From INSERM U25, 161 rue de Sèvres, 75743 Paris cedex 15, France
Received for publication, December 13, 2000, and in revised form, February 28, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The transporters associated with antigen
processing (TAP1/TAP2) provide peptides to MHC class I molecules in the
endoplasmic reticulum. Like other ATP-binding cassette proteins, TAP
uses ATP hydrolysis to power transport. We have studied peptide binding to as well as translocation by TAP proteins with mutations in the
Walker A and B sequences that are known to mediate ATP binding and
hydrolysis. We show that a mutation in the TAP1 Walker B sequence reported to abrogate class I expression by a lung tumor does not affect
ATP binding affinity, suggesting a defect restricted to ATP hydrolysis.
This mutation reduces peptide transport by only 50%, suggesting that
TAP function can be highly limiting for antigen presentation in
non-lymphoid cells. Single substitutions in Walker A sequences
(TAP1K544A, TAP2K509A), or their complete replacements, abrogate
nucleotide binding to each subunit. Although all of these mutations
abrogate peptide transport, they reveal distinct roles for nucleotide
binding to the two transporter subunits in TAP folding and in
regulation of peptide substrate affinity, respectively. Alteration of
the TAP1 Walker A motif can have strong effects on TAP1 and
thereby TAP complex folding. However, TAP1 Walker A mutations
compatible with correct folding do not affect peptide binding. In
contrast, abrogation of the TAP2 nucleotide binding capacity has little
or no effect on TAP folding but eliminates peptide binding to TAP at
37 °C in the presence of nucleotides. Thus, nucleotide binding to
TAP2 but not to TAP1 is a prerequisite for peptide binding to TAP.
Based on these results, we propose a model in which nucleotide
and peptide release from TAP are coupled and followed by ATP binding to
TAP2, which induces high peptide affinity and initiates the transport cycle.
The transporters associated with antigen processing
(TAP)1 belong to the family
of ATP binding cassette (ABC) transporters, a large group of proteins
that use energy provided by nucleotide triphosphates to translocate a
vast variety of substrates across intracellular or cell surface
membranes (1). All ABC transporters possess two transmembrane domains,
each generally composed of six membrane-spanning segments, and two
nucleotide binding domains (NBDs) with primary sequence homology across
the protein family. Whereas substrate interaction is generally thought
to involve the transmembrane domains, the NBDs bind and hydrolyze ATP.
Both of the latter events have been shown to lead to conformational changes that upon transmission to substrate binding domains in an
undefined fashion result in substrate translocation (2).
Given the important role of ABC transporters in diseases such as
mucoviscidosis or cancer, the mechanism of substrate transport has been
subject to intense scrutiny. Mutagenesis studies and crystallographic
analysis of HisP and MalK, the NBDs of bacterial histidine and maltose
transporters, respectively, have elucidated the role of several
conserved sequence motifs contained in ABC transporter NBDs (3, 4).
Thus, the Walker A consensus sequence (5), also termed P-loop, engages
in limited interactions with the ATP adenine ring and extensive
interactions with its Several models have been formulated that link ATP metabolism to
substrate transport (2). These models generally postulate that,
depending on the nature of the bound nucleotide, NBDs can assume at
least three states: an ATP-associated state, followed by a short-lived
transition state with ADP and inorganic phosphate (Pi)
bound, and an ADP-bound state. Transition from one state to the next is
thought to be coupled to re-orientation of the corresponding substrate
binding site and/or a change in substrate affinity. ATP metabolism by
the two NBDs of an ABC transporter is generally believed to be linked
in an allosteric fashion, as mutations in one NBD affect ATP processing
by the other NBD, at least in most cases. Indirect evidence suggests
that ATP metabolism by some ABC transporters proceeds in an alternating
fashion, such that only one NBD at a time hydrolyzes ATP. This
conclusion is mainly based on "vanadate trapping" experiments in
which only one of the two NBDs of P-glycoprotein (P-gp), a drug efflux
pump overexpressed in many tumors, can be "frozen" in the
ADP-Pi bound transition state (6). Recent studies suggest
that LmrA, a bacterial analog of P-gp, also uses a "two cylinder
engine" model and elegantly provide evidence on how orientation and
affinity of the (in this case presumably two) substrate binding sites
change during the catalytic cycle (7). Whereas the P-gp NBDs are
thought to metabolize ATP in an alternating but otherwise identical
fashion (8), recent evidence suggests that other ABC transporters may
have assigned distinct roles to the two NBDs. Thus, the multidrug
resistance protein (MRP1), an organic anion exporter also transporting
many anti-cancer drugs, appears to hydrolyze ATP preferentially or initially after substrate binding at its NBD2 (9-11). Among several interpretations of these non-equivalent roles of the MRP1 NBDs, the
hypothesis that nucleotide binding to NBD1 may regulate nucleotide hydrolysis by NBD2 is noteworthy (9).
TAP1 and TAP2 form a heterodimeric complex in the membrane of the
endoplasmic reticulum and proximal Golgi that provides antigenic peptides to newly synthesized and assembling MHC class I molecules (12). TAP is essential for MHC class I-restricted antigen presentation, as demonstrated by absent or low MHC class I expression in
TAP-deficient mice and cell lines (13). Tumor cell lines frequently
show impaired TAP expression and/or function, underlining the key role
of TAP in MHC class I-mediated immune surveillance and prevention of tumor growth (14). Several herpes virus proteins block TAP function, presumably through different mechanisms involving conformational change
(15). Thus, diminution or loss of TAP function plays a role in the
pathogenesis of viral and malignant diseases. These perturbations of
TAP function may be related to mutations or protein interactions
interfering with ATP metabolism; indeed, a case of a human lung tumor
with impaired TAP function because of a mutation in the Walker B region
has been described (16).
