From the Department of Microbiology and Immunology
and § Department of Pharmacology, University of Michigan
Medical School, Ann Arbor, Michigan 48109-0620
Received for publication, October 17, 2000, and in revised form, November 15, 2000
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ABSTRACT |
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We generated mutants of the transporter
associated with antigen-processing subunits TAP1 and TAP2 that were
altered at the conserved lysine residue in the Walker A motifs of the
nucleotide binding domains (NBD). In other ATP binding cassette
transporters, mutations of the lysine have been shown to reduce or
abrogate the ATP hydrolysis activity and in some cases impair
nucleotide binding. Mutants TAP1(K544M) and TAP2(K509M) were expressed
in insect cells, and the effects of the mutations on nucleotide
binding, peptide binding, and peptide translocation were assessed. The mutant TAP1 subunit is significantly impaired for nucleotide binding relative to wild type TAP1. The identical mutation in TAP2 does not
significantly impair nucleotide binding relative to wild type TAP2.
Using fluorescence quenching assays to measure the binding of
fluorescent peptides, we show that both mutants, in combination with
their wild type partners, can bind peptides. Since the mutant TAP1 is
significantly impaired for nucleotide binding, these results indicate
that nucleotide binding to TAP1 is not a requirement for peptide
binding to TAP complexes. Peptide translocation is undetectable for
TAP1·TAP2(K509M) complexes, but low levels of translocation
are detectable with TAP1(K544M)·TAP2 complexes. These results suggest
an impairment in nucleotide hydrolysis by TAP complexes containing
either mutant TAP subunit and indicate that the presence of one intact
TAP NBD is insufficient for efficient catalysis of peptide
translocation. Taken together, these results also suggest the
possibility of distinct functions for TAP1 and TAP2 NBD during a single
translocation cycle.
The transporter associated with antigen processing
(TAP)1 is a critical
component of the major histocompatibility complex (MHC) class I antigen
presentation (1-3). TAP functions to translocate peptides from the
cytosol to the ER. Binding of peptides to newly synthesized MHC class I
molecules in the ER stabilizes the MHC class I heterodimer and allows
transit of MHC class I-peptide complexes to the cell surface for immune
surveillance by T cells (4). Two structurally related subunits of the
TAP transporter, TAP1 and TAP2, form a complex on the ER membrane that
is necessary and sufficient for peptide translocation from the cytosol
into the ER. The cytosolic face of TAP1·TAP2 complexes contains a
binding site for peptides (5), which can function in the sequestration of peptides derived from proteasomal proteolysis. A recently discovered protein called tapasin is associated with the TAP1·TAP2 complex (6,
7). Tapasin has been shown to enhance the expression level of TAP1 and
increase peptide transport by TAP complexes (8). However, tapasin is
not required for peptide binding by TAP1·TAP2 complexes or for
translocation per se, since TAP1·TAP2 complexes expressed
heterologously in insect cells can bind and transport peptides (9).
TAP is a member of the ATP-binding cassette (ABC) family of
transmembrane transport proteins (10). Members of this family transport
various substrates across cellular membranes in an
ATP-dependent manner. Known substrates of ABC transporters
include amino acids, peptides, proteins, sugars, and lipids. The
typical ABC transporter has two hydrophobic membrane-spanning regions
with multiple membrane-spanning segments and two cytosolic nucleotide
binding domains (NBD). The NBD contains several conserved sequence
motifs. These include the Walker A and Walker B sequence motifs, which
are characteristic of nucleotide binding folds (11). The
membrane-spanning segments and the NBD can occur in a single
polypeptide as in the mammalian cystic fibrosis transmembrane
conductance regulator and P-glycoprotein, as two separate polypeptides
as in the TAP transporters, or as four separate polypeptides as in many
bacterial ABC transporters (10).
TAP1 and TAP2 each comprise one membrane-spanning region with several
membrane-spanning segments and one cytosolic NBD. In vitro
experiments with intact TAP domains as well as with each truncated NBD
have shown that both TAP1 and TAP2 bind ATP and ADP (12-15). Indirect
evidence for the occurrence of ATP hydrolysis during translocation
comes from observations that the addition of exogenous ATP is required
for peptide translocation by TAP and that translocation is not
supported by nonhydrolyzable ATP analogs or ADP (16). However, the
catalysis of ATP hydrolysis by TAP complexes or either NBD remains to
be directly demonstrated. It is not known whether ATP hydrolysis by
TAP1, TAP2, or the complex is required for a completion of a catalytic
cycle. Furthermore, the requirement for nucleotide binding to each TAP
subunit upon substrate (peptide) interactions with the TAP complex also
remains to be unambiguously defined. Peptide binding to TAP was
initially suggested to be nucleotide binding-independent, based
upon studies of peptide binding to microsomal membranes expressing wild
type TAP1 and TAP2 in the presence or absence of apyrase, an enzyme that depletes ATP and ADP (5). However, based upon more recent studies
using TAP1 and TAP2 mutants that both lacked nucleotide binding
activity, it was suggested that peptide binding by TAP is dependent
upon nucleotide interactions with one or both subunits of the
TAP1·TAP2 complex (17). Further experiments are required to resolve
this discrepancy and to establish whether nucleotide binding to TAP1,
TAP2, or both subunits alters the affinity of TAP1·TAP2 complexes for peptides.
Toward a definition of the requirement for nucleotide binding and
hydrolysis by each TAP subunit for peptide binding and translocation, we generated mutants of TAP1 (K544M) and TAP2 (K509M) that were altered
at a conserved lysine residue of the Walker A motif
(GXXGXGK(S/T)) of each protein. Based upon the
crystal structure of the ATP binding subunit of histidine permease, a
bacterial ABC transporter, the Walker A lysines of TAP1 and TAP2 are
predicted to be located on the phosphate binding loop (P-loop) and form
contacts with the Construction of Baculoviruses Encoding Mutant TAP
Subunits--
The cDNAs for TAP1 and TAP2 were obtained from Dr.
