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Vol. 14, Issue 5, 1835-1851, May 2003
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* Institut de Biologie et Chimie des Protéines, UMR5086,
CNRS/Université Lyon I, IFR 128 BioSciences Lyon-Gerland, 7, Passage du
Vercors, 69367 Lyon cedex 07, France;
Laboratoire de Biochimie et Biophysique des Systémes
Intégrés, 38054 Grenoble Cedex 9, France;
241 Patterson Laboratories, Section of Molecular Cell and Developmental
Biology, The University of Texas at Austin, Austin, Texas 78712;
Laboratoire de Biologie Moléculaire et Cellulaire/UMR 5665 Ecole
Normale Supérieure de Lyon, 69364 Lyon Cedex 07, France; and
¶ Université de Genève, Centre Médical Universitaire,
Département de Morphologie, CH-1211 Genève 4, Switzerland
Submitted October 2, 2002;
Revised November 20, 2002;
Accepted December 27, 2002
Monitoring Editor: Randy Schekman
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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-adaptin AP-1 subunit was cloned and shown to belong to a
Golgi-localized 300-kDa protein complex. Time-lapse analysis of cells
expressing -adaptin tagged with the green-fluorescent protein
demonstrates the dynamics of AP-1–coated structures leaving the Golgi
apparatus and rarely moving toward the TGN. Targeted disruption of the AP-1
medium chain results in viable cells displaying a severe growth defect and a
delayed developmental cycle compared with parental cells. Lysosomal enzymes
are constitutively secreted as precursors, suggesting that protein transport
between the TGN and lysosomes is defective. Although endocytic protein markers
are correctly localized to endosomal compartments, morphological and
ultrastructural studies reveal the absence of large endosomal vacuoles and an
increased number of small vacuoles. In addition, the function of the
contractile vacuole complex (CV), an osmoregulatory organelle is impaired and
some CV components are not correctly targeted. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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;
Rothman and Wieland, 1996).
One of the best characterized coat protein is clathrin, which, in association
with adaptor complexes (APs), participates in multiple transport steps
(Hirst and Robinson, 1998;
Smith and Pearse, 1999).
Four different AP complexes have been identified (AP-1 to AP-4;
Boehm and Bonifacino, 2001).
All APs share a similar composition. They are comprised of two large subunits
(80–130 kDa), one medium size subunit (50 kDa), and one small subunit
(20 kDa). For instance, AP-1 contains 
1- and -adaptin large
chains, a µ1A or µ1B medium chain, and a 
1A or 1B small
chain (Scales et al.,
2000). Cargo sorting relies on -adaptin and µ chains.
These two chains recognize tyrosine- and leucine-based sorting signals found
in the cytoplasmic domains of cargo proteins
(Ohno et al., 1995;
Rapoport et al.,
1998), thereby concentrating them into budding vesicles.
Each individual AP complex functions at a distinct intracellular site
(Kirchhausen, 1999). Hence,
AP-1 is mainly recruited to membranes of the trans-Golgi network
(TGN) and participates in transport from the TGN to endocytic compartments.
Recently, the function of AP-1 in membrane traffic has been carefully
reassessed. Gene targeted disruption of µ1A or -adaptin in mice and
of µ1 in Saccharomyces cerevisiae suggested that AP-1 is required
for retrograde transport from endosomes to the TGN of mannose 6-phosphate
receptors (MPR) in mammalian cells as well as chitin synthase III and syntaxin
Tlg1p in yeast cells (Zizioli et
al., 1999; Meyer et
al., 2000; Valdivia
et al., 2002). However, studies on the dynamics of AP-1
followed in living cells expressing yellow-fluorescent-protein–tagged
µ1 revealed that AP-1 is mainly associated with transport structures moving
from the TGN toward the cell periphery, whereas AP-1–associated
retrograde transport appears to rarely occur
(Huang et al., 2001).
In addition, AP-1 has been implicated both in the transport of the transferrin
receptor from apical to basolateral membranes in epithelial cells
(Futter et al., 1998)
and the recycling of the low-density lipoprotein receptor and the transferrin
receptor to the basolateral membrane (Gan
et al., 2002). AP-1 was also reported to mediate
transport from the TGN to the basolateral membrane of many membrane proteins
(Folsch et al., 1999,
2001). Thus AP-1 could
function in more than one location; however, how a single vesicular coat could
play multiple roles is not understood yet.
The amoeba Dictyostelium discoideum is a genetically tractable
eukaryotic cell that is commonly used as cellular model to study membrane
trafficking in the endocytic pathways. Being a professional phagocyte,
Dictyostelium binds to and internalizes large size particles (>1
µm) by phagocytosis (Maniak,
1999; Neuhaus and Soldati,
1999; Cardelli,
2001; Maniak,
2001; Rupper and Cardelli,
2001). Laboratory strains can also grow in axenic liquid culture
medium. Fluid-phase nutrients are then mainly internalized by
macropinocytosis, concentrated in endosomes, and degraded in lysosomes.
Undigested material can eventually be returned to the cell surface via
postlysosomal vacuoles (Neuhaus and
Soldati, 1999). The biosynthesis of lysosomal enzymes in
Dictyostelium is similar to that in mammalian cells
(Cardelli, 1993). Newly
synthesized lysosomal hydrolases are first synthesized as membrane-bound,
N-glycosylated precursor proteins in the ER and then transported to the Golgi.
However, in contrast to mammalian cells where lysosomal enzymes are targeted
to lysosomes through the recognition of mannose 6-phosphate (M6P) sugars by
MPRs, the sorting machinery recognizing M6P sugars is poorly characterized in
Dictyostelium, and in particular MPRs have not been identified yet
(Cardelli, 1993). The fact that
the clathrin heavy chain is required for proper sorting of lysosomal enzymes
(Ruscetti et al.,
1994) suggests that clathrin-coated vesicles play a crucial role
in this process in Dictyostelium.