Little is known on ATP metabolism by the TAP transporters. ATP
hydrolysis is required for peptide transport by TAP (17). We have
recently described that at 37 °C, microsomal TAP requires stabilization by nucleotide di- or triphosphates to maintain a conformation with high substrate affinity (18). Similarly, Knittler et al.(19) reported that substitutions in the TAP Walker A
sequences not only abolished peptide binding to, and transport by, TAP
but also inhibited the physiological dissociation of peptide-assembled MHC class I molecules from TAP. In this study, we used an insect cell
expression system to study substrate interaction with TAP proteins
carrying mutations in Walker A and B sequences. Our results shed light
on the nucleotide requirements for TAP folding and for peptide binding
and transport and suggest that the NBDs of TAP1 and TAP2 fulfill
distinct functions in the catalytic TAP cycle.
Mutant TAP Proteins--
Baculoviruses encoding human wild type
(wt) TAP1.A and TAP2.A proteins with mutations in the Walker sequences
were produced by oligonucleotide-directed "loop out" mutagenesis on
uracil-enriched single-stranded DNA as described previously (20).
cDNAs containing the desired mutations, confirmed by sequencing,
were cloned into the baculovirus transfer vector pVL1392 (Invitrogen,
San Diego, CA), and co-transfected with BaculoGoldTM (PharMingen, San
Diego, CA) virus DNA into Sf9 insect cells to produce
recombinant viruses as described previously (20).
Production of Microsomes--
Insect cell microsomes expressing
mutant TAP proteins were produced by sucrose gradient fractionation of
mechanically lysed infected Sf9 cells according to previously
published procedures (21). A mixture of protease inhibitors was added
before insect cell lysis (chymostatin, phenylmethylsulfonyl fluoride,
pepstatin, aprotinin, and leupeptin; see Ref. 20 for details).
Peptide Binding and Transport Assays--
TAP function was
evaluated in peptide binding and transport assays performed exactly as
described previously, using 125I-labeled reporter peptides
R-9-L (RRYNASTEL) and R-10-T (RYWANATRST) for binding and transport
assays, respectively (20). In preincubation experiments, microsomes
(15-30 µl in 150 µl of phosphate-buffered saline with 0.1 mg/ml
bovine serum albumin, 1 mM dithiotreitol, and 2 mM MgCl2) were incubated for 15 min at
37 °C, cooled to 0 °C in a water/ice bath, pelleted (20,000 × g for 5 min), resuspended in 150 µl of assay buffer,
and used in a standard binding assay.
Immunoprecipitation, Western Blot, and Flow Cytometric
Analysis--
These experiments were also performed as described
previously by us (18, 22). For immunoprecipitation, 30-µl (for
vesicles prepared up to three months prior to the experiment) or
60-µl microsomes (for older vesicles) were lysed in a total volume of 600 µl using Nonidet P-40, and TAP complexes were immunoprecipitated with monoclonal antibody (mAb) 148.3 specific for human TAP1 (23). Proteins blotted onto polyvinylidene difluoride membranes were visualized with mAbs 148.3 or 429.3 (anti-TAP2) and a standard enhanced
chemiluminescence protocol. For flow cytometric analysis, Sf9
cells were infected with TAP1 and TAP2 viruses, both at a multiplicity
of infection of 6, together with a virus encoding HLA-B27 and human
ATP Binding Assay--
5 × 106 Sf9
cells were infected with one or two TAP viruses, each at a multiplicity
of infection of 10 and incubated for 72 h before harvesting.
Pelleted cells were lysed for 40 min at 4 °C in 750 µl of a 10 mM Tris, pH 7.4 buffer containing 10 mM sodium phosphate, 130 mM NaCl, 5 mM MgCl2,
and 1% Triton X-100. After lysate clarification by centrifugation
(40,000 × g, 40 min), 75-250 µl of lysate
(corresponding to 0.5-1.6 × 106 cells) was diluted
to 300 µl and incubated for 2 h at 4 °C with 50 µl of
packed ATP-agarose beads (11 atom spacer, Sigma-Aldrich, St.-Quentin,
France) in the presence or absence of ATP or ADP. Then beads were
washed three times with 1 ml of the same buffer, with 0.5% Triton
X-100, and finally resuspended in 30 µl of SDS-PAGE sample buffer. 7 µl were then loaded on 7.5% SDS-PAGE gels, and precipitated TAP
proteins were detected by Western blot analysis using mAb 148.3 and
429.3, as described above.
Insect Cell Expression of Mutant TAP Proteins--
To study the
role of nucleotides for TAP function, TAP proteins with mutant Walker
sequences were designed and expressed in the Sf9 insect
cell/baculovirus system. Mutants included single residue
substitutions as well as complete replacement or deletion of
Walker A sequences (Table I).
Point substitutions in the Walker A motif were designed according to
published studies. Substitution of the highly conserved Lys residue
(mutants T1K544A and T2K509A), and of the amino-terminal Gly residue
(mutant T1G538Q) have been reported to abrogate ATP hydrolysis and in
some cases also binding by different ABC transporters (2, 25-27). In
HisP, the corresponding residues interact directly with each other, and
the Lys residue forms several hydrogen bonds with the
Recombinant baculoviruses containing mutant TAP cDNAs were
generated, and expression and assembly of TAP proteins were studied in
Western blotting and immunoprecipitation experiments (Fig. 1). All mutant TAP1 and TAP2 proteins
were expressed and migrated as expected in SDS-PAGE gels. However,
whereas most mutant TAP subunits were overexpressed to a similar extent
as wild-type TAP1 and TAP2, only small amounts of mutant TAP1 subunit
T1Del could be immunoprecipitated, and recovery of mutant T1G538Q was
also reduced. Reduced levels of immunoprecipitable T1Del and T1G538Q proteins reflected reduced levels of mutant proteins in microsome lysates prepared using Nonidet P-40 (see below); however, relatively large amounts of these subunits were detected in microsome "debris" that could not be solubilized in Nonidet P-40 (not shown). This suggested that mutants T1Del and T1G538Q folded inefficiently and
formed insoluble aggregates. Nevertheless, even in these cases, immunoprecipitable material, presumably corresponding to the correctly folded fractions, assembled efficiently in heteromeric TAP complexes, as indicated by co-immunoprecipitation (Fig. 1). Thus, none of the
mutations had an adverse effect on assembly with wild-type complementary subunits.