John Trowsdale. Polymerase chain reaction was used to introduce a
sequence encoding a His6 tag to the 3'-end of TAP1 for the
TAP1-His construct, to introduce BamHI sites at both ends of the TAP1
cDNA, and to introduce BglII sites at both ends of the
TAP2 cDNA. The modified cDNAs were subcloned into pPCR2.1.
Site-directed mutagenesis was performed by the Kunkel method (24). The
mutated cDNAs were sequenced, excised from pPCR 2.1, and ligated
into pAcUW51 transfer vector. The transfer vectors and BaculoGold DNA
(PharMingen) were cotransfected into insect cells as described in Ref.
43. Plaque assays were used to isolate pure viruses, which were used to
reinfect cells for virus amplification.
Insect Cell Infection, Lysis, Immunoprecipitation, and Binding to
ATP Beads--
2 × 107 Sf21 cells were
infected with the appropriate baculovirus at a multiplicity of
infection of 5-20, depending on the expression level of the individual
virus. After ~72 h, the cells were harvested and lysed in 1 ml of
lysis buffer (10 mM Tris, 10 mM phosphate, 130 mM NaCl, 1% Triton X-100, pH 7.5) supplemented with
protease inhibitors (Complete mini-EDTA free tablets; Roche Molecular
Biochemicals). The cells were lysed on ice for 45 min, and the lysates
were cleared by centrifugation at 40,000 × g for 45 min at 4 °C. 0.5 ml of the lysate was then immunoprecipitated with
0.75 µl of anti-His ascites, 200 µl of anti-TAP1 (148.3) hybridoma
supernatant (9), or 200 µl of anti-TAP2 (435.3) (25) hybridoma
supernatant. The samples were centrifuged again at 14,000 × g for 5 min at 4 °C, and the supernatants were
transferred to an appropriate volume of protein A-Sepharose beads
prewashed in wash buffer (10 mM Tris, 10 mM
phosphate, 130 mM NaCl, 0.5% Triton X-100, pH 7.5) and
incubated for 1 h at 4 °C with shaking. The beads were then
washed three times in wash buffer before adding SDS-PAGE sample buffer.
Samples were separated on 8% SDS-PAGE gels, transferred to
nitrocellulose membranes, and immunoblotted with 5 µl of anti-His
ascites, 150 µl of anti-TAP1 hybridoma supernatant, or 250 µl of
anti-TAP2 hybridoma supernatant overnight. The membranes were then
incubated with 5 µl of goat anti-mouse alkaline
phosphatase-conjugated antibody (American Qualex) for 2 h and
developed with the Bio-Rad alkaline phosphatase conjugate substrate kit.
Cells were infected and lysed in an identical manner for the ATP beads
binding experiment. Cells were lysed using the lysis buffer described
above, containing 5 mM MgCl2. 0.5 ml of each cleared lysate was incubated for 2 h with an appropriate amount of
preswollen ATP-, ADP-, or AMP-conjugated agarose beads (Sigma) or
immunoprecipitated with antibodies as described above. The beads were
then washed three times with lysis buffer, run on gels, transferred to
membranes, and immunoblotted in the same manner as above.
Microsome Preparations--
Microsomes were prepared as
described previously (9). Briefly, 1 × 108 SF21 cells
were infected with the appropriate baculovirus at an multiplicity of
infection of 5-20, depending on the expression level of the individual
virus. After 72 h, the cells were harvested, washed once in
ice-cold PBS, and then resuspended in 800 µl of ice-cold cavitation
buffer (250 mM sucrose, 25 mM potassium
acetate, 5 mM magnesium acetate, 0.5 mM calcium
acetate, 50 mM Tris, pH 7.4) supplemented with a protease
inhibitor mixture. Cells were lysed by repeatedly drawing the
suspension through a 26-gauge needle. After centrifugation at
~500 × g for 5 min, the supernatant was added to 5 ml of 2.5 M sucrose in gradient buffer (2.5 M
sucrose, 150 mM potassium acetate, 5 mM
magnesium acetate, 50 mM Tris, pH 7.4). This was then
overlaid with 3 ml each of 2.0 M sucrose in gradient
buffer, then 1.3 M sucrose in gradient buffer, and finally
with 800 µl of cavitation buffer. This was centrifuged overnight at
80,000 × g, and the microsomal fraction at the
interface of the 2.0 and 1.3 M sucrose layers was
collected. This fraction was diluted into 5 ml of PBS, 1 mM
dithiothreitol, and centrifuged for 1 h at 100,000 × g, and the pellet was resuspended in 1 ml of PBS, 1 mM dithiothreitol and frozen in aliquots at Peptide Translocation Assay--
For the translocation assays, a
peptide of the sequence RRYNASTEL was synthesized, and iodinated using
the chloramine-T labeling chemistry to a specific activity of 100-150
µCi/µg. Microsome-based translocation assays were carried out as
previously described (9). 30 µg of insect cell microsomes containing
wild type or mutant TAP1·TAP2 complexes or single subunits were added
to 150 µl of assay buffer (PBS, 0.1% bovine serum albumin, 1 mM dithiothreitol, pH 7.3) containing 10 mM
MgCl2 and 5 mM ATP (+ATP samples) or 0.03 units/µl of apyrase ( Peptide Binding Assay--
Peptide binding assays were carried
out using a recently described procedure (27). All peptides were
synthesized using Fmoc (N-(9-fluorenyl)methoxycarbonyl)
chemistry. For the binding experiments, a peptide was synthesized with
the sequence RRYQKCTEL. For fluorochrome labeling, the peptide was
dissolved in PBS, 33% N,N-dimethylformamide. 5-Iodoacetamidofluorescein (Molecular Probes, Inc., Eugene, OR) was
then added at a 1.2:1 molar ratio to peptide. The reaction was allowed
to incubate in the dark for 2 h at room temperature. The modified
peptide was purified by reverse-phase HPLC. Peptide concentrations were
normalized between experiments by measuring the absorbance at 495 nm
(A495 = 0.28 for 0.01 mg/ml peptide).