As a first step toward a better understanding of AP-1–dependent transport mechanisms, we decided to investigate AP-1 function in the model organism Dictyostelium. Here we report for the first time the molecular and functional characterization of the AP-1 complex in this organism. Together our data provide new evidence for the sorting function of AP-1 in the endocytic pathway and uncover the role of AP-1 in CV biogenesis.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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) was cultured at 22°C in HL5 medium. To measure growth on
bacterial lawns, cells were mixed with 400 µl of overnight liquid culture
of Klebsiella aerogenes and plated on SM nutrient agar plates
(Kay, 1987). For developmental
analysis, cells were plated on 0.45-µm membrane filters (Whatman,
Maidstone, UK) laid on Na/K phosphate buffer and incubated at 24°C in
humid chambers (Sussman,
1987). To assess constitutive secretion in HL5 culture medium,
cells were resuspended in HL5 at 5 x 106 cells/ml and
incubated at 22°C. After 10 h, culture medium was TCA precipitated and
pellets were analyzed by Western blotting.
Electron Microscopy
For conventional electron microscopy the cells were processed as described
(Orci et al., 1973).
Sections were photographed in a Philips CM-10 transmission electron microscope
(Philips, Eindhoven, The Netherlands) at calibrated magnifications.
Antibodies and Immunofluorescence Microscopy
Polyclonal antibodies to Dictyostelium µ1 and -adaptin
were raised in rabbits using KLH-coupled peptides
(320VPPDADTPKFRC331; Covalab, Lyon, France) and
GST- (592–896) recombinant protein, respectively. Rabbit
polyclonal antibodies to Dictyostelium cathepsin D
(Journet et al.,
1999) and clathrin heavy chain (CHC) were gifts from Dr. J. Garin
(CEA, Grenoble, France) and Dr. T. O'Halloran (University of Texas, Austin,
TX), respectively. Mouse monoclonal antibodies (mAb) against
-mannosidase (2H9; Mierendorf
et al., 1983), comitin (190-68-1;
Weiner et al., 1993),
and a Golgi unknown antigen (1/39; Graf
et al., 1999), were kind gifts from Dr. H. Freeze
(University of California, La Jolla, CA), Dr. A. Noegel (University of
Köln, Germany), and Dr. R. Gräf (Muenchen, Germany). Monoclonal
antibodies (mAbs) against coronin (176-2-5), vacuolin B (221-1-1), and the A
subunit of the vacuolar H+-ATPase (221-35-2) have been
characterized previously (Zhu and Clarke,
1992; Jenne et al.,
1998; Neuhaus et al.,
1998; de Hostos,
1999). The rat antiphagosome mAb M12A9 was reported to cross-react
with Dictyostelium -adaptin
(Morrissette et al.,
1999). The mAb H161, specific for a p80 cell surface marker, has
been described (Ravanel et al.,
2001). Rh50 was stained with a rabbit polyclonal antibody
(Benghezal et al.,
2001). To allow simultaneous labeling with an anti-p80 antibody
and another mouse mAb, the H161 antibody was directly coupled to Alexa Fluor
488 (Molecular Probes, Leiden, Netherlands) according to manufacturer's
instructions, and cells were processed for immunofluorescence as described
(Ravanel et al.,
2001). For indirect immunofluorescence analysis, unless indicated,
cells were grown on glass coverslips for 3 days, fixed with —20°C
methanol for 10 min, incubated with the indicated antibodies for 1 h, and then
stained with corresponding fluorescent secondary antibodies. Cells were
visualized with a Zeiss confocal microscope (LSM510) (LePecq, France). For
fluorescence microscopy in living cells, just prior observation cells were
compressed under a thin layer of agarose
(Yumura et al.,
1984). For excitation of all GFP constructs, the 488 band of an
argon-ion laser was used together with a 515–565 filter for emission.
For epifluorescence microscopy cells were observed using a Zeiss Axioplan 2
microscope.
Immunoprecipitation, Binding Assay, and Western Blotting
For immunoprecipitations, 107 cells were lysed in lysis buffer
(PBS, 10 mM EDTA, 1% Triton X-100, protease inhibitors) and cleared by
centrifugation for 15 min at 13,000 rpm in a microfuge. About 1 mg of protein
extract was incubated overnight at 4°C with mAb M12A9 and Gammabind
Sepharose beads (Amersham Pharmacia, Orsay, France). The beads were then
washed three times in lysis buffer and twice in 50 mM Tris-HCl, pH 7.5, and
bound proteins analyzed by immunoblots. SDS polyacrylamide electrophoresis and
immunoblotting were performed as previously described
(Cornillon et al.,
2000). Bands were visualized using an ECF kit (Amersham Pharmacia)
and a STORM Imager (Molecular Dynamics, Sunnyvale, CA). Binding assays between
GST- fusion proteins and cytosol were carried out as described
(Doray and Kornfeld,
2001).
Gel Filtration, Sucrose Gradient Fractionation
To prepare cytosol, Dictyostelium cells were washed twice in
breaking buffer (1 mM EDTA, 150 mM NaCl, 20 mM MES-Na buffer, pH 6.5, 10 mM
iodoacetamide, protease inhibitor cocktail), suspended at 4 x
108 cells/ml in the same buffer and then broken by six strokes in a
ball-bearing cell cracker (Balch and
Rothman, 1985). Unbroken cells were removed by low speed
centrifugation (1000 x g, 5 min, 4°C). The supernatant was
then centrifuged at 100,000 x g for 1 h in a SW50Ti rotor.
Cytosol was collected and then loaded onto a HiPrep 16/60 Sephacryl S-300 HR
gel filtration column (Amersham Pharmacia) equilibrated in breaking buffer.
One-milliliter fractions were collected, and protein contents were TCA
precipitated before SDS-PAGE and immunoblotting analysis. Membrane
preparations and sucrose gradient fractionation were carried out as reported
(Bogdanovic et al.,
2000).