Peptide Translocation and Supply to HLA-B27 Molecules by Mutant TAP
Complexes--
To analyze the effect of the mutations on TAP transport
function, insect cells expressing mutant TAP subunits together with wild-type complementary subunits were produced and tested in two assays. One of these evaluates accumulation of a glycosylated radiolabeled peptide in microsomes (17), whereas the other uses appearance of HLA-B27 molecules on the surface of infected insect cells
as a readout for TAP function. We have previously shown that insect
cell-expressed B27 molecules are loaded with TAP-supplied peptides in
the absence of other human accessory molecules, notably tapasin, and
that cell surface expression in the Sf9 insect cell system
reflects intracellular peptide loading of HLA-B27 (22). In this study,
we analyzed peptide loading of HLA-B27 as an additional and
physiologically relevant readout for TAP function. Moreover, because
tapasin-independent loading of B27 with TAP-supplied peptides is likely
to be related to the exceptionally high compatibility between TAP and
B27 peptide preferences (28), we reasoned that this assay might also
provide information on potential changes in TAP selectivity resulting
from mutations.
As expected, substitutions T1K544A, T1G538Q, and T2K509A, as well as
Walker A replacements and deletions in either TAP subunit, abolished
accumulation of glycosylated peptide in the ER, whereas control
mutation T2ST510/1TS did not affect the rate of peptide accumulation
and T1ST545/6TS increased it (Fig. 2).
Thus, ATP binding to and/or hydrolysis by both NBDs is required for
peptide transport by TAP. Moreover, similar to the CFTR (25), mutation T1ST545/6TS may enhance efficiency of ATP hydrolysis by TAP1. Surprisingly, mutation T1R659Q, reported to create a TAP-deficient cell
phenotype (16), reduced peptide accumulation by about 50% rather than
abolishing it. ATP-independent peptide accumulation after prolonged
microsome incubation at 37 °C is likely to reflect vesicle
"leakiness." ADP, which is known to act as a competitor for ATP in
ABC transporters (25), reduced peptide accumulation in microsomes
expressing wild-type TAP or control mutations (ST to TS inversion) to
levels below those observed without addition of nucleotides. This may
indicate that ADP competed with ATP carried over in microsome
purification for binding to the TAP NBDs.
Analysis of HLA-B27 expression confirmed results obtained in transport
assays (Fig. 3). In accordance with our
previous observation (22), B27 was not detectable on the surface of
insect cells expressing HLAB27/ Peptide Binding to Mutant TAP Complexes at Low Temperature and
37 °C--
Peptide transport is preceded by a high affinity
interaction of peptide with the TAP substrate binding site, which can
be measured in a binding assay performed at low temperature (21). We
have previously shown that wild-type TAP complexes incubated at
37 °C in the absence of nucleotide di- or triphosphates change conformation and lose the ability to bind peptide with high affinity (18). We have therefore proposed that low TAP affinity for peptide substrate characterizes a "nucleotide-off" conformation that may be
related albeit not identical to a transition state associated with
peptide release during the transport cycle. Here we asked how Walker
motif mutations impairing nucleotide binding and/or hydrolysis affect
peptide binding to TAP at different temperatures.
In standard peptide binding assays with TAP-expressing vesicles not
exposed to temperatures above 27 °C (the temperature used for insect
cell culture), microsomes expressing TAP complexes with deletion of the
Walker A sequence in either TAP subunit consistently showed very low
but significant peptide binding. This binding exceeded binding to
control vesicles expressing TAP1 by a factor of 10-15 (Fig.
4). Microsomes expressing mutant T1G538Q
also consistently displayed significantly reduced peptide binding
capacity. Peptide binding to mutant T2Rep was moderately reduced,
whereas other TAP1 mutants, including those with Walker A replacement,
bound normal peptide amounts.
When microsomes were incubated for 15 min at 37 °C prior to binding
assays, all TAP complexes lost binding capacity (Fig. 5). This included complexes composed of
two mutant subunits, for example T1Rep/T2Rep or T1K544A/T2K509A (not
shown). Thus, ATP hydrolysis was not required for loss of high affinity
peptide binding by TAP. Addition of ATP or ADP during incubations at
37 °C preserved peptide binding capacity by some but not all TAP complexes containing mutated subunits. TAP1 mutations T1Del and T1G538Q
abolished TAP complex stabilization by ATP. Complexes containing other
TAP1 mutations, including mutation T1Rep, retained peptide binding
capacity in the presence of ATP or ADP to a similar extent as wild-type
TAP complexes. However, complexes with similar (T2Rep) or identical
(T2K509A) mutations in TAP2 did not retain peptide binding capacity in
the presence of nucleotides. This suggested that binding of nucleotide
di- or triphosphates to TAP2 is essential for maintaining TAP peptide
binding capacity whereas nucleotide binding to TAP1 is not. Moreover,
TAP1 mutants prone to aggregation also could not be stabilized by
nucleotides.