For the peptide binding assays, the fluorescence emission signal
(
After stabilization of fluorescence signals, unlabeled peptide was
added at a concentration of ~30 µM, and the
fluorescence recovery was monitored over several seconds as bound
fluorescent peptide dissociated. The dissociation rate constants,
kd, were estimated by fitting the fluorescence
recovery signal to a monoexponential function, y = Fe +
Each set of binding experiments included a control with microsomal
membranes prepared from uninfected Sf21 cells. Fluorescence quenching signals were also sometimes observed with microsomes derived
from uninfected cells, but recovery was typically not observed (Figs. 4
and 5). At peptide concentrations below 80 nM, the
quenching signals observed for the control microsome preparations (if
any) were typically significantly lower than those observed with
microsome preparations containing wild type or mutant TAP complexes
(Fig. 5, A-C). Variations in the quenching signal observed for microsome preparations from uninfected cells (ranging from negative
amplitudes to low positive amplitudes as shown in Fig. 5,
A-C) might arise due to peptide degradation during storage. In general, the lowest background signals were observed for experiments conducted within 1 week of HPLC purification of peptide, when the
purified peptide was stored at Expression and ATP Binding by TAP1 and TAP2 Mutants--
We
modified the TAP1 cDNA by polymerase chain reaction, to introduce a
sequence encoding a C-terminal hexahistidine tag (TAP1-His), to facilitate the biochemical analyses of TAP1 expression and function.
The TAP1-His sequence was further modified by site-directed mutagenesis (24), in order to alter a conserved lysine residue in the
Walker A motif (Lys544) to a methionine. A similar
modification was introduced into the cDNA sequence encoding TAP2,
in order to alter the Walker A lysine (Lys509) to a
methionine. Mutant clones were sequenced and ligated into the
baculovirus transfer vector pAcUW51. Recombinant baculoviruses were
generated encoding histidine-tagged TAP1 (TAP1-His), the corresponding
Walker A lysine mutant (TAP1(K544M)-His), and the TAP2 Walker A lysine
mutant (TAP2(K509M)). The anti-His antibody (Covance Scientific) was
used to screen for TAP1 expression, and the TAP2-specific antibody
435.3 (anti-TAP2) (25) was used to screen for TAP2 expression.
Baculoviruses encoding wild type TAP1 (TAP1) and TAP2 (TAP2) were
obtained from the laboratory of Dr. Robert Tampé (9). For
comparisons of nucleotide binding by each mutant or wild type TAP
subunit, insect cells were infected with baculoviruses encoding
TAP1-His, TAP1(K544M)-His, TAP2, or TAP2(K509M) for ~72 h. Detergent
lysates of cells from each infection were incubated with ATP, ADP, or
AMP-Sepharose beads for 2 h. Subsequently, the beads were washed,
and proteins were eluted from the beads by boiling in the presence of
SDS-polyacrylamide gel electrophoresis buffer. Proteins associated with
the beads were analyzed by SDS-polyacrylamide gel electrophoresis and
immunoblotting with anti-His or anti-TAP2 antibodies (Fig.
1). Lysates were also immunoprecipitated
with anti-His or anti-TAP2 antibodies to directly visualize the
relative expression levels of mutant versus wild type
proteins. At comparable expression levels of TAP1-His and TAP1(K544M)-His (Fig. 1A, lanes 4 and
8), strong binding of TAP1-His to ATP-Sepharose and
ADP-Sepharose beads is visualized (Fig. 1A, lanes
1 and 2), whereas TAP1(K544M)-His binding to ATP
and ADP beads is barely detectable (Fig. 1A,
lanes 5 and 6). These observations indicate that the K544M mutation impairs nucleotide binding to TAP1. By
contrast, TAP2(K509M) binding to ATP and ADP beads does not appear to
be significantly impaired relative to wild type TAP2 (Fig.
1B, lanes 1 and 2 compared
with lanes 5 and 6).
Subunit Association and Peptide Translocation by Wild Type and
Mutant TAP Complexes--
It has previously been shown TAP1 and TAP2
associate into stable complexes and that both subunits are required for
peptide binding and translocation (28, 29). Prior to analysis of the peptide binding and translocation properties of the mutants, it was
therefore necessary to demonstrate that each mutant could form
TAP1·TAP2 complexes. To assay the formation of TAP1(K544M)-His·TAP2 complexes, cells were infected with viruses encoding TAP1(K544M)-His and TAP2, or with TAP1-His and TAP2 as the positive controls. Detergent
lysates were immunoprecipitated with either anti-His or anti-TAP2
antibodies. The samples were subsequently separated by
SDS-polyacrylamide gel electrophoresis and transferred to
nitrocellulose membranes, and the membranes were stained with anti-His
or anti-TAP2. We observed that TAP1(K544M)-His associates with TAP2 as
does TAP1-His (Fig. 2A).
Likewise, TAP2(K509M) associates with TAP1 as does wild type TAP2, as
measured by coimmunoprecipitation analyses with the TAP1-specific
antibody 148.3 (anti-TAP1) (9) and anti-TAP2 (Fig. 2B).
Peptide translocation experiments were carried out as previously
described (9, 26). The model substrate RRYNASTEL was used, which was
125I-labeled. Microsomal membrane preparations containing
wild type or mutant TAP complexes were incubated with the radiolabeled
peptide in the presence or absence of ATP for 15 min at 37 °C. Since
the peptide sequence contains an NAS glycosylation motif, translocated peptide was retained in the ER rather than being exported (30). Free
peptide was separated by centrifugation, the microsomal membranes were
lysed in detergent, and glycosylated peptide was quantified by
determining their binding to ConA-Sepharose beads. The experiments were
carried out with microsome preparations expressing wild type and mutant
TAP1·TAP2 complexes. Microsome preparations expressing single TAP
subunits were used as negative controls, since it has previously been
shown that microsomal preparations expressing wild type TAP1 alone or
wild type TAP2 alone do not translocate peptides (9, 25). The results
of a representative experiment are indicated in Fig.