Plasmids and Cell Transfection
Full-length (nucleotides 1–1287) and truncated (nucleotides
1–1233) apm1 cDNAs, encoding µ1 and µ1
Ct,
respectively, were amplified by PCR, sequenced (MWG-Biotech, Ebersberg,
Germany), and cloned into the Dictyostelium expression vector pDXA-3C
(Manstein et al.,
1995). Plasmid were transfected in Dictyostelium by
electroporation as described (Cornillon
et al., 2000). -Adaptin encoding cDNA was isolated
by PCR screening of a cDNA library kindly provided by Dr. Fuller (University
of California, San Diego, CA) with oligonucleotides derived from a partial
gene sequence (contig6354) encoding part of Dictyostelium
-adaptin. A full-length cDNA insert (GenBank accession number AY144597
[GenBank]
)
was sequenced on both strands and cloned into pDXA-3C. For GFP tagging of
-adaptin, a GFP insert was obtained by PCR using pTX-GFP as template
(Levi et al., 2000)
and inserted before the stop coding of cDNA. -GFP was included
in AP-1 complexes and partitioned into membranes and cytosol as native
-adaptin and, very few monomeric -GFP proteins were detected in
cytosol fractionated by gel filtration on a Sephacryl S300 column (our
unpublished results). GST- constructs were made by PCR and subcloned
into the vector pGEX-4T (Amersham Pharmacia).
Apm1 Knockout
DNA fragments comprising nucleotides 373–578 (5' fragment) and
579-1287 (3' fragment) of apm1 cDNA were amplified by PCR from
DH1-10 genomic DNA and cloned into pCRII-TOPO (Invitrogen, Groningen,
Netherlands). The apm1 knockout construct was made by inserting the
blasticidin resistance cassette (Sutoh,
1993) between the 5' and 3' apm1 fragments.
The final construct was linearized by SalI and electroporated into
DH1-10 cells as for complementation studies. Transformants were selected in
the presence of 10 µg/ml blasticidin, and individual colonies were tested
by Southern blot and immunoblotting. Four independent clones were identified
and displayed identical phenotypes. One clone was chosen for further
characterization.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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). A rabbit polyclonal antibody against µ1 was raised, and
this antiserum recognized a protein with an electrophoretic mobility of 50 kDa
(Figure 1A) as predicted from
its cDNA sequence. Next, the identification of the -adaptin subunit was
facilitated by the characterization of an antimurine phagosome mAb (M12A9)
that cross-reacts with Dictyostelium phagosomes
(Morrissette et al.,
1999). In Dictyostelium, this antibody recognized a
100-kDa protein (Figure 1A),
consistent with the molecular mass of APs large subunits. Molecular cloning of
the corresponding cDNA suggested that this protein could correspond to
Dictyostelium -adaptin because the deduced protein sequence
shares 44% of identity with the human 1-adaptin
(Figure 2).
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To determine whether µ1 and -adaptin associate with other
subunits to form a bona fide AP-1 complex, Dictyostelium
cytosol was fractionated by gel filtration on a Sephacryl S300 column. µ1
and -adaptin were detected in fractions 16–26, peaking at
fraction 20, which corresponds to an apparent size of 
300 kDa
(Figure 1A) as observed for
AP-1 complexes in different species
(Kirchhausen, 1999). The
assumption that both chains were subunits of the same adaptor complex was
further strengthened by the finding that the µ1 subunit, but not the
lysosomal enzyme Cathepsin D (CatD), can be coimmunoprecipitated in cell
lysates with the -adaptin chain recognized by the anti- M12A9
antibody (Figure 1B). Taken
together, these results demonstrate the existence of an AP-1 complex in
Dictyostelium.
Intracellular Localization of the AP-1 Complex in Dictyostelium
In mammalian cells, AP-1 is found not only in the cytosol but also in
association with Golgi membranes. To establish the intracellular localization
of µ1-containing AP complexes in Dictyostelium, membranes were
fractionated on linear (15–57%) sucrose gradients
(Figure 1C). As previously
reported (Bogdanovic et al.,
2000), lysosomal compartments were concentrated in high-density
fractions (fractions 2–5), whereas plasma membrane, contractile vacuole,
and Golgi membranes were contained in low-density fractions (fractions
8–11; unpublished data). Comitin (p24), an actin-binding protein
associated with the Golgi apparatus, was previously found in two distinct
fractions in sucrose gradients (Weiner
et al., 1993). As shown in
Figure 1C, µ1 cofractionated
with low-density comitin (peaking at fraction 9) in fractions containing Golgi
membranes. This result was further confirmed using an antibody (mAb 1/39)
against an unknown antigen specifically localized to the Golgi apparatus
(Graf et al., 1999).
Indeed distribution of µ1 on sucrose gradient was similar to that of the
protein detected by mAb 1/39 (Figure
1C). Together, these data indicate that the AP-1 complex is mainly
associated with Golgi-enriched fractions in Dictyostelium.
The AP-1 localization was next examined by confocal microscopy experiments.
Double immunostaining with polyclonal anti–-adaptin and
anticomitin antibodies revealed that -adaptin and comitin (a marker for
the Golgi) were mainly concentrated in the same Golgi area as well as in
vesicular structures throughout the cytoplasm
(Figure 3A). The localization
of AP-1 subunits to the Golgi was also confirmed by assessing the distribution
of GFP-tagged -adaptin (for the biochemical characterization of
-GFP see MATERIALS AND METHODS). When expressed in
Dictyostelium cells, -GFP localized to the Golgi area labeled
with clathrin (Figure 3A),
suggesting that in vivo, AP-1 subunits recruit clathrin coats. Some
clathrin-coated structures did not colocalize with -GFP and could
correspond to vesicles including other adaptor proteins. Interestingly,
-GFP and native -adaptin associated with Golgi membranes were
insensitive to Brefeldin A (BFA), a fungal metabolite that triggers
dissociation of vesicular coats (e.g., COP1 and AP-1) from membranes of
mammalian cells (Donaldson et
al., 1990; Robinson and
Kreis, 1992; Wong and Brodsky,
1992). This lack of sensitivity to BFA was already reported for
-COP in Dictyostelium cells and is likely due to the presence
of a BFA resistance gene (Mohrs et
al., 2000).