Nucleotide Binding Affinity of Mutant TAP Subunits--
To provide
direct evidence for the role of nucleotide binding, as opposed to
hydrolysis, for peptide binding to TAP, we measured the binding
affinity of mutant TAP subunits for ATP immobilized on agarose beads
(Fig. 6). Initially, we studied ATP
binding of isolated or co-expressed wild-type TAP subunits (Fig.
6A). ATP binding of isolated wild-type TAP1 was efficient
and inhibited with equal efficiency by ADP and ATP but not AMP. In
contrast, isolated TAP2 subunits showed relatively low levels of
Nonidet P-40 soluble material, and ATP binding was hardly detectable. This suggested that isolated TAP2 folded poorly and that correctly folded isolated TAP2 bound ATP poorly. Co-expression of the two wild-type subunits had no effect on the efficiency of TAP1 binding to
ATP but induced a significant increase in the amount of Nonidet P-40
solubilized TAP2, suggesting that correct TAP2 folding is enhanced in
the presence of TAP1. Simultaneously, the amount of TAP2 recovered by
incubation with ATP-agarose increased dramatically and by a factor
largely exceeding the effect of TAP1 on TAP2 folding (Fig.
6A). The latter phenomenon indicated either indirect
recovery of TAP2 (itself with low ATP affinity) via ATP-bound TAP1, or an increase in the ATP binding affinity of TAP1-associated
TAP2.
Because isolated TAP1 subunits can bind ATP and be stabilized by it,
ATP binding affinities of TAP1 mutants were first studied in the
absence of TAP2 (Fig. 6B). Western blot analysis of Nonidet P-40-solubilized material confirmed low expression levels of Nonidet P-40-solubilized T1Del and T1G538Q mutants, whereas all other mutants,
including T1Rep, were expressed at similar high levels. Control
mutation T1ST545/6TS and Walker B mutation T1R659Q had no effect on ATP
binding affinity, whereas all other TAP1 mutations abolished ATP
binding completely. Identical results were obtained with cells
co-expressing TAP1 mutants with wild-type TAP2 (not shown). The latter
experiments also demonstrated that complex formation with TAP1 does not
increase the ATP binding affinity of TAP2. Co-expression of T1K544A
(being itself unable to bind ATP) increased the amount of Nonidet
P-40-solubilized TAP2 as efficiently as wild-type TAP1; however, TAP2
binding to ATP remained very low in its presence (not shown).
Because of the poor folding and ATP binding affinity (Fig.
6A), isolated mutant TAP2 proteins could not be used for ATP
binding experiments. To increase the amount of correctly folded TAP2
material, we chose to co-express a mutant TAP1 protein (T1K544A) that
itself could not bind to immobilized ATP (Fig. 6B).
Moreover, to compensate for inefficient binding of TAP2 to ATP-agarose,
experiments were performed on a 4-fold higher amount of cell lysate.
Western blot analysis of Nonidet P-40 lysates confirmed that, among
TAP2 mutants, only T2Del showed slightly reduced expression levels,
possibly because of reduced folding efficiency (Fig. 6C).
Using relatively large amounts of cell lysate, efficient binding of
T2wt to ATP-agarose was detected. Control mutant T2ST510/1TS also bound
ATP, whereas the other three mutants completely lacked ATP binding.
Thus, both in TAP1 and TAP2, substitutions in as well as replacement
and deletion of the Walker A motifs completely abolished ATP binding.
Conformational Changes in Mutant TAP Complexes at 37 °C--
We
have previously shown that loss of peptide binding capacity by TAP at
37 °C is associated with a conformational change that reduces TAP
recognition by several antibodies to TAP1 and TAP2 (18). Because
peptide binding to TAP complexes containing mutant TAP1 subunits unable
to bind nucleotides was preserved by ATP, we wondered whether this
reflected a dissociation between TAP1 conformation and TAP peptide
binding capacity. An alternative explanation was that nucleotide
binding to TAP2 might stabilize TAP1 indirectly. We therefore
precipitated TAP complexes that had been incubated at 4 or 37 °C for
15 min in the presence or absence of ATP with antibodies to TAP1 and
determined the efficiency of precipitation by Western blot analysis.
These experiments exploit the phenomenon that incubation of TAP1 at
37 °C in the absence of nucleotides induces a conformational change
that reduces recognition of native Nonidet P-40-solubilized TAP1 by mAb
148.3 without affecting its recognition in a denatured state,
i.e. in Western
blots.2 Because wild-type
TAP1 subunits can bind and be stabilized by ATP whereas TAP2 subunits
cannot (18), experiments were also performed with microsomes expressing
TAP1 only.
Similar to wild-type TAP complexes, incubation of isolated mutant TAP1
subunits or TAP complexes containing such subunits at 37 °C leads to
a strong decrease in the amount of recovered TAP proteins (Fig.
7 and data not shown). TAP1 mutations
T1K544A, T1G538Q, and T1Rep abrogated stabilization of the isolated
TAP1 subunit whereas mutations T1ST545/6TS and T1R659Q did not (Fig. 7
and Table II). However, dimeric TAP
complexes containing mutants T1K544A and T1Rep were stabilized
efficiently by ATP, which could only be due to ATP binding to TAP2. ATP
did not stabilize mutant TAP2 subunits expressed in complexes with
wild-type TAP1 or complexes containing mutant T1G538Q (Table II). Taken
together, these experiments demonstrate that peptide binding capacity
of complexes containing mutant TAP subunits is closely associated with
conformational changes. Like peptide binding capacity, TAP conformation
is maintained by nucleotide binding to the TAP2 subunit, which controls
the conformation of the entire TAP complex and stabilizes TAP1 in an
indirect manner.