3. Within an experiment, we defined a
translocation signal as positive when the average
cpm+ATP/cpm
For mutant TAP1(K544M)-His·TAP2 complexes, a low but positive
translocation signal was observed in three independent translocation experiments. In each experiment, the average
cpm+ATP/cpm Peptide Binding by Wild Type and Mutant TAP Complexes--
A
fluorescence quenching assay has recently been described for
quantitation of peptide binding to TAP complexes, under
nucleotide-depleting conditions (27). We used these assays to quantify
peptide binding by wild type and mutant TAP complexes. For this
purpose, the peptide RRYQKCTEL was fluorescein-labeled using
5-iodoacetamidofluorescein and HPLC-purified. Microsomal membrane
preparations containing each mutant TAP complex, the corresponding wild
type TAP complexes, or neither TAP subunit (uninfected microsomes) were
added to different concentrations of the fluorescent peptide, and the
fluorescence emission signal was monitored. For the wild type as well
as for the mutant TAP complexes, increased fluorescence quenching was observed as the peptide concentration was increased (Fig.
4, A-C). After stabilization
of fluorescence signals, unlabeled peptide was added at a concentration
of ~30 µM, and the fluorescence recovery was monitored
over several seconds as bound fluorescent peptide dissociated. As for
wild type TAP complexes, fluorescence recovery was also observed for
both TAP mutants, and the magnitude of the recovery was in proportion
to the magnitude of the quenching. The magnitude of the fluorescence
quenching and recovery signals obtained for wild type and mutant TAP
complexes were critically dependent upon the expression levels of both
TAP subunits. In the experiments shown in Fig. 4, A and
C, the same microsome preparations of TAP1·TAP2 and
TAP1·TAP2(K509M) were used as for the translocation assays shown in
Fig. 3 (Fig. 3B, lanes 1 and
2). At lower expression levels, reliable fluorescence
quenching and recovery was not observed. For example, with microsomal
preparations of TAP1-His·TAP2 (Fig. 3B, lane
3), fluorescence recovery was not observed upon the addition of excess unlabeled peptide. Increasing the expression level of both
TAP1-His and TAP2, however, resulted in binding profiles that resembled
wild type TAP1·TAP2 complexes (data not shown).
To verify the TAP dependence of the fluorescence quenching and recovery
signals shown in Fig. 4, A-C, changes in fluorescence were
also monitored upon the addition of microsomal preparations from
uninfected insect cells (cells that expressed neither TAP subunit) to
different concentrations of the fluorescent peptide (Fig.
4D). The amplitudes of fluorescence quenching signals
observed for uninfected microsomes typically ranged from negative
values to low positive values over the concentration range of 2.5-80 nM in several different experiments. Positive quenching
profiles, if observed, were linear rather than exponential, and the end point amplitudes were significantly lower than observed for
TAP-containing microsomes. Finally, in cases where positive quenching
signals were observed with uninfected microsomes, fluorescence recovery was typically not observed upon the addition of excess unlabeled peptide (an example is indicated in Fig. 4D, 40 nM). Thus, the fluorescence quenching and recovery profiles
for uninfected microsomes are distinct from those seen for wild type
TAP complexes, as previously described (27), or for either mutant
complex, as indicated in Fig. 4, B and C.
The steady state fluorescence quenching was determined from an
exponential fit of the association data (averaged over two independent
experiments for the same microsome preparation). The steady state
fluorescence quenching values are plotted as a function of peptide
concentration for microsomes containing wild type and mutant TAP
complexes (Figs. 5, A-C).
Each experiment summarized in Fig. 5, A-C, included a
negative control with uninfected microsomes (also averaged over two
independent experiments for the same microsome preparation). For most
of the data points shown in Fig. 5, the amplitudes of the fluorescence
quenching signals obtained with microsomes containing wild type or
mutant TAP complexes were significantly greater than that obtained for
microsomes from uninfected cells, over the indicated concentration
range. Taken together with the fluorescence recovery data, we infer
from these results that both sets of mutant TAP complexes are capable
of binding peptides. The apparent binding constants
(Kd) were calculated from the steady state
fluorescence quenching versus peptide concentration plots
(Fig. 5, A-C). In addition, the dissociation rate
constants, kd, were estimated by fitting the
fluorescence recovery signal to a monoexponential function (Fig. 4,
A-C). The calculated binding constants for the
TAP1·TAP2(K509M) mutant indicated that the peptide binding affinity
of this mutant was slightly weaker compared with wild type TAP
complexes. The observations that this mutant TAP complex can bind
peptides (Figs. 4C and 5C) and nucleotides (Fig.
1B) but does not translocate peptides (Fig. 3A)
indicate that the mutant is arrested at a stage that follows peptide
binding and nucleotide binding, most likely at a step that requires
nucleotide hydrolysis.
The calculated binding and rate constants for peptide interactions with
TAP1·TAP2 complexes and TAP1-His·TAP2 are very similar to each
other and to TAP1(K544M)-His·TAP2 complexes (Fig. 4, A and
B, and Fig. 5, A and B; data not shown
for TAP-His·TAP2 complexes). As mentioned above, the peptide binding
experiments shown in Figs. 4 and 5 were carried out in the presence of
apyrase (ATP and ADP-depleting conditions). It is a formal possibility
that TAP-associated nucleotides cannot be accessed by apyrase and that
one or both TAP subunits in wild type TAP complexes remain
nucleotide-occupied in the presence of apyrase. However, since
TAP1(K544M)-His is significantly impaired for nucleotide binding (Fig.