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Disruption of the actin network using DMSO leads to the reversible
fragmentation of the Golgi apparatus and the redistribution of Golgi markers
(e.g., -COP) in the cytoplasm of mammalian and Dictyostelium
cells (Weiner et al.,
1993; Mohrs et al.,
2000). To determine whether -GFP redistributes as
endogenous -adaptin during DMSO treatment in vivo,
-GFP–expressing cells were incubated in 5% DMSO for 0, 20, and 40
min, stained with a polyclonal anti–-adaptin antibody, and
analyzed by epifluorescence microscopy
(Figure 3B). After 20 min of
treatment, both -GFP and endogenous -adaptin showed a diffuse
cytosolic stain and also localized to many vesicular structures. However,
after 40 min, both proteins reassembled in a perinuclear region with identical
kinetics. These results demonstrate that -GFP and native
-adaptin are localized to the same organelle and behave identically
when cells are treated with an agent that affects Golgi integrity.
Dictyostelium AP-1 Recruits Clathrin
The colocalization of -GFP with clathrin suggested that AP-1
recruits clathrin coats in living cells. To assess clathrin binding of
Dictyostelium -adaptin, we constructed GST- fusion
proteins and assayed their ability to bind clathrin from
Dictyostelium cytosol. Analysis of the protein sequence of
Dictyostelium -adaptin revealed only one putative copy of the
clathrin box motif, at position 630 (Figure
2), identified in mammalian chain as responsible for
clathrin binding (ter Haar et
al., 2000; Doray and
Kornfeld, 2001). Both GST- 592–896 (hinge and
appendage, with the clathrin box) and GST- 663–896 (appendage,
without the clathrin box) displayed clathrin binding capacities
(Figure 3C). As the presence of
the hinge domain resulted in an enhanced recruitment of clathrin in pull-down
experiments, the hinge and appendage domains could present independent
clathrin binding domains as previously reported for mammalian chain
(Doray and Kornfeld,
2001).
Intracellular Dynamics of -GFP in Living Cells
To determine the dynamic localization of AP-1 in living cells, we observed
the intracellular dynamics of -GFP–expressing cells by time-lapse
confocal microscopy. The main event was the rapid movement of AP-1 structures
emerging from the Golgi area toward the cell periphery. These AP-1 structures
included vesicles but also tubules and/or queues of vesicles along a same
track (Figure 4A). Most AP-1
structures rapidly disappeared when reaching their destination (or were moving
out the confocal plane), but in some cases, AP-1 vesicles moved back to the
Golgi following the same track (Figure
4B). Together these observations are consistent with the role of
AP-1 in the transport of cargo proteins from the TGN to endosomes. In
addition, AP-1 structures moving toward the TGN were also observed but at a
very low frequency (our unpublished results), although these events were
difficult to reliably quantify because AP-1 structures moving along
microtubules hardly stayed in a same confocal plane and, consequently, AP-1
structures docking to Golgi membranes could be easily overlooked. This result
suggests that AP-1 could be also implicated in retrograde transport between
endosomes and the TGN in Dictyostelium cells as reported for
mammalian and yeast cells.
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The µ1 Chain Is Not Required for AP-1 Assembly and Golgi
Localization
To determine the role of AP-1 in Dictyostelium, we decided to
invalidate genes encoding AP-1 subunits. For unknown reasons, we failed to
inactivate the -adaptin gene, and therefore the apm1 gene,
encoding the µ1 AP-1 subunit, was disrupted by targeted integration of the
blasticidin selection marker (see MATERIALS AND METHODS). The µ1 protein
was not detected in mutant cells (Figure
5A), whereas stable transfection of native µ1
(apm1— + µ1), or µ1 truncated by deletion of
its 18 C-terminal residues (apm1— + µ1Ct),
restored µ1 expression to levels comparable to wild-type cells (DH1;
Figure 5A). The deleted domain
in µ1Ct corresponds to a region in murine µ2 required for the
interaction between µ chains and tyrosine-based signals
(Owen and Evans, 1998). This
truncated µ1 chain was unable to bind mammalian tyrosine signals in a yeast
two-hybrid assay (our unpublished results). Therefore the µ1Ct
construct was used hereafter to determine whether the observed defects in
mutant apm1— cells were due to the absence of a
functional µ1 subunit or to any aberrant functions of partial AP-1
complexes deprived of µ1 chains.
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To determine the structure of AP-1 complexes in mutant
apm1— cells, cytosol was fractionated by gel
filtration and fractions were analyzed by immunoblotting with the
anti–-adaptin antibody. In contrast to wild-type cells,
-adaptin from apm1— cytosol was detected in
fractions 20–26, peaking at fraction 22, which corresponds to an
apparent size of 250 kDa (Figure
5B). Therefore AP-1 subunits are able to assemble without µ1
chains.
A characteristic of AP complexes is to cycle between cytosolic and
membrane-bound pools. To evaluate whether AP-1 complexes without µ1 chains
partitioned into cytosolic and membrane fractions as native AP-1, we
determined the relative amount of -adaptin in cytosol (C) and total
membrane (M) fractions prepared by centrifugation. In wild-type cells, 87% of
total cellular -adaptin associated with membranes, whereas only 52% was
detected in membranes of apm1— mutant cell
(Figure 5C). We next assessed
the intracellular localization of AP-1 partial complexes in
apm1— cells. As observed in
Figure 5D, -adaptin was
detected in the Golgi area of apm1—,
apm1— + µ1Ct and wild-type cells. Taking
together these results demonstrate that µ1 is not essential for specific
recruitment of AP-1 to Golgi membranes in Dictyostelium cells but
participates in membrane binding efficiency.
apm1 Disruption Causes Defects in Cell Growth and Development
Mutant apm1— cells were viable but grew poorly on
bacterial lawns (Figure 6A) and
in liquid culture (Figure 6B).
