded in wells of a 24-well plate. At each
release that might require WASH in a similar manner.
time point, the contents of triplicate wells were resuspended, pelSeveral groups have also reported the disruption of endosomal
leted by centrifugation, and frozen. Pellets were then lysed in
and lysosomal networks when WASH is ablated in mammalian cells
100 μl 150 mM NaCl, 10 mM Tris, 1 mM ethylene glycol tetraacetic
(Derivery et al., 2009; Gomez et al., 2012; Piotrowski et al., 2012).
2722 | J. S. King et al.
Recycling
X
Recycling
Lysosomal hydrolases
Recycling
B
Molecular Biology of the Cell
157
acid (EGTA), 1 mM EDTA, 1% NP40 (pH 7.5), and total protein
measures were taken using Precision Red reagent (Cytoskeleton,
Denver, CO). Protein levels were then normalized to the unstarved
protein measurements for each strain.
Cloning and gene disruption
For generating an Atg1 knockout construct, the 5′ and 3′ fragments of the gene were amplified using the primers AAACAA­
ATGAACCCTTTGCC/aagcttTTGGATCCCATAAGGAAGTGAAGAGGCG and GGATCCaaaagcttTATTCATCACCAACCGAGGC/
TGAGTTCTCACTTCAAATGC. These were then combined by
PCR, and the floxed blasticidin cassette from pLPBLP (Faix et al.,
2004) was inserted as a BamHI/HindIII fragment to generate the
final Atg1 knockout construct pJSK471. The Bsr cassette from
WASH-null cells was removed using Cre recombinase (Faix et al.,
2004), and pJSK471 was used to make both a WASH/Atg1 double
and an Ax2 Atg1 single knockout. The FAM21-GFP construct used
was described by Park et al. (2013), and the golvesin-GFP plasmid
was generated by amplifying the full-length golvesin gene from
cDNA, adding 5′BamHI/3′XbaI sites, and subcloning into the GFPfusion vector pDM450 (Veltman et al., 2009).
Live-cell imaging and pH determination
Autophagosomes were observed using the GFP-Atg8 plasmid
pDM430 (King et al., 2011). High-resolution images were obtained
by overlaying cells with a layer of 1% agarose and compressing
them by capillary action, as previously described (Yumura et al.,
1984; King et al., 2011). Images were obtained on a Nikon (Tokyo,
Japan) Eclipse TE 2000-U microscope equipped with a 100×/1.45
numerical aperture (NA) Nikon total internal reflection fluorescence
oil-immersion objective, 473-nm diode laser illumination (Omicron,
Rodgau, Germany), and a Cascade 512F EMCCD camera (Photometrics, Tucson, AZ).
For determination of autophagosome induction upon starvation,
Z-series of images (0.25-μm step size) of uncompressed cells were
taken at each time point, using an Olympus IX81 inverted wide-field
microscope equipped with a 60×/1.42 NA Plan-ApoN objective.
The number of visible puncta in maximum intensity projections was
scored for >100 cells at each point. Differential interference contrast
(DIC) images were captured on an inverted microscope fitted with a
100×/1.40 NA oil-immersion objective (Nikon).
For phospholipid labeling, cells were incubated with HCS
LipidTOX Red (Molecular Probes) diluted 2000× in SIH or SIH–
Arg/Lys for 24 h. The endocytic pathway was labeled by also
including 0.2 mg/ml Cascade Blue and 0.4 mg/ml FITC-dextran
(Sigma-Aldrich, St. Louis, MO). Live-cell images were captured
on an Olympus FV1000 confocal microscope, using a
Plan-ApoN 60×/1.4 NA objective. Ax2 control cells were used to
calibrate the microscope laser intensities such that the pH-sensitive
FITC fluoresced only in a subset of vesicles, and identical settings
were used with all strains and conditions. For direct determination
of vesicular pH, cells were incubated overnight with
0.2 mg/ml LysoSensor yellow/blue dextran (Life Technologies;
Diwu et al., 1999). Confocal images were then collected, using
365-nm excitation and detecting the emission at both 450 and
510 nm. pH was determined by comparing the ratio of 450/510-nm
emission to a standard curve generated by imaging LysoSensor in
SIH medium at different pHs (Figure S1B). The false-colored vesicular pH images were then generated using ImageJ (http://
rsbweb.nih.gov/ij) by thresholding the 450-nm channel to generate a mask and converting the 450/510-nm emission ratios to pH
values. The emission ratio of individual vesicles was determined by
Volume 24 September 1, 2013
manually outlining each vesicle and measuring the mean intensity
of each channel.