1), we expect that in the presence of apyrase, the nucleotide occupancy
will be low for the TAP1(K544M)-His mutant in TAP1(K544M)-His·TAP2
complexes. The observation that the calculated binding constants and
dissociation rate constants for TAP1(K544M)-His·complexes do not
differ significantly from the values derived for wild type TAP1·TAP2
complexes when apyrase is present or when ADP is present (Table
I) indicates that nucleotide interaction
with TAP1 is not a requirement for peptide interactions with
TAP1·TAP2 complexes.
It has previously been observed that the isolated nucleotide
binding domains of TAP1 and TAP2 are capable of binding nucleotide (9,
13, 15). In addition, it has been shown that wild type TAP1·TAP2
complexes bind to Here we report that the K509M mutation in TAP2 abrogates peptide
transport by TAP1·TAP2(K509M) complexes, although ATP binding by this
mutant is not significantly different from wild type. Impairment in
peptide translocation does not arise from structural disruptions
induced by the mutation, since TAP1·TAP2(K509M) complexes are capable
of binding both peptides and nucleotides (Figs. 1 and 3). Furthermore,
based upon limited proteolytic digestion analysis, the proteolysis
profiles observed for TAP1·TAP2(K509M) complexes closely parallel the
profiles seen for TAP1·TAP2
complexes.2 Thus, nucleotide
hydrolysis by TAP complexes containing mutant TAP2 appears to be
impaired. Likewise, the TAP1(K544M) mutation reduces peptide
translocation efficiency of TAP complexes even when no residue
modifications are introduced into the TAP2 subunit. Thus, nucleotide
hydrolysis by TAP complexes containing mutant TAP1 also appears to be
impaired. We are presently unable to directly demonstrate impaired
ATPase activity of TAP complexes containing either mutant TAP subunit,
since we do not have a working assay to measure the ATPase activities
of TAP1 or TAP2 NBD or of TAP1·TAP2 complexes. ATPase activities have
been previously demonstrated for purified NBD of other ABC transporters
(for example, Refs. 31-33). Using purified TAP1 and TAP2 NBD and TAP
NBD complexes (15), we were unable to unambiguously demonstrate ATPase
activities. The lack of measurable ATPase activity for TAP NBD
suggested that intact membrane-associated TAP complexes might be
required for observation of the enzymatic activity. Expression of the
human MDR1 gene in Sf9 cells has previously
been shown to generate a membrane ATPase activity that is significantly
stimulated by drugs known to interact with P-glycoprotein (34). We are
attempting to develop an analogous assay using microsomal membrane
preparations expressing wild type TAP1·TAP2 complexes but thus far
have been unable to demonstrate a TAP-specific ATPase activity.
The observation that the TAP2(K509M) mutation impairs translocation by
TAP1·TAP2(K509M) complexes although no residue alterations were
introduced into TAP1 indicates that the ATPase activity at TAP1, if
present, is insufficient for completion of a peptide translocation
cycle. Taken together with the observation that the TAP1(K544M)
mutation significantly reduces peptide translocation efficiency of TAP
complexes when no residue modifications are introduced into TAP2, these
results indicate a coupling between nucleotide interactions with TAP1
and TAP2. The mechanistic model that emerges from these studies differs
from that recently described for histidine permease. Functional
analyses of mutants of histidine permease, which contains two copies of
a single NBD (HisP), have shown that presence of one intact ATP
hydrolysis site is sufficient to support ATPase activity and substrate
translocation (35). In chimeric histidine permease complexes containing
one ATPase-active and one ATPase-inactive NBD, ligand translocation
occurred at half the rate of the wild type. These observations are
consistent with a mechanism for histidine permease, whereby, in wild
type complexes, two molecules of ATP are hydrolyzed within a single turnover of the catalytic cycle, and hydrolysis at either NBD can
result in ligand translocation. By contrast to histidine permease, in
the case of the P-glycoprotein-based drug transport system, mutations
of the Walker A lysine at either one NBD completely blocked
drug-stimulated ATPase activity of P-glycoprotein and drug transport,
although no residue alterations were introduced at the second site (19,
21). Other mutants, as well as single site chemical modifications of
P-glycoprotein were observed to show similar effects (36-38). Similar
results were obtained for maltose permease (39), another bacterial
transporter, which contains two copies of a single NBD (MalK). When the
Walker A lysine was replaced with asparagine in both MalK subunits,
maltose transport and ATPase activities were reduced to 1% of those of the wild type. When the mutation was present in only one of the two
subunits, the complex had 6% of the wild-type activities. Functional
coupling of ATPase activities observed for P-glycoprotein and MalK
might arise if ATP hydrolysis is required at two distinct steps during
a single turnover of the catalytic cycle as has been recently suggested
for P-glycoprotein (40, 41). Activation of the ATPase activity at each
NBD might require transient conformational states that are acquired at
specific steps of a translocation cycle. Modification at either ATPase
site would then be expected to cause conformational trapping, resulting
in an inability to complete a translocation cycle. Other models
invoking alternating catalytic cycles (42) at each NBD are also
consistent with the observations of functional coupling between the two
NBD.
Our studies indicate that TAP may belong to a group of transporters
with mechanistic similarities to P-glycoprotein and MalK. It is
interesting that the TAP2(K509M) mutation abrogates peptide translocation by TAP1·TAP2(K509M) complexes but that the TAP1 mutant
with a significant impairment in TAP1 nucleotide binding appears to,
with low efficiency, mediate peptide translocation by
TAP1(K544M)-His·TAP2 complexes. One interpretation of these observations is that nucleotide hydrolysis by TAP2 constitutes the
first ATP hydrolysis step during a single peptide translocation cycle.
This first hydrolytic event might be accompanied by the release of
bound peptide and its translocation. Nucleotide hydrolysis by TAP1
might then be required to reset the transporter for another cycle of
peptide translocation. Additional experiments are required to obtain
further evidence in support of such a mechanism and to obtain greater
insights into the precise role of TAP1 and TAP2 during translocation.