Expression of native µ1, but not of µ1Ct, in
apm1— cells reverted the growth defect. In old
cultures, some giant cells with 3–5 nuclei per cell appeared, suggesting
cytokinesis defects in this mutant.
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On nutrient depletion, Dictyostelium cells aggregate and proceed
to a developmental cycle leading to the formation of fruiting bodies. Previous
studies on clathrin heavy-chain null mutants (chc—) revealed
the important role played by clathrin during Dictyostelium
development (Niswonger and O'Halloran,
1997). chc— cells are delayed in early
development and fail to produce any spores. To evaluate the role of AP-1 in
development, mutant cells were layered onto filters soaked with Na/K phosphate
buffer. In contrast to chc— cells, the early stages of
development in apm1— cells (e.g., aggregation) were
normal as well as the morphology of each developmental step (our unpublished
results). However, the overall kinetics of development was delayed
(Figure 6C). After 29 h,
although development of wild-type cells was complete,
apm1— cells showed finger-like structures. After 44
h of development, fruiting body-like structures appeared, though the stalks
were very thin in comparison to wild-type structures. Interestingly, in HL5
medium, only 10% of mutant spores germinated and gave rise to amoeba compared
with 70% for control spores. Stable transfection of native µ1 complemented
both growth and developmental defects. However, µ1 truncated of the 18
C-terminal residues (µ1Ct) did not restore these functions (our
unpublished results).
Thus, in Dictyostelium, AP-1 seems to be important for cell
growth, development, and spore germination. Notably, the µ1Ct did
not rescue any defects establishing that the absence of a fully functional
µ1 chain accounts for the observed phenotypes in
apm1— cells.
Lysosomal Enzyme Precursors Are Secreted in apm1—
Cells
In mammalian cells, AP-1 is required for the transport of newly synthesized
lysosomal enzymes between the TGN and lysosomes. Accordingly, defects in
lysosomal enzymes transport lead to secretion of enzyme precursor forms into
the culture medium (Le Borgne and Hoflack,
1998). To test if AP-1 could play a similar role in
Dictyostelium, we analyzed whether the precursor forms of two
lysosomal enzymes (-mannosidase and cathepsin D) could be detected in
culture medium from apm1— mutant cells.
Dictyostelium -mannosidase is synthesized as a 140-kDa
precursor, which, upon transport to lysosomes, is processed into peptides of
60 and 58 kDa (Mierendorf et al.,
1983,
1985;
Cardelli et al.,
1986). In wild-type cells, no precursor form (pro) was detected in
the culture medium after 10 h of incubation
(Figure 7A) and most of the
enzyme in the intracellular fraction (P) was found in its mature lysosomal
form (mat). In contrast, in apm1— cells, the
-mannosidase precursor was detected in the culture medium (48% of total
enzyme secreted after 10 h) and less mature form (mat) accumulated relative to
the amount of secreted precursor (Figure
7A). Complementation of mutant cells with native µ1, but not
with µ1Ct (38% of enzyme secreted after 10 h), reverted this
phenotype, although more intracellular mature form was detected in
apm1— + µ1Ct than in
apm1— cells, suggesting that µ1Ct could
partially restore AP-1 sorting functions. The fate of cathepsin D, another
lysosomal enzyme, was also assayed and identical maturation and secretion
defects were observed (our unpublished results).
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These results demonstrate that AP-1 is required for the targeting of two
lysosomal enzymes to lysosomes, indicating that Dictyostelium and
mammalian AP-1 adaptor complexes share similar functions. To exclude the
possibility that apm1— mutant cells display a
general membrane trafficking defect, we analyzed the internalization rate of a
cell surface transmembrane protein (p80) known to be constitutively
internalized (Ravanel et al.,
2001). Wild-type cells showed 80% of p80 endocytosis after 30
min monitored by flow cytometry, and the rate of endocytosis was similar in
apm1— cells (our unpublished results), excluding a
major defect in membrane protein endocytosis in mutant cells.
Endocytic Compartments in apm1— Cells
As the transport of lysosomal components is impaired in
apm1— cells, the apm1—
deletion could also affect the overall biogenesis and/or morphology of the
endocytic compartments. To assess this possibility, we analyzed the
intracellular distribution of a transmembrane endosomal protein, p80, in
wild-type and mutant cells. The p80 protein is present at the cell surface but
also throughout the endocytic pathway. As previously described
(Ravanel et al.,
2001), in wild-type cells, p80 was detected in early endocytic
compartments (characterized by a low concentration of p80) and also in late
endocytic compartments (characterized by a high concentration of p80;
Figure 7B). In
apm1— cells, p80 was similarly distributed in
endocytic compartments with high and low p80 content. However, although this
marker was normal in location, the size of these endocytic compartments
appeared smaller and the number of vacuole structures was significantly
increased (Figure 7B). To
demonstrate that these morphological defects were not restricted to
p80-containing vacuoles, cells were allowed to internalize the fluid-phase
marker FITC-dextran for 60 min to label all endocytic compartments and
analyzed by epifluorescence microscopy. In these conditions, identical defects
in size and number of vacuoles were observed and the expression of
µ1Ct in apm1— cells did not restore the
wild-type phenotype (Figure
7C). Electron microscopy analysis further confirmed these results.
In contrast to wild-type cells that displayed several large vacuoles,
apm1— cells displayed only one or two medium-size
vacuoles surrounded by many small vacuoles
(Figure 8). Finally the
localization of two other endosomal markers, the actin binding protein coronin
(early and late endosomes) and the coat protein vacuolin (postlysosomal
vacuoles), was also found identical in apm1— and
wild-type cells (Figure 7B and
our unpublished results). Therefore, apm1— mutant
cells present a normal organization of the endocytic pathway but an altered
morphology of endocytic organelles.
|
AP-1 Is Required for Transport of Proteins to the Contractile Vacuole
Complex
To live in hypo-osmotic condition, Dictyostelium cells have
developed a specialized compartment, the contractile vacuole complex (CV).