The mutants produced in this study provide insight into the
regulation of TAP function by nucleotides. We find that the Walker sequences of the two TAP subunits have distinct roles and affect TAP
structure as well as peptide binding and transport. Whereas the TAP1
NBD appears to be important for TAP1 folding and overall complex
stability, the TAP2 NBD controls substrate affinity and more limited
conformational changes.
Analysis of mutant expression, solubility in Nonidet P-40, and
immunoprecipitation provided some insight into the mechanism of TAP
folding and assembly. In the case of TAP1, not only deletion of the
Walker A sequence, but also point mutation T1G538Q strongly decreased
the amount of Nonidet P-40 soluble protein, suggesting that TAP1
folding is influenced by its NBD. However, in the case of mutants
T1K544A and remarkably also T1Rep, high expression levels and normal
peptide binding capacity at 4 °C and 37 °C argue against an
effect on folding. Whereas folding of the isolated TAP2 subunit is much
less efficient than that of TAP1, mutants T2K509A and T2Rep also have
no detectable effect on folding and even deletion of its Walker A motif
affects folding moderately. Taken together, these observations suggest
a model in which assembly of TAP complexes proceeds via initial folding
of the TAP1 subunit, possibly bound to nucleotides, which in turn
serves as a scaffold supporting TAP2 folding and simultaneous dimer formation.
ATP binding assays provided clear evidence that substitution of the Lys
residue in the Walker A sequence, which engages in extensive
interactions with the Not surprisingly, we find that all mutations abolishing nucleotide
binding to either TAP subunit eliminate peptide transport by TAP,
demonstrating that nucleotide interaction of both subunits is required
for transport. This observation is in agreement with a previous study
by Knittler and associates (19). However, to our knowledge,
substitution T1R659Q is the first case to be studied of a TAP mutation
likely to exclusively affect ATP hydrolysis. This mutation reduced TAP
transport by only 50%. Impairment rather than abolition of TAP
function evidently may reflect reduced but not absent ATP hydrolysis.
An interesting alternative interpretation can be formulated with
reference to other ABC transporters such as MRP1. In that case, NBD1
has been proposed to regulate NBD2, thereby explaining a complete
functional knockout by mutation of NBD2 in contrast to 70% loss of
function only by mutation of NBD1 (9). Elucidation of the role of ATP
hydrolysis by TAP1 and TAP2 will require an assay measuring ATP
hydrolysis by TAP, which is as yet unavailable. It is important to
underline that mutations impairing exclusively ATP hydrolysis by TAP2
have not been reported and studied so far, so that a requirement for
ATP hydrolysis by TAP2 remains hypothetical.
Whatever the reason for the limited effect of the T1R659Q mutation on
TAP transport and B27 expression, a 50% function by T1R659Q mutant TAP
proteins is a surprising finding, given that the human small cell lung
cancer line H1436, in which the mutation was found, was described as
essentially "HLA class I negative" (16). Several explanations may
account for this discrepancy. It appears unlikely that H1436 harbors
additional defects in antigen presentation genes as TAP1 transfection
restored class I expression (16). However, Chen et al.
(16) showed that H1436 expresses low TAP1 levels and that
expression of high levels of T1R659Q in H1436 resulted in significant
class I expression by some, albeit not all, tumor cells. Hence, cancer
cells expressing high amounts of T1R659Q may resemble insect cells
overexpressing it, and the phenotype of H1436 may be caused by a
combination of low TAP expression and impaired TAP function. It is also
conceivable that mutation T1R659Q affects TAP interaction with class I
loading complexes, for example by inhibiting release of peptide-loaded
class I molecules (19) or by impairing tapasin interaction (29). In any
case, it is interesting to note that a 50% reduction in TAP function can result in loss of class I expression. TAP may act as a factor limiting antigen presentation by non-lymphoid cells and is therefore an
especially "promising" target for mutations or regulatory proteins that allow for tumor or viral escape from immune surveillance (15,
30).
This study clearly demonstrates that peptide binding to TAP complexes,
i.e. peptide affinity, is controlled by nucleotide interaction with TAP2 but not TAP1. Our results suggest that the TAP
substrate binding site can assume at least two conformations distinguished by their peptide affinities. The low affinity state can
be produced by three experimental manipulations: (i) incubation of
microsomal wild-type TAP complexes at 37° in the absence of nucleotides, (ii) incubation at 37 °C, in the presence or absence of
nucleotides, of TAP complexes with substitutions in the TAP2 Walker A
sequence abolishing nucleotide binding, and (iii) deletion of the
Walker A sequence in TAP2. In other words, low peptide affinity can
either be induced in a temperature-independent fashion by deletion of
the TAP2 Walker A sequence or at 37 °C by absence of nucleotide
binding to it. In contrast, reduced peptide binding to T1Del/T2wt and
T1G538Q/T2wt complexes reflects a reduced amount of correctly folded
TAP complexes formed by these mutants rather than a conformation with
low peptide affinity. The amount of peptide bound by these complexes
corresponds to the amount of immunoprecipitable and presumably
correctly folded complexes (Figs. 1 and 4). Absence of nucleotide
stabilization of these complexes at 37 °C may then indicate that
nucleotide binding to TAP2 cannot prevent aggregation of these
presumably unstable TAP1 subunits. At 37 °C, complexes with
replacements of both Walker A sequences (T1Rep/T2Rep) undergo the
same conformational change associated with loss of peptide binding
capacity as wild-type TAP complexes. Thus, transition to the TAP state
with low peptide affinity can take place in the absence of nucleotide
interaction and appears to reflect a spontaneous conformational change
at 37 °C that is reversed or prevented by nucleotide binding to TAP2.
Nucleotide binding to TAP2 not only controls TAP peptide affinity but
also is sufficient to prevent a substantial conformational change of
TAP complexes at 37 °C. It is important to underline that this
change is different from that induced by mutations T1Del and T1G538Q.