We analyzed the ability of each mutant TAP subunit in combination with
the wild type partner to bind peptides. Using fluorescence quenching
assays, the equilibrium binding constant we calculate for peptide
interaction with wild type TAP1·TAP2 complexes (Kd = 19.4 ± 4.8 nM) is quite similar to that previously
reported (Kd = 12 ± 1 nM) (27).
However, the calculated dissociation rate constant
(kd = 0.013 ± 0.005 s The finding that the K544M mutation in TAP1 significantly reduces
nucleotide binding also allowed us to explore the linkage between
nucleotide binding by TAP1 and peptide binding to TAP1·TAP2 complexes. Using assays with fluorescently labeled peptides, we studied
peptide binding to wild type TAP1·TAP2 complexes or
TAP1(K544M)-His·TAP2 complexes in the presence of apyrase
(nucleotide-depleting conditions) or in the presence of ADP. For the
latter experiments, microsomes were first exposed to 1 mM
ADP and subsequently diluted 10-fold into buffers containing
fluorescent peptide. Under these conditions, wild type TAP1 is expected
to be nucleotide-bound in TAP1·TAP2 complexes, based upon previous
studies of inhibition of 8-azido-ATP labeling of TAP1 in the presence
of 100 µM to 1 mM cold ADP (14). For
TAP1(K544M)-His·TAP2 complexes that were treated with apyrase, the
TAP1(K544M)-His is expected to be significantly nucleotide-depleted, based upon the result that nucleotide binding to the TAP1(K544M)-His mutant is significantly impaired, and that apyrase might further facilitate nucleotide depletion. Since the apparent
Kd and kd values for wild type
TAP1·TAP2 and mutant TAP1(K544M)-His·TAP2 complexes in the presence
of ADP and apyrase, respectively, are very similar, there does not
appear to be a strong correlation between nucleotide binding to TAP1
and peptide binding. Our interpretation of this result is that a
nucleotide-depleted TAP1 is functional for peptide binding. For the
peptide binding experiments described here, we were careful to maintain
the mutant TAP proteins at levels equal to or greater than that present
for wild type TAP1·TAP2 complexes, at which unambiguous fluorescence
quenching and recovery signals were obtained. At low expression levels
of either TAP1 or TAP2, we found that a peptide binding signal could
not be unambiguously discerned even for wild type TAP1·TAP2
complexes. These observations raise the question of whether expression
levels of TAP1 or TAP2 in other systems could influence the results of
binding experiments, which indicated that nucleotide binding is
required for peptide interactions with the TAP complex (17).
It has previously been reported that the presence of ATP and
nonhydrolyzable ATP analogs enhances the dissociation of peptides from
TAP1·TAP2 complexes (25). At room temperature, we find no significant
differences in the calculated dissociation rates for
peptide-TAP1·TAP2 complexes in the presence of ADP and apyrase (Table
I). We also analyzed peptide binding to wild type TAP1·TAP2 complexes
in the presence of ATP and observed that the ATP does not alter the
Kd or Kd values, relative to
those observed in the presence of ADP and apyrase (data not shown). This result was somewhat surprising, since ATP induces peptide translocation, which entails a prior release of TAP-associated peptides. In this context, it is noteworthy that the binding
experiments were carried out at room temperature, whereas the
translocation experiments require a 37 °C incubation. It is possible
that ATP hydrolysis is significantly inhibited at the lower
temperature. The nucleotide insensitivity of peptide interactions with
TAP1·TAP2 complexes at room temperature point to a mechanism in which
ATP hydrolysis rather than binding per se might be
associated with the structural changes that accompany the cycles of
peptide binding and release. Further experiments are required to
establish the occurrence of ATP-induced structural changes in TAP
complexes at 37 °C and their effect on peptide binding.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-phosphate of ATP (18). Mutations of this lysine
in other ABC transporters have been shown to impair nucleotide binding and/or hydrolysis and impair substrate translocation (19-23). We observe that the Walker A lysine mutations in TAP1 and TAP2 have distinct effects upon nucleotide binding to each subunit, with nucleotide binding being significantly impaired in the TAP1(K544M) mutant but not in the TAP2(K509M) mutant. The observation that the
TAP1(K544M) mutation is significantly impaired for nucleotide binding
by TAP1 also allowed us to determine the correlation between nucleotide
binding by TAP1 and peptide binding to TAP1·TAP2 complexes. Additionally, assays of peptide translocation by each mutant allowed insights into the requirement for nucleotide hydrolysis by TAP2 during
a translocation cycle. Taken together, these results allow for the
refinement of a model for the mechanistic steps involved during a
single peptide translocation cycle.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
70 °C. The
total protein content was determined by a BCA assay (Pierce).
ATP) (Sigma). Radioiodinated RRYNASTEL peptide
was then added, and the samples were incubated at 37 °C for 15 min.
The samples were then centrifuged at 4 °C at 8,800 × g and washed once with 250 µl of the appropriate assay
buffer. The pellets were then resuspended in 250 µl of lysis buffer
(50 mM Tris, 150 mM NaCl, 1% Nonidet P-40, pH
7.4) and incubated on ice for 1 h. After centrifugation at
8,800 × g for 5 min at 4 °C, the supernatants were
transferred to ConA-Sepharose (Amersham Pharmacia Biotech) beads and
incubated for 2 h at 4 °C with agitation. Beads were washed
twice with the appropriate assay buffer, and the radioactivity was
determined using a Beckman 5500
-counter. Results from
microsome-based translocation assays were verified using Sf21
cell-based assays, as per previously described protocols (26).
ex/em = 470/515 nm) was recorded using a PTI
fluorimeter equipped with a microstirring device. A 5-mm round quartz
cuvette was filled with 180 µl of assay buffer (PBS, 1 mM
dithiothreitol, 5 mM MgCl2, pH 7.4) at room
temperature, and the fluorescent peptide RRYQKCTEL was added with
vigorous stirring (2.5-80 nM final concentration). The
fluorescence signal was then allowed to stabilize. 20 µl of insect
cell microsomes treated either with 0.03 units/µl of apyrase or with
1 mM ADP or ATP (for at least 30 min on ice) were added to
the cuvette. The decrease in fluorescence due to
TAP-dependent quenching of peptide fluorescence was
monitored until equilibrium was achieved (280 s). The time dependence
of quenching was determined by fitting the data to an exponential
function, y = Ff +
F*e
kt (where
Ff is the fluorescence value at steady state,
F is the net change in fluorescence, and k is
the rate constant) using nonlinear least squares analysis in the Prism
software package (Graph Pad software). The total fluorescence quenching
signal (
F) was plotted as a function of peptide
concentration. The curve was fitted to the equation y =
F * [P]/(Kd + [P]), and Kd values were determined by nonlinear least squares analysis.