This osmoregulatory organelle is composed of large cisternae (bladder) and
interconnecting ducts. The cisternae fuse periodically with the plasma
membrane to expel water from the cells. Because clathrin-deficient cells
display osmoregulatory defects and the CV complex is missing
(O'Halloran and Anderson,
1992), we first assayed the function of the CV complex in
apm1— mutant cells. Wild-type and mutant cells were
shifted from culture medium to water and observed >30 min. Rapidly, both
cell types swelled and rounded up because of water import. Although wild-type
cells rapidly adjusted to the osmotic stress and returned to their original
shape, apm1— mutant cells stayed swollen
(Figure 9). The expression of
native µ1, but not of µ1Ct, allowed
apm1— mutant cells to regain the ability to adjust
to osmotic stress (Figure
9).
|
Next we examined the integrity of the CV complex by studying the
localization of two CV resident proteins, Rh50 (Dictyostelium rhesus
protein; Benghezal et al.,
2001) and VatA (a subunit of the peripheral
vacuolar-H+-ATPase V1 domain;
Zhu and Clarke, 1992;
Jenne et al., 1998;
Neuhaus et al., 1998;
de Hostos, 1999) in
apm1— mutant cells. In wild-type and
apm1— + µ1 cells, Rh50 localized exclusively to
the CV network, whereas VatA was mainly present in the CV network but also was
detected in early endosomes. In contrast, Rh50 and VatA did not colocalized to
the CV network in apm1— and
apm1— + µ1Ct mutant cells
(Figure 10A). Instead, Rh50
was mainly found on punctuate structures in the Golgi area stained with the
anticomitin antibody, whereas VatA was mainly detected in vacuoles, which were
likely endosomes because they contained p80
(Figure 10B). In addition,
Calmodulin, another CV component (Ultricht
and Soldati, 1999), also redistributed in endosomes in
apm1— cells, and the characteristic CV network was
not detected in apm1— mutant cells incubated with FM
4–64 (a fluorescent dye known to label the CV complex; our unpublished
results). Thus, no recognizable CV network can be detected in
apm1— cells, although it is not clear whether
Rh50-positive structures correspond to altered nonfunctional contractile
vacuoles or to mislocalized Rh50, for instance to the Golgi apparatus.
|
The absence of functional CV and the mis-localization of CV components
suggested that the morphology of the CV complexes was altered in
apm1— mutant cells. In electron microscopy images,
CV appears as large translucent vacuoles
(Chastellier et al.,
1978; Quiviger et
al., 1978). Electron microscopy analysis of thin section
series of apm1— mutant cells did not reveal the
presence of large vacuoles characteristic of CV organelles
(Figure 8).
|
DISCUSSION |
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|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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Molecular Characterization and Dynamics of Dictyostelium AP-1
The high homology of -adaptin and µ1 chains with corresponding
mammalian AP-1 subunits strongly suggested that these chains were subunits of
a same AP-1 complex. This assumption was confirmed by showing that these
chains assemble with putative 1 and 1 subunits to form in a
high-molecular-weight complex of 300 kDa, consistent with the size of an
heterotetrameric AP complex (Figure
1A). Although these 1 and 1 chains were not
characterized in this study, blast searches in available EST and genomic data
basis indicate potential genes encoding 1 (contig11712 encoding a
protein of 927 aa sharing 52% identify and 28% similarity with mouse 1)
and 1 subunits (cDNA clone SSF227 encoding a deduced protein sequence
of 154 aa sharing 55% identity and 28% similarity with mouse 1).
The identification of this AP complex as Dictyostelium AP-1 was
further established by studying its intracellular localization. Subcellular
fractionation studies indicate that the AP-1 complex is mainly associated with
Golgi-enriched fractions (Figure
1C). Confocal microscopy analyses show that AP-1 is concentrated
in the Golgi area (Figure 3A)
as well as vesicular structures. Finally, a -GFP fusion protein
expressed in Dictyostelium cells localizes to this same perinuclear
region (Figure 3A), which
undergoes a reversible fragmentation when cells are treated with DMSO, an
agent that affects Golgi integrity (Figure
3B). Therefore the intracellular localization of AP-1 in
Dictyostelium is similar to that observed for mammalian AP-1
complexes (Robinson,
1989).
In mammalian and yeast cells, AP-1 has been reported to be involved in
retrograde (endosomes to TGN) transport of several proteins
(Zizioli et al.,
1999; Meyer et al.,
2000; Valdivia et
al., 2002). However, live microscopy studies of cells
expressing YFP-tagged-µ1A chains indicates that AP-1–coated structure
moving toward the TGN are hardly observed
(Huang et al., 2001),
thus leaving open the debate on the transport pathway in which operate AP-1
complexes. The incorporation of a functional -GFP protein into AP-1
complexes allowed us to examine the dynamics of the AP-1 coat in
Dictyostelium living cells. As described in mammalian cells
(Huang et al., 2001),
AP-1 labeled with -GFP is detected in vesicles, spherical and tubular
structures moving along a same track from the TGN to the cell periphery and
very rarely toward the TGN. Therefore the main function of AP-1 in
Dictyostelium appears to be in anterograde (TGN to endosomes)
transport. Surprisingly, we also detected vesicles moving back and forth
between TGN and endosomes. This observation suggests that AP-1 vesicles could
rapidly deliver cargo proteins to endosomal compartments without fusion
between vesicles and endosomes and the loss of vesicles integrity.
Alternatively, this movement could reflect the Golgi dynamics, which has been
reported to varied between a compact organization and a dispersed structure
accompanied by fast protrusions of Golgi tubules and vesicles
(Schneider et al.,
2000). Further studies will be required to test these
hypotheses.