Whereas the mutations lead to formation of aggregates that are
insoluble in Nonidet P-40, incubation at 37 °C does not. Thus, while
TAP1 serves as a scaffold that folds under the control of its NBD and
permits secondary folding and association of TAP2, nucleotide binding
to TAP2 induces a distinct conformation with high peptide affinity. In
controlling substrate affinity by nucleotide interaction with a single
subunit, the TAP complex differs from some extensively studied ABC
transporters such as P-gp that possess two functionally identical NBDs
(6). However, TAP may resemble other ABC transporters such as CFTR or
MRP1 in which the two NBDs seem to have distinct functions (6, 9, 25).
TAP also appears to differ from other ABC transporters with respect to
the nucleotides bound in the state with high substrate affinity.
Whereas other transporters display low substrate affinity (required for
substrate release) when in the ADP-bound state (2, 7), ADP is clearly
indistinguishable from ATP with respect to preserving high peptide
binding affinity of TAP. Thus, TAP complexes may release peptide either
after complete nucleotide dissociation or during the transition state
associated with ADP and Pi. In contrast to P-gp, which can
be frozen in the latter state by incubation with orthovanadate and ATP
(31), an experimental technique for inducing the transitional
conformation in TAP has not been described. However, our observation
that low peptide affinity can be induced by incubation at 37 °C of
TAP complexes unable to bind nucleotides argues for a nucleotide-off
state as an intermediate with low substrate affinity. Our data suggest that this state is followed by nucleotide binding to TAP2, which allows
TAP complexes to acquire a high peptide affinity and initiate a new
transport cycle.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
- and
-phosphates whereas the less
conserved Walker B motif is likely to be involved in attacking the
-phosphate bond during MgATP hydrolysis. Consequently, substitutions
in the Walker sequences generally impair ATP hydrolysis and transporter
function partly or completely (2).
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
2-microglobulin at a multiplicity of infection of 3. Viruses coding for two unrelated proteins, 65-kDa glutamic acid
decarboxylase (GAD), and IA-2, the intracellular fragment of a tyrosine
phosphatase, were used in control infections for cytometric analysis
(24). Cells were harvested 40 h after infection and stained for
cell surface expression of HLA-B27 with mAbs B27.M1 and W6/32. Dead
cells staining with propidium iodide were excluded from analysis.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-phosphate of
ATP (4). Inversion of the carboxyl-terminal Ser-Thr residues (mutants
T1ST545/6TS and T2ST510/1TS) has been reported to enhance efficiency of
MgATP utilization by the cystic fibrosis transmembrane conductance
regulator (CFTR) (25). Finally, mutation T1R659Q corresponds to a
natural mutation in the TAP1 Walker B motif previously identified in a human small cell lung cancer line and was described to be associated with a TAP-deficient cell phenotype (16). Replacements of entire Walker
A motifs by synthetic linker peptides (mutants T1Rep and T2Rep), or
complete Walker A deletions (mutants T1Del and T2Del) were included to
study TAP proteins highly likely to be unable to hydrolyze and bind
ATP.
Sequence of TAP Walker motifs and mutations
View larger version (33K):
[in a new window]
Fig. 1.
Recovery by immunoprecipitation and complex
formation of mutant TAP proteins. 60 µl of microsome solution
were lysed in a buffer containing 1% Nonidet P-40, and TAP complexes
were precipitated from cleared lysates with mAb 148.3 that is specific
for TAP1. Equal parts of the immunoprecipitated material were separated
in two 7.5% SDS-PAGE gels, blotted on polyvinylidene difluoride
membranes, and stained with mAb 148.3 (upper panels) or
429.3 specific for TAP2 (bottom panels). Mutant or wild-type
(wt) TAP proteins expressed by microsomes are indicated
above or below the panels,
respectively. In this and the following figures, inversion mutants
(T1ST545/6TS and T2ST510/1TS) are designated Co (for
control).
View larger version (36K):
[in a new window]
Fig. 2.
Peptide transport by mutant TAP
complexes. Accumulation of glycosylated peptide R-10-T in insect
cell microsomes expressing the indicated TAP1/TAP2 combinations was
measured after incubation at 37 °C for the time indicated. ATP and
ADP were used at 1 mM concentrations.
2-m together with the two unrelated
control proteins GAD and IA-2 (mean fluorescence 5). Co-expression of wild-type TAP complexes induced B27 expression by about 50% of the
cells (mean 21). Expression was limited to 50% because cells were
infected with a low multiplicity of infection of the B27/
2-m virus
together with a high multiplicity of infection of TAP viruses to ensure
TAP expression by all B27-expressing cells. Whereas control mutations
(ST to TS inversions) increased B27 expression to the same extent as
wild-type TAP (means of 20), TAP mutants abolishing peptide transport
had no significant effect on its cell surface density (means between 6 and 7). Importantly, mutation T1R659Q displayed about 50% of the
effect of wild-type TAP (mean 14). Thus, TAP-mediated peptide supply to
HLA-B27 also required ATP binding to and/or hydrolysis by both TAP
subunits and was reduced but not abolished by the Walker B mutation
described to eliminate HLA class I expression on the surface of a tumor
cell line.
View larger version (39K):
[in a new window]
Fig. 3.
Peptide supply by mutant TAP complexes to
HLA-B27/ 2m molecules co-expressed
in insect cells. Sf9 insect cells were co-infected with
baculoviruses coding for HLA-B27/
2m together with
control viruses GAD and IA-2 (upper left panel) or with
viruses encoding indicated TAP subunits. 40 h after infection,
live cells were stained with the B27-specific mAb B27.M1. Equivalent
results were obtained with mAb W6/32.