F *
e
kdt (where Fe is
the end point fluorescence value,
F is the net change in
fluorescence, and kd is the dissociation rate constant).
20 °C under dessication.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Binding of TAP1-His, TAP1(K544M)-His, TAP2,
and TAP2(K509M) to nucleotide-agarose beads. Insect cells were
infected with the indicated baculoviruses for ~72 h. The binding of
each protein to ATP-, ADP-, or AMP-agarose beads was assessed, and the
total expression levels of wild type and mutant TAP1 and TAP2 were
assessed using immunoprecipitations (IP) and immunoblotting
with anti-His and anti-TAP2 antibodies, respectively. A,
lanes 1 and 2 show that TAP1-His binds
to ATP and ADP. Lanes 5 and 6 show
that TAP1(K544M)-His is severely impaired in its ability to bind ATP
and ADP relative to wild type TAP1-His. B, lanes
1 and 2 show that TAP2 binds to ATP and ADP.
Lanes 5 and 6 show that the binding
pattern for TAP2(K509M) is similar to wild type TAP2 and indicate that
the mutant is not deficient in nucleotide binding.
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Fig. 2.
Association of wild type and mutant TAP1 and
TAP2. Insect cells were infected with baculoviruses encoding the
indicated constructs. Detergent lysates of these cells were
immunoprecipitated (IP) with the indicated monoclonal
antibody, and immunoblotting analysis was carried out with an antibody
directed against the same or a partner subunit. A,
TAP1-His/TAP2 or TAP1(K544M)-His/TAP2 interactions.
Immunoprecipitations with anti-His indicate the presence of TAP2, and
immunoprecipitations with anti-TAP2 indicate the presence of TAP1.
B, TAP1·TAP2, or TAP1·TAP2(K509M) interactions.
Immunoprecipitations and Western blots were done identically to those
shown in Fig. 2A, but an anti-TAP1 monoclonal antibody (9)
was used instead of anti-His to detect the untagged wild type TAP1
protein. Immunoprecipitations with anti-TAP1 indicate the presence of
TAP2, and immunoprecipitations with anti-TAP2 indicate the presence of
TAP1.
ATP ratio was at least
2-fold above the ratio observed for the single subunit control. By this
criterion, results from three independent translocation experiments
indicated that TAP1·TAP2(K509M) complexes were impaired for
translocation. Impaired translocation by TAP1·TAP2(K509M) complexes
was not due to reduced expression of either TAP1 or TAP2 (Fig.
3B). Indeed, microsomes containing TAP1-His·TAP2 complexes yielded higher cpm+ATP/cpm
ATP
ratios, although the expression levels of TAP1 and TAP2 were
significantly lower than that present in microsomal preparations of
TAP1·TAP2(K509M) complexes (Fig. 3B, compare
lane 2 with lane 3).
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Fig. 3.
Peptide translocation by wild type or mutant
TAP complexes. A, microsomal vesicles expressing the
indicated TAP complexes were assayed for their ability to import
125I-labeled RRYNASTEL. Bar graphs
represent the average of experiments done in triplicate, and the
average cpm+ATP/cpm ATP ratio is
indicated. In three independent translocation experiments, the average
cpm+ATP/cpm
ATP ratios for
TAP1(K544M)-His·TAP2 complexes were 2-fold higher than for single
subunit controls, whereas the average
cpm+ATP/cpm
ATP ratios for
TAP1·TAP2(K509M) complexes were at the same level as the single
subunit controls. B, expression levels of TAP1 and TAP2 in
the translocation experiments from A. Microsomal membranes
used in the translocation assays shown in A were run on
SDS-polyacrylamide gels and immunoblotted with anti-TAP1 or anti-TAP2
monoclonal antibodies. The resulting blot shows that while no
translocation signal is observable for TAP1·TAP2(K509M) complexes,
both TAP subunits are expressed at levels comparable with the wild type
complex (compare lanes 1 and 2). The
TAP1(K544M)-His·TAP2 complex shows a positive translocation signal,
but the translocation efficiency is low. Despite a significantly lower
expression level of TAP1-His and TAP2 in the TAP1-His·TAP2 microsome
preparation, the translocation efficiency is higher than that observed
with the TAP1(K544M)-His·TAP2 preparation (compare lanes
3 and 4).
ATP ratio for the
mutant complex was approximately 2-fold above the single subunit
background (ratios for the single subunit controls ranged from 0.6 to
1.7 in the different experiments). The low translocation efficiency is
apparent from the higher
cpm+ATP/cpm
ATP ratio observed for
wild type complexes although the expression level was significantly
reduced compared with the mutant (Fig. 3B, compare
lanes 3 and 4).
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Fig. 4.
Association and dissociation of fluorescent
peptide·TAP complexes. Microsomal vesicles expressing the
indicated TAP complexes or microsomes prepared from uninfected
Sf21 cells were assayed for their ability to bind
fluorescein-labeled RRYQKCTEL at the indicated concentrations.