Immunohistochemistry
Two methods of fixation were used. All antibody staining was done
using cells fixed in ultracold methanol, as described by Hagedorn
et al. (2006). Briefly, cells were seeded onto thin (no. 0) acid-washed
coverslips and left to adhere. They were then plunged into a beaker
of −80°C methanol on dry ice and left to fix for 30 min. After several
washes in phosphate-buffered saline (PBS), coverslips were blocked
with 3% bovine serum albumin (BSA) and stained. The rabbit
polyclonal anti-catD antibody was a kind gift from Agnes Journet
(Journet et al., 1999), and PDI was stained using a cocktail of five
monoclonal antibodies. For Bodipy-pepstatin A staining, cells were
fixed with 4% paraformaldehyde, 0.1% Brij35 in citrate buffer
(50 mM sodium citrate, pH 4.1) for 20 min before being washed and
then stained with 1 μM Bodipy-pepstatin A (Life Technologies; Chen
et al., 2000). Coverslips were kept in citrate buffer at all stages to
maintain cathepsin in the active conformation. All coverslips were
mounted in ProLong Gold with 4′,6-diamidino-2-phenylindole (Life
Technologies). Images were captured on an Olympus IX81 widefield fluorescence microscope with a 100×/1.4 NA objective and an
additional 1.6× optovar magnification. For deconvolution, a Z-stack
of images (0.25-μm step size) was captured with a CoolSNAPHQ
camera (Photometrics). Images were then deconvolved by Volocity
software (Perkin Elmer-Cetus, Waltham, MA) using calculated pointspread functions.
Phagocytosis and in vivo proteolysis assay
Bead uptake was measured as previously described (Sattler et al.,
2013). Fluorescent beads of 1-μm diameter (fluorescent YG-carboxylated beads; Polysciences, Warrington, PA) were added to cells at
a 200:1 dilution; at each time point, cells were washed free of unbound beads, and the number of ingested beads was determined
by flow cytometry. Proteolysis experiments were adapted from Yates
et al. (2005) and carried out as described previously (Gopaldass
et al., 2012). Briefly, cells were fed beads coupled to Alexa Fluor 594
succinimidyl ester (Life Technologies) and BSA labeled with DQ
Green at a self-quenching concentration (Molecular Probes). On
BSA proteolysis, DQ Green fluorescence increases due to dequenching of the fluorophore. Results were normalized to bead uptake by plotting the ratio of DQ Green to Alexa Fluor 594 fluorescence as a function of time.
TEM
For investigating subcellular structures using TEM, adherent Dictyostelium cells were fixed in HL5c medium with 2% glutaraldehyde and
1% paraformaldehyde for 1 h. Subsequently the cells were scraped
off the dish, washed with buffer, and incubated for 1 h with 0.5%
OsO4. The samples were washed several times; this was followed by
an incubation step with 1% tannin for 20 min at room temperature.
The cells were washed again, dehydrated, embedded in Epon resin,
and processed for conventional electron microscopy. Grids were examined with a Tecnai Spirit transmission electron microscope (FEI,
Eindhoven, Netherlands). Images were quantified by counting the
number of MLBs in >20 cells for each strain/condition.
Cathepsin and hexosaminidase assays
For enzymatic assays, cells growing in HL5 were resuspended,
washed twice in KK2 buffer (0.1 M potassium phosphate, pH 6.1),
and lysed in 50 mM sodium citrate buffer (pH 4.2) with 1% Triton
X100 at 5 × 104 cells/ml. For cathepsin activity, lysates were
WASH and lysosomal recycling | 2723 158
incubated at 37°C with 50 μM cathepsin fluorogenic substrate
(BML-P145-001; Enzo Life Sciences, Exeter, UK) in 50 mM citrate
buffer (pH 4.2), and samples were removed and quenched by being
diluted at 1:10 in 10% trichloroacetic acid. Fluorescence at 330-nm
excitation/410-nm emission was then detected in a fluorometer. For
β-hexoaminidase activity, 2 mM 4-methylumbelliferyl-β-d-Nacetylglucosamine (Calbiochem, San Diego, CA) was used as substrate, and reactions were quenched by dilution at 1:10 in 0.2 M
glycine, 0.2 M Na2CO3. Fluorescence was measured at 365-nm
excitation/445-nm emission. For calculation of β-hexoaminidase
activity, 4-methylumbelliferone was used to generate a standard
curve.