View larger version (27K):
[in a new window]
Fig. 4.
Peptide binding capacity of mutant TAP
complexes at low temperature. Binding of radiolabeled peptide
R-9-L to insect cell microsomes expressing the indicated TAP1/TAP2
combinations was measured at 4 °C. As precise control of protein
expression in the baculovirus system is impossible, absolute cpm bound
in peptide binding assays vary with microsome batches. Moreover,
peptide binding decreases with prolonged microsome storage at
80 °C. Variations by a factor of 2 (e.g. 1wt/2wt
versus T1K544A/T2wt or T1wt/T2K509A) are therefore
considered not significant.
View larger version (26K):
[in a new window]
Fig. 5.
Effect of nucleotides on peptide binding
capacity of mutant TAP complexes at 37 °C. Microsomes
expressing the indicated TAP1/TAP2 combinations were incubated for 15 min at 37 °C in the absence or presence of 0.5 or 2.0 mM
ATP or ADP, as shown. Then vesicles were cooled to 4 °C, pelleted,
resuspended in fresh assay buffer without nucleotides, and binding of
iodinated peptide R-9-L was measured in a standard binding assay at
4 °C. Peptide binding after preincubation at 4 °C in the absence
of nucleotides is set at 100%.
View larger version (54K):
[in a new window]
Fig. 6.
Binding of mutant TAP proteins to immobilized
ATP. Cells expressing individual TAP subunits or combinations
thereof were lysed in a buffer containing Triton X-100, and TAP
proteins binding ATP were recovered by incubation with ATP-agarose
beads. TAP proteins binding to the beads, and in parallel total
solubilized TAP proteins, were visualized in Western blots stained with
mAb 148.3 or 429.3. In A, Triton X-100-solubilized total TAP
proteins in cells expressing (top to bottom) TAP1 only, TAP2
only, or both wild-type subunits are shown on the left. TAP
proteins recovered from 2.3 × 105 cells by incubation
with ATP-agarose, in the presence of the nucleotides indicated under
the panels, are shown on the right. In
B, the top panel shows total TAP1 proteins
recovered from cells expressing isolated mutant TAP1 subunits, whereas
the lower panel shows ATP-binding TAP1 proteins recovered
from the same cells (1.2 × 105 cell
equivalents/lane). In C, cells expressed TAP1 mutant K544A
together with indicated mutant TAP2 proteins. Total solubilized TAP2
protein and ATP-binding TAP2 protein corresponding to 4 × 105 cell equivalents are shown in the top and
bottom panels, respectively. TAP1 protein expression was
identical in all samples (data not shown).
View larger version (40K):
[in a new window]
Fig. 7.
Nucleotide binding to TAP2 controls TAP1
conformation. Microsomes expressing mutant T1Rep or T1R659Q
subunits alone (five left lanes), or together with the
wild-type TAP2 subunit (right lanes) were incubated for 15 min at 4 °C or 37 °C, in the absence or presence of 0.2 or 2 mM ATP, as indicated between the two panels.
Then vesicles were cooled to 4 °C, and TAP1 protein was
immunoprecipitated using mAb 148.3 and visualized in a Western blot
stained with the same antibody. More efficient stabilization of
T1R659Q/T2wt complexes, as compared with T1Rep/2wt complexes, is caused
by incomplete assembly of overexpressed TAP1 subunits with TAP2
(non-assembled T1R659Q is stabilized by ATP, whereas T1Rep is
not).
ATP stabilization of mutant TAP proteins and complexes
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-phosphate of bound ATP in HisP (4), or
replacement of the entire sequence, abolishes nucleotide binding to
TAP1 and TAP2. Thus, TAP differs from other ABC transporters such as
P-gp in which Lys substitution has been reported to affect ATP
hydrolysis but not binding (27). ATP binding assays also suggested that
TAP2, expressed as an isolated subunit or assembled with TAP1, has
lower ATP binding affinity than TAP1. Finally, comparison of these
assays with our "nucleotide stabilization assay," in which TAP is
immunoprecipitated after incubation at 37 °C, demonstrates that the
latter assay reflects nucleotide binding and can replace ATP binding
assays. Mutation T1R659Q is the only one without effect on ATP binding
and TAP1 stabilization at 37 °C. It is therefore likely to
selectively impair ATP hydrolysis, a conclusion in agreement with the
proposed role of the Walker B residues in coordinating MgATP hydrolysis
(3, 4).
![]() |
FOOTNOTES |
---|
* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed. Tel.: 33-1-44492563;
Fax: 33-1-43062388; E-mail: vanendert@necker.fr.
Published, JBC Papers in Press, April 4, 2001, DOI 10.1074/jbc.M011221200
2 P. van Endert, unpublished results.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: TAP, transporter associated with antigen processing; ABC, ATP binding cassette; CFTR, cystic fibrosis transmembrane conductance regulator; ER, endoplasmic reticulum; GAD, glutamic acid decarboxylase; HLA, human leukocyte antigen; MHC, major histocompatibility complex; mAb, monoclonal antibody; MRP1, multidrug resistance protein 1; NBD, nucleotide binding domain; P-gp, P-glycoprotein; wt, wild type; PAGE, polyacrylamide gel electrophoresis.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Higgins, C. F. (1992) Annu. Rev. Cell Biol. 8, 67-113[CrossRef] |
2. | Schneider, E., and Hunke, S. (1998) FEMS Microbiol. Rev. 22, 1-20[CrossRef][Medline] [Order article via Infotrieve] |
3. |
Diederichs, K.,
Diez, J.,
Greller, G.,
Muller, C.,
Breed, J.,
Schnell, C.,
Vonrhein, C.,
Boos, W.,
and Welte, W.