Microsomes containing wild type TAP complexes or either mutant showed
increased fluorescence quenching and recovery profiles, as the peptide
concentration was increased over the indicated range. Fluorescence
quenching was sometimes also observed with uninfected microsome
preparations, as indicated, but fluorescence recovery was typically not
observed upon the addition of excess unlabeled peptide. In addition, if
quenching was observed, the magnitude of the quenching was lower than
that seen for TAP-containing microsomes, and the quenching profiles
were linear rather than exponential. The quenching and recovery
profiles shown indicate that both mutant TAP complexes are capable of
binding peptides. The dissociation rate constants,
kd, estimated by fitting the fluorescence recovery
signals to a monoexponential function are indicated for wild type and
mutant TAP complexes. For wild type TAP1·TAP2 and
TAP1·TAP2(K509M), the same microsomes were used as in the
translocation assays indicated in Fig. 3. For TAP1(K544M)-His·TAP2
complexes, a different microsome preparation was used that clearly
demonstrated quenching and recovery signals for this mutant. The total
protein concentration on each microsome preparation was 0.7 mg/ml
TAP1·TAP2 (A), 1.8 mg/ml TAP1(K544M)-His·TAP2 (B), 1.3 mg/ml TAP1·TAP2(K509M) (C), and 1.3 mg/ml uninfected
(D).
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Fig. 5.
Equilibrium binding curves for fluorescent
peptide/TAP interactions. Microsomal vesicles expressing the
indicated TAP complexes or microsomes prepared from uninfected
Sf21 cells were assayed for their ability to bind
fluorescein-labeled RRYQKCTEL over the indicated concentration range.
At each peptide concentration, the steady state fluorescence quenching
was determined and plotted as a function of peptide concentration, and
the Kd values were estimated. The averages of data
points from two separate experiments with the same microsome
preparation are indicated on each plot. Each set of binding experiments
included a control with microsomal membranes prepared from uninfected
cells, which are also indicated on each plot. For all of the
TAP-containing microsomes indicated, the same microsome preparations
were used as in the translocation assays indicated in Fig. 3. The total
protein concentration on each microsome preparation was 0.7 mg/ml
TAP1·TAP2 (A), 1.1 mg/ml TAP1(K544M)-His·TAP2
(B), and 1.3 mg/ml TAP1·TAP2(K509M) (C). For
the uninfected microsomes, the total protein concentration was 1.3 mg/ml. Four independent sets of binding experiments comparing peptide
interactions with TAP1(K544M)-His·TAP2 microsomes and uninfected
microsomes verify that these mutant complexes are capable of binding
peptides by the following criteria: (i) the overall fluorescence
quenching and recovery profiles resemble those obtained with wild type
TAP complexes rather than uninfected microsomes, and (ii) the
amplitudes of the quenching signals are greater than those obtained for
uninfected microsomes. By similar criteria, four separate sets of
experiments comparing peptide binding by TAP1·TAP2(K509M) microsomes
and uninfected microsomes verify that this mutant complex is also
capable of binding peptides.
Comparisons of peptide binding to wild type TAP1·TAP2 or
TAP1(K544M)-His·TAP2 complexes in the presence of ADP and apyrase
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-32P-labeled 8-azido-ATP and to ATP
and ADP-Sepharose beads (14, 17). Here we show that detergent extracts
of isolated TAP subunits are also capable of binding nucleotide. We
generated mutants of TAP1 and TAP2 that were altered at structurally
analogous residues in the Walker A motif of TAP1. We observed that the
TAP1(K544M) mutation significantly reduced nucleotide binding by TAP1
but that the TAP2(K509M) mutation did not significantly alter
nucleotide binding by TAP2. These observations point to structural
differences in the nucleotide binding pockets of TAP1 and TAP2. Indeed,
the NBD of TAP1 and TAP2 show only about 51% sequence identity; thus, structural differences might be expected at the atomic level. These
differences are likely to contribute to differences in the respective
nucleotide binding pockets, which might further lead to differing
affinities for nucleotide and differences in catalytic properties for
nucleotide hydrolysis. It remains to be addressed whether such
differences exist and, if so, whether there are any functional consequences.
1) is severalfold faster than that
previously reported (kd = 0.002 s
1) (27). The difference in the dissociation
rate might arise because the binding experiments were carried out at
room temperature and 10 °C, respectively. Using similar sets of
fluorescence quenching assays, we show here that TAP1(K544M)-His·TAP2
and TAP1·TAP2(K509M) complexes are capable of binding peptides,
although the binding affinity of TAP1·TAP2(K509M) complexes appears
weaker than wild type.
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ACKNOWLEDGEMENTS |
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We thank Dr. Robert Tampé for the baculovirus constructs encoding wild type TAP1 and TAP2 and for the anti-TAP1 antibody 148.3, Shikha Arora for doing the protease digestion experiments, the University of Michigan Hybridoma core for ascites preparations and for the maintenance and storage of hybridoma lines, the University of Michigan Biomedical Research Core facilities for DNA sequencing and peptide syntheses and purifications, and the University of Michigan Cell Biology laboratories for use of computer resources. We thank Dr. Wesley Dunnick for critical review of the manuscript.
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FOOTNOTES |
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* This work was supported by National Institutes of Health Grants RO1 AI44115-01 (to M. R.) and GM39561 (to R. N.) and by University of Michigan Multipurpose Arthritis and Musculoskeletal Diseases Center Grant 5P60AR20557.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: Dept. of Microbiology and Immunology, 5641 Medical Science Bldg. II, University of Michigan Medical School, Ann Arbor, MI 48109-0620. Tel.: 734-647-7752; Fax: 734-764-3562; E-mail: malinir@umich.edu.
Published, JBC Papers in Press, November 30, 2000, DOI 10.1074/jbc.M009448200
2 S. Arora and M. Raghavan, unpublished observations.
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ABBREVIATIONS |
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The abbreviations used are: TAP, transporter associated with antigen processing; MHC, major histocompatibility complex; ABC, ATP-binding cassette; NBD, nucleotide binding domain(s); PBS, phosphate-buffered saline; HPLC, high pressure liquid chromatography.
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