For measuring secreted enzyme activity, cells were washed three
times in fresh media and seeded at 2 × 106 cells per well of a 24-well
plate in 500 μl of fresh medium. At each time point, the medium was
carefully removed and spun for 30 s at 3000 rpm in a benchtop microfuge to pellet any cells, and 400 μl of the supernatant was taken.
Activity was determined by incubating samples of the media with
the appropriate substrates as above for 1 h at 37°C.
Mass spectrometry and analysis
Phagosomes were isolated by density centrifugation after cells were
fed latex beads, as described previously, using a pulse–chase of
15 min/2 h 45 min (Gotthardt et al., 2006b). The reduction, alkylation, digestion, and tandem mass tag (TMT) labeling was mainly
performed as described by Dayon et al. (2008). Briefly, phagosome
pellets were resuspended at 100 μg proteins per 33 μl of triethylammonium hydrogen carbonate buffer (TEAB: 0.1 M, pH 8.5, 6 M
urea). After addition of 50 mM Tris-(2-carboxyethyl) phosphine
hydrochloride (TCEP), samples were reduced for 1 h at 37°C. Alkylation was then performed at room temperature in the dark for 30 min
after addition of 1 μl of 400 mM iodoacetamide. Then 67 μl of TEAB
and 2 μg trypsin were added, and digestion was performed overnight at 37°C. Each sample was then labeled with a TMT reagent
according to the manufacturer’s instructions (Proteome Sciences,
Frankfurt, Germany) and evaporated under a speed-vacuum.
Off-gel electrophoresis was performed according to the manufacturer’s instructions (Agilent, Santa Clara, CA). After being desalted, pooled samples were reconstituted in OFFGEL solution.
Isoelectric focusing was performed using a 12-well frame on an
Immobiline DryStrip (pH 3–10, 13 cm) and run at 8000 V, 50 μA,
200 mW, until 20 kVh was reached. The 12 fractions were then
recovered and desalted using C18 MicroSpin columns.
ESI LTQ-OT (electrospray ionization linear trap quadropole orbitrap) mass spectrometry (MS) was performed on a LTQ Orbitrap Velos (Thermo Electron, Waltham, MA) equipped with a NanoAcquity
system (Waters). Peptides were trapped on a 5-μm, 200-Å Magic
C18 AQ (Michrom) 0.1-mm × 20-mm precolumn and separated on
a 5-μm, 100-Å Magic C18 AQ (Michrom, Auburn, CA) 0.75-mm ×
150-mm column with a gravity-pulled emitter. Analytical separation
was run for 65 min using a gradient of H2O/FA 99.9%/0.1% (solvent
A) and CH3CN/FA 99.9%/0.1% (solvent B). The gradient was run as
follows: 95% A and 5% B at 0–1 min, then to 65% A and 35% B at
55 min, and 20% A and 80% B at 65 min at a flow rate of 220 nl/min.
For MS survey scans, the OT resolution was set to 60,000 and the
ion population was set to 5 × 105, with an m/z window of 400–2000.
A maximum of three precursors were selected for both collision-induced dissociation (CID) in the LTQ and high-energy C-trap dissociation (HCD) with analysis in the OT. For tandem MS (MS/MS) in the
LTQ, the ion population was set to 7000 (isolation width of 2 m/z),
while for MS/MS detection in the OT, it was set to 2 × 105 (isolation
width of 2.5 m/z), with resolution of 7500, first mass at m/z = 100,
2724 | J. S. King et al.
and maximum injection time of 750 ms. The normalized collision
energies were set to 35% for CID and 60% for HCD.
Protein identification was performed with the EasyProt platform
(Gluck et al., 2012). After peak list generation, the CID and HCD
spectra were merged for simultaneous identification and quantification (Dayon et al., 2010). The parent ion tolerance was set to 10 ppm.
TMT-sixplex amino terminus and TMT-sixplex lysine (229.1629 Da),
carbamidomethylation of cysteines were set as fixed modifications.
All data sets were searched once in the forward and once in the reverse Dictyostelium Uniprot_sprot database. For identification, only
proteins matching two different peptide sequences were kept.
Isobaric quantification was performed using the IsoQuant module of EasyProt. TMT sixplex was selected as the reporter, and
a mass tolerance of 0.05 m/z was used to calculate WASH/Ax2
and FAM21/Ax2 protein ratios. Statistics were performed using
EasyProt’s Mascat and Libra statistical methods to generate a list of
proteins that significantly differed between samples.