An essential feature of AP-1 complexes in all species is to facilitate the
recruitment of cytosolic clathrin onto nascent vesicles. This process occurs
through a direct interaction between clathrin and sequences in the hinge
domains of 1 and -adaptin
(Shih et al., 1995;
ter Haar et al.,
2000; Doray and Kornfeld,
2001). As expected, immunofluorescence microscopy studies in
Dictyostelium cells establish that -adaptin mainly localizes
to clathrin-coated vesicles (Figure
3A). Furthermore, GST- fusion proteins comprised of either
the hinge and appendage domains or only the appendage region displayed
clathrin-binding abilities (Figure
3C) as reported for mammalian subunit of AP-1
(Doray and Kornfeld, 2001).
Taken together, these results support the conclusion that we have identified a
bona fide AP-1 complex in Dictyostelium with features comparable to
that of mammalian AP-1.
Dictyostelium, A Model Organism To Study the Function of AP-1
Yeast genetic studies aimed at understanding the function of AP-1 have been
hindered by the absence of major trafficking defects in µ1-deleted strains
(Stepp et al., 1995).
On the contrary, deletions of µ1A and chains in mice lead to
embryonic lethality, also preventing functional analysis of AP-1 in vivo
(Zizioli et al.,
1999; Meyer et al.,
2000). To overcome this problem, we took advantage of the
molecular characterization of AP-1 in Dictyostelium and of the
molecular genetic techniques well developed in this model organism. The
apm1 gene, encoding Dictyostelium µ1 chain, was disrupted
by inserting a blasticidin resistance marker gene. In contrast to yeast cells
and mice, apm1 deletion results in viable cells with several
phenotypic alterations, indicating that Dictyostelium is a unique and
valuable model to study the function of AP-1 in vivo.
µ1 Is Not Required for AP-1 Subunits Assembly and Membrane
Recruitment
In apm1— mutant cells, other AP-1 subunits are
still able to form partial AP-1 complexes
(Figure 5B), which are
correctly localized to the Golgi apparatus
(Figure 5D). These results
demonstrate that µ1 is dispensable for the specific docking of AP-1 to
Golgi membranes. However, these partial complexes are less stably associated
with membranes because only 52% of total -adaptin are detected in
membranes prepared from apm1— mutant cells compared
with 87% for wild-type cells (Figure
5C). Because AP µ chains are involved in the recognition of
tyrosine-based signals found on transported proteins, our results demonstrate
that the recognition of cargo proteins by µ1 chains is not necessary for
membrane recruitment of AP-1 in Dictyostelium cells but contributes
to the efficiency or/and the stability of the interaction.
How is AP-1 deprived of µ1 subunit still targeted to the right
compartment? One possibility is that another AP-1 subunit (e.g., 1)
could also participate in the recognition of cargo proteins and therefore this
interaction could be sufficient for specific binding of partial AP-1 complexes
to Golgi membranes. On the other hand, determinants responsible for membrane
localization of APs have been identified in and chains of AP-2
and AP-1 complexes, respectively (Page and
Robinson, 1995). More recently, AP-2 complexes have been shown to
interact with phosphoinositides through two phospholipid binding domains in
the chain and the surface of µ2 Ct domain
(Gaidarov and Keen, 1999;
Collins et al., 2002;
Rohde et al., 2002).
Sequence alignment of and µ2 with AP-1 and µ1,
respectively, indicates that AP-1 could also bind phospholipids
(Collins et al.,
2002). Accordingly, the recognition of Golgi specific
phospholipids by -adaptin could be sufficient for membrane binding of
partial AP-1 complexes in apm1— cells. This
hypothesis is in agreement with the finding that AP-1 can be recruited in an
ARF-1 dependent manner to protein-free soybean liposomes
(Zhu et al., 1999).
However, the presence of peptides containing tyrosine-based signals in
liposomes clearly enhances AP-1 recruitment to soybean liposomes, indicating
that cargo proteins are also involved in AP-1 recruitment
(Crottet et al.,
2002). Our results are in agreement with these in vitro studies
because native AP-1 associates with membranes more efficiently than AP-1
complexes without µ1 chains (Figure
5C). Notably, AP-1 complexes deprived of mouse µ1A subunits do
not associate with membranes (Meyer et
al., 2000) suggesting that in mammalian cells, lipid and
cargo interactions through other AP-1 subunits are not sufficient for the
recruitment of partial AP-1 complexes to Golgi membranes. This discrepancy
between Dictyostelium and mammalian AP-1 complexes could reflect
their respective affinity for sorting signals and membrane lipids between
species.
µ1 Deletion Affects AP-1 and Clathrin Functions
Clathrin heavy chain null mutant in Dictyostelium
(chc— mutant) displays pleiotropic defects, including growth
and developmental defects, as well as the inhibition of several transport
events along the endocytic pathway
(O'Halloran and Anderson,
1992; Ruscetti et
al., 1994; Niswonger and
O'Halloran, 1997). Clathrin is known to be recruited by several
proteins at different intracellular locations, preventing thus to analyze the
precise role of clathrin-coated structures in any individual transport
pathway. Our study allows for the first time to ascribe some defects observed
in chc— mutant cells to the AP-1 adaptor complex.
As reported for chc— cells,
apm1— cells are viable but grow slower than wild-type
cells (Figure 6, A and B). In
addition, mutant cells show a much weaker defect in Dictyostelium
development compared with chc— cells, suggesting that other
clathrin-associated coats are involved in Dictyostelium development.
The fact that Dictyostelium AP-4 medium chain is upregulated during
development reinforces this possibility
(de Chassey et al.,
2001).