(2000)
EMBO J.
19,
5951-5961 |
4. | Hung, L. W., Wang, I. X., Nikaido, K., Liu, P. Q., Ames, G. F., and Kim, S. H. (1998) Nature 396, 703-707[CrossRef][Medline] [Order article via Infotrieve] |
5. | Walker, J. E., Saraste, M., Runswick, M. J., and Gay, N. J. (1982) EMBO J. 1, 945-951[Medline] [Order article via Infotrieve] |
6. | Senior, A. E., and Gadsby, D. C. (1997) Semin. Cancer Biol. 8, 143-150[CrossRef][Medline] [Order article via Infotrieve] |
7. |
van Veen, H. W.,
Margolles, A.,
Muller, M.,
Higgins, C. F.,
and Konings, W. N.
(2000)
EMBO J.
19,
2503-2514 |
8. | Senior, A. E., al-Shawi, M. K., and Urbatsch, I. L. (1995) FEBS Lett. 377, 285-289[CrossRef][Medline] [Order article via Infotrieve] |
9. |
Gao, M.,
Cui, H. R.,
Loe, D. W.,
Grant, C. E.,
Almquist, K. C.,
Cole, S. P.,
and Deeley, R. G.
(2000)
J. Biol. Chem.
275,
13098-13108 |
10. |
Hou, Y.,
Cui, L.,
Riordan, J. R.,
and Chang, X.
(2000)
J. Biol. Chem.
275,
20280-20287 |
11. |
Nagata, K.,
Nishitani, M.,
Matsuo, M.,
Kioka, N.,
Amachi, T.,
and Ueda, K.
(2000)
J. Biol. Chem.
275,
17626-17630 |
12. | Elliott, T. (1997) Adv. Immunol. 65, 47-109[Medline] [Order article via Infotrieve] |
13. | Spies, T., and Demars, R. (1991) Nature 351, 323-324[CrossRef][Medline] [Order article via Infotrieve] |
14. | Algarra, I., Cabrera, T., and Garrido, F. (2000) Hum. Immunol. 61, 65-73[CrossRef][Medline] [Order article via Infotrieve] |
15. | Tortorella, D., Gewurz, B. E., Furman, M. H., Schust, D. J., and Ploegh, H. L. (2000) Annu. Rev. Immunol. 18, 861-926[CrossRef][Medline] [Order article via Infotrieve] |
16. | Chen, H. L., Gabrilovich, D., Tampe, R., Girgis, K. R., Nadaf, S., and Carbone, D. P. (1996) Nat. Genet. 13, 210-213[Medline] [Order article via Infotrieve] |
17. | Neefjes, J. J., Momburg, F., and Hammerling, G. J. (1993) Science 261, 769-771[Medline] [Order article via Infotrieve] |
18. |
van Endert, P. M.
(1999)
J. Biol. Chem.
274,
14632-14638 |
19. | Knittler, M. R., Alberts, P., Deverson, E. V., and Howard, J. C. (1999) Curr. Biol. 9, 999-1008[CrossRef][Medline] [Order article via Infotrieve] |
20. | Daniel, S., Caillat-Zucman, S., Hammer, J., Bach, J. F., and van Endert, P. M. (1997) J. Immunol. 159, 2350-2357[Abstract] |
21. | van Endert, P. M., Tampé, R., Meyer, T. H., Tisch, R., Bach, J.-F., and McDevitt, H. O. (1994) Immunity 1, 491-500[Medline] [Order article via Infotrieve] |
22. |
Lauvau, G.,
Gubler, B.,
Cohen, H.,
Daniel, S.,
Caillat-Zucman, S.,
and van Endert, P. M.
(1999)
J. Biol. Chem.
274,
31349-31358 |
23. | Meyer, T. H., van Endert, P. M., Uebel, S., Ehring, B., and Tampé, R. (1994) FEBS Lett. 351, 443-447[CrossRef][Medline] [Order article via Infotrieve] |
24. | Bach, J. M., Otto, H., Nepom, G. T., Jung, G., Cohen, H., Timsit, J., Boitard, C., and van Endert, P. M. (1997) J. Autoimmun. 10, 375-386[CrossRef][Medline] [Order article via Infotrieve] |
25. | Anderson, M. P., and Welsh, M. J. (1992) Science 257, 1701-1704[Medline] [Order article via Infotrieve] |
26. |
Szabo, K.,
Welker, E.,
Bakos,
Muller, M.,
Roninson, I.,
Varadi, A.,
and Sarkadi, B.
(1998)
J. Biol. Chem.
273,
10132-10138 |
27. |
Urbatsch, I. L.,
Gimi, K.,
Wilke-Mounts, S.,
and Senior, A. E.
(2000)
J. Biol. Chem.
275,
25031-25038 |
28. |
Daniel, S.,
Brusic, V.,
Caillat-Zucman, S.,
Petrovsky, N.,
Harrison, L.,
Riganelli, D.,
Sinigaglia, F.,
Gallazzi, F.,
Hammer, J.,
and van Endert, P. M.
(1998)
J. Immunol.
161,
617-624 |
29. | Cresswell, P., Bangia, N., Dick, T., and Diedrich, G. (1999) Immunol. Rev. 172, 21-28[Medline] [Order article via Infotrieve] |
30. | Garrido, F., Ruiz-Cabello, F., Cabrera, T., Perez-Villar, J. J., Lopez-Botet, M., Duggan-Keen, M., and Stern, P. L. (1997) Immunol. Today 18, 89-95[CrossRef][Medline] [Order article via Infotrieve] |
31. |
Urbatsch, I. L.,
Sankaran, B.,
Weber, J.,
and Senior, A. E.
(1995)
J. Biol. Chem.
270,
19383-19390 |