Western blotting
Cells growing in HL5 were resuspended, washed in KK2 buffer, and
lysed in 150 mM NaCl, 10 mM Tris, 1 mM EGTA, 1 mM EDTA, 1%
NP40 (pH 7.5). Samples were then normalized to total protein content and boiled in SDS buffer (Life Technologies) before being
loaded on a 10% Bis-Tris acrylamide Nu-PAGE gel (Life Technologies). Gels were then transferred onto nitrocellulose membrane before being probed with anti-catD antibody diluted 1:5000 in PBS
(Journet et al., 1999) and a DyLight 800 anti-rabbit fluorescent secondary antibody (Pierce, Rockford, IL). As a loading control, we used
Alexa Fluor 680–labeled streptavidin (Life Technologies), which, in
Dictyostelium, recognizes a single protein at ∼77 kDa corresponding to the mitochondrial methylcrotonyl-CoA carboxylase (Davidson
et al., 2013). Blots were analyzed on a fluorescent gel imager, and
images were quantified using ImageJ.
ACKNOWLEDGMENTS
We thank Agnes Journet for kindly providing us with catD antibody;
Pierre Cosson for sharing the bacterial strains; David Strachan for
help with image processing; and Alex Scherl, Carla Pasquarello,
Patrizia Arboit, and Alexandre Hainard from the Proteomic Core
Facility (CMU, Geneva) for their proteomics expertise and help. We
are also very grateful to Pete Thomason, Tobias Zech, and Laura
Machesky for many helpful discussions and critiques of the manuscript and to Pete Watson for informed suggestions. This work was
supported by Cancer Research–UK core funding to R.H.I. and a grant
from the Swiss National Science Foundation to the T.S. laboratory.
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Supplemental Materials
Molecular Biology of the Cell
King et al.
162
Supplementary Figure 1. (A) Normal phagocytosis of beads in WASH complex
mutants. (B) Calibration curve of lysosensor blue/green fluorescence at different
pH’s. The ratio of emission at 450nm/510nm at each pH is plotted. A best fit curve
was then fitted using the least squares method, and the equation, and R2 value are
indicated on the graph.
Supplementary table 1. Full list of proteins reduced by at least 30% in both WASH
and FAM21 mutants. Proteins are grouped by subcellular compartment. *P<0.05,
**P<0.005, ***P<0.001 (T-test).
1
163
164
165
5.4-Phagocytosis of mycobacteria is decreased in LmpB ko cells
Figure 1 from Sattler et al. (Sattler et al., in preparation)
Uptake of different fluorescent particles by wild type, lmpA kd, lmpB ko or lmpC ko cells was
measured by flow cytometry. Relative fluorescence units are normalised to the fluorescence of wild
type cells after 60 minutes of uptake. Symbols and error bars indicate the mean and SEM of three to
four experiments. Phagocytosis of the following particles was monitored: A. 1 µm YG-beads. B. 4.5
µm YG-beads. C. Alexa-488 labeled K. pneumoniae. D. Alexa-labeled B. subtilis. E. GFP-expressing
M. marinum. F. GFP-expressing M. marinum-L1D. G. GFP-expressing M. smegmatis.
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185 Acknowledgments
I would like to thank Professor Thierry Soldati for offering me the opportunity to do my PhD in his lab
and for the interesting and challenging project. I especially thank him for always being available.
I want to thank Professor Hubert Hilbi for accepting to be part of my thesis committee.
I also thank Professor Pierre Cosson for being a member of my jury and of my thesis exam jury, and
for his helpful comments and input as a member of my TAC meeting.
I thank Professor Jean-Charles Sanchez for being a member of my jury and for his participation to my
TAC meeting.
A big thank to my nice colleagues of the Soldati lab for the nice working atmosphere and for the nice
discussions during lunch and coffee breaks. I especially thank Regis who taught me everything when I
arrived in the lab and Monica who taught me everything about mycobacteria. A big thank you to Sonia
who had to share the office with me and had to deal with my stress. I know she particularly
appreciated my music. A very special thank you to Natascha who was always there to support me
during those years and who made my life much better in Geneva.
I also thank the people of the biochemistry department for the nice atmosphere, especially my
colleagues of the PhD course: Chrystelle, Charlotte, Aline… A big thank you to Christin, especially
for her support during the two months of writing.
Merci à Domitille et Romain pour les kilos de pates carbos ainsi qu’à Fanny et Vincent d’avoir été la
pour me changer les idées de ma thèse.
Je remercie mes parents, mes soeurs et Laurent pour les bons moments passés pendant leurs visites,
pour les heures passées au téléphone et tout simplement pour m’avoir soutenue pendant ces années de
thèse. Elo et Lulu, BFAV!
186