In Dictyostelium as in mammalian cells, clathrin has been
implicated in the transport of lysosomal enzymes between the Golgi apparatus
and lysosomes. For instance, chc— mutant cells secrete
immature forms of the hydrolase, -mannosidase
(Ruscetti et al.,
1994). In apm1— cells, the precursor
form of this enzyme is also secreted
(Figure 7A), supporting the
conclusion that Dictyostelium AP-1 is required for the transport of
lysosomal enzymes from TGN to lysosomes. However, this defect is only partial
because mature -mannosidase is still detected intracellularly. Cell
surface internalization and further processing of the secreted immature
precursor is likely to account for this residual mature enzyme as reported for
mannose-6-phosphate receptors in µ1A deleted fibroblasts
(Meyer et al., 2000).
Alternatively, Dictyostelium hydrolases could be transported to
lysosomes through an AP-1–independent mechanism, for instance through a
GGA (Golgi-localized -ear–containing Arf-binding proteins)
mediated transport pathway as described for several proteins in other
organisms (Black and Pelham,
2000; Dell'Angelica et
al., 2000; Hirst et
al., 2000; Zhdankina
et al., 2001).
In mammalian cells, transport of lysosomal enzymes is mediated by MPRs. As
MPRs have not been identified in Dictyostelium, our results suggest
the existence of another lysosomal enzyme receptor whose trafficking is AP-1
dependent. Recently, targeted disruption of the AP-1 µ1A gene in mice has
revealed that AP-1 may act in MPRs retrieval from early endosomes to the TGN
(Meyer et al., 2000).
This retrieval function was also described in yeast, where the disruption of
the µ1 gene inhibits Golgi retrieval of chitin synthase III and syntaxin
Tlg1p (Valdivia et al.,
2002). Our data in Dictyostelium are consistent with the
role of AP-1 in this endosomes to TGN retrograde pathway, because a block in
recycling of MPR like receptor would also lead to a general defect in
lysosomal enzyme targeting. However, the Golgi steady state localization of
AP-1 and the fact that AP-1–coated structures moving toward the Golgi
area are hardly observed suggest that in Dictyostelium, AP-1 is
mainly involved in sorting putative MPR-like proteins at the TGN level. The
reason for this discrepancy is unknown. Studies on other clathrin-dependent
transport processes from the TGN could provide missing clues.
The morphology of the endocytic pathway is affected both in
apm1— and chc— cells. Despite of
lysosomal enzyme targeting defects, apm1— mutant
cells present a normal localization of several other endocytic protein markers
(Figure 7B), suggesting that
the overall organization of the endocytic pathway is not altered. In contrast,
the morphology of the endocytic organelles is modified in
apm1— as well as in chc— cells.
Conventional thin-section electron microscopy reveals the absence of large
vacuoles in AP-1–deficient cells and a higher density of small vacuoles
(Figure 8). Endo-lysosomal
organelles are known to undergo repeated homotypic and heteropypic fusion
events, leading to the formation of big vacuoles referred to as postlysosomes
or secretory lysosomes (Neuhaus et
al., 2002). It is tempting to propose that clathrin/AP-1
complexes could be implicated in sorting events essential for the biogenesis
of late endocytic compartments. For instance, essential components of the
fusion machinery (e.g., SNARES) required for specific fusion events between
vacuoles could be incorrectly addressed in the absence of functional AP-1
complexes, thus preventing the formation of large vacuoles. Conversely,
AP-1–coated vesicles could contribute to the constant flux of membrane
required to establish and maintain the structure of these organelles.
Interestingly, apm1— mutant cells display a reduced rate of fluid-phase endocytosis and phagocytosis (our unpublished results), whereas endocytosis of a plasma membrane receptor is not affected. In chc— cells, all these endocytic pathways are altered to some extent. Although receptor-mediated endocytosis in Dictyostelium is likely to involved AP-2 clathrin associated adaptor complexes, our results suggest that AP-1 could play a role in pinocytosis and phagocytosis. This possibility is currently under investigation.
Finally, the contractile vacuole complex, an osmoregulatory organelle in
Dictyostelium also involved in Ca2+ regulation
(Moniakis et al.,
1999), is an integral compartment distinct from endosomes and the
plasma membrane. Although endosomes and CV share common components
(Bush et al., 1994;
Temesvari et al.,
1996), these two compartments are apparently not connected to each
other because no protein and membrane exchanges occur
(Gabriel et al.,
1999; Clarke et al.,
2002). Clathrin-coated vesicles have been proposed to play an
essential role in the biogenesis of CV complexes by transporting proteins and
membrane required for CV formation
(O'Halloran and Anderson,
1992). Here we show for the first time, that AP-1 is involved in
the targeting of two CV components. Rh50, a membrane protein exclusively
restricted to the CV network (Benghezal
et al., 2001), is mostly retained in the Golgi apparatus
in AP-1 deficient cells whereas the vacuolar H+-ATPase (VatA) is
routed to endosomes, its other physiological localization
(Figure 10). Targeting defects
of these CV components correlate with the absence of functional and
morphologically detectable CV complexes (Figures
8 and
9). Together these results
suggest that AP-1 is implicated in the biogenesis of the CV complex.
In conclusion, our study demonstrates that the genetically tractable model organism, D. discoideum, is a valuable model for studying the function of AP vesicular coat proteins in vivo. Further genetic approaches are now being used to identify additional components implicated in the AP-1–dependent transport machinery.
|
ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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|
Footnotes |
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# Corresponding author. E-mail address: f.letourneur{at}ibcp.fr.
|
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)
and apm1— cells (
) was fractionated on a
Sephacryl S-300 HR gel filtration column. Fractions were collected and
proteins were analyzed by Western blotting using the
anti–
) cells grow slower
than wild-type (
)
cells. (C) The kinetics of development of the apm1—
mutant is delayed. For development analysis, cells were plated on 0.45-µm
membrane filters laid on Na/K phosphate buffer and incubated at 24°C in
humid chambers for the indicated periods of time. Although after 29 h the
development of wild-type cells was complete, apm1—
cells showed finger-like structures. After 44 h of development, fruiting
body-like structures appeared, though the stalks were very thin in comparison
to wild-type structures.
