From the Department of Microbiology and Immunology, University of Michigan Medical School, Ann Arbor, Michigan 48109-0620
Received for publication, August 30, 2002, and in revised form, December 2, 2002
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ABSTRACT |
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The transporter associated with antigen
processing (TAP) contains two nucleotide-binding domains (NBD) in the
TAP1 and TAP2 subunits. When expressed as individual subunits or
domains, TAP1 and TAP2 NBD differ markedly in their nucleotide binding
properties. We investigated whether the two nucleotide-binding sites of
TAP1/TAP2 complexes also differed in their nucleotide binding
properties. To facilitate electrophoretic separation of the subunits
when in complex, we used TAP complexes in which one of the subunits was
expressed as a fluorescent protein fusion construct. In binding experiments at 4 °C using the photo-cross-linkable nucleotide analogs 8-azido-[ The transporter associated with antigen processing
(TAP)1 complex is an integral
part of the major histocompatibility complex class I antigen
presentation pathway (1). TAP is comprised of two related subunits,
TAP1 and TAP2, which are necessary and sufficient for peptide
translocation from the cytosol into the endoplasmic reticulum (2, 3).
Both proteins contain an N-terminal membrane-spanning region (MSR) and
a C-terminal nucleotide-binding domain (NBD). Peptide translocation by
TAP complexes is preceded by peptide binding to the cytosolic face of
TAP1/TAP2 complexes, a step that appears to be nucleotide
binding-independent, at least at low temperatures (4, 5). However, the
presence of ATP or ADP is critical for maintaining the structural
stability of TAP complexes at physiological temperatures (6), and TAP
mutants that are defective in nucleotide binding at the TAP2 site lose their ability to bind peptide at 37 °C but not at lower temperatures (7). ATP hydrolysis is critical for peptide translocation across the
endoplasmic reticulum membrane (8), and a peptide-stimulated ATPase activity has been described for TAP complexes (9).
The role of each NBD during peptide translocation is not well
understood. Mutations in both NBD have been shown to affect peptide
translocation efficiency, although mutations on TAP2 NBD generally have
more severe consequences. Based upon mutagenesis studies at
structurally analogous residues on TAP1 and TAP2, experiments from
several labs have resulted in the postulation of functional distinctions between a TAP1 and a TAP2 NBD during transport (5, 7, 10,
11). Our subsequent studies indicated that chimeric TAP complexes
containing two TAP1 NBD and two TAP2 NBD are capable of peptide
translocation but with reduced efficiency relative to wild type
complexes (12). TAP complexes containing two TAP2 NBD were less
efficient in nucleotide binding and peptide translocation, relative to
TAP complexes containing two TAP1 NBD. Likewise the isolated TAP2
subunit and TAP2 NBD bound nucleotides with reduced efficiency compared
with the isolated TAP1 subunit and TAP1 NBD (5, 7, 11, 13). These
observations raised the question of whether the TAP1 and TAP2
nucleotide-binding sites of TAP1/TAP2 complexes also had distinct
nucleotide binding properties.
TAP is a member of a family of transporter proteins called ATP-binding
cassette (ABC) transporters. ABC transporters have similar domain
organizations and have considerable sequence identity in their NBD.
Conserved sequence motifs within the NBD include the Walker A and
Walker B motifs, the consensus C (signature) motif, and the switch
region (reviewed in Ref. 14). There is increasing evidence in the
literature that the NBD of ABC transporters interact in a manner
similar to that observed in the DNA repair enzyme Rad50 (15). Indeed,
recent structures of the intact ABC transporter (BtuCD) (16) and of NBD
dimers of the ABC transporter MJ0796 (17) and biochemical studies
analyzing the products of vanadate-catalyzed photo-cleavage of the
maltose transporter NBD (18) have provided strong evidence in favor of
a Rad50-like NBD-NBD interface for ABC transporters. In such an
interaction, each ATP-binding site is comprised of residues that derive
from the interface of two NBD. If TAP1 and TAP2 NBD interact in a
Rad50-like manner, the TAP1 nucleotide-binding site would contain
residues primarily from the Walker A motif of TAP1 and from the
consensus C motif of TAP2, whereas the TAP2 nucleotide-binding site
would include residues primarily from the Walker A motif of TAP2 and the TAP1 consensus C motif (see "Discussion"). A Rad50-like
interface for ABC transporter NBD could explain the high degree of
conservation of the consensus C motif. Furthermore, a Rad50-like
interface provides a greater number of protein-nucleotide contacts than those observed in structures of ABC transporter NBD monomers (19-21). Therefore, it was conceivable that the nucleotide binding properties of
isolated TAP1 and TAP2 subunits were not reflective of their nucleotide
binding properties when in complex.
Human TAP1 and TAP2 are not well resolved on SDS-PAGE gels; however, by
creating a fusion protein of TAP1 and TAP2 with enhanced green
fluorescent protein (TAP1-eGFP) and enhanced yellow fluorescent protein
(TAP2-eYFP), respectively, the complexes of one tagged subunit and the
partner untagged subunit could be easily resolved and quantified on a
gel. By cross-linking radiolabeled 8-azido nucleotides at varying
concentrations, we were able to derive affinity constants for 8-azido
nucleotide binding to the individual TAP subunits when expressed alone
and in complex. We found that in a TAP1/TAP2 complex, but not in the
individual subunits, the nucleotide binding affinities derived from
labeling of the TAP1 and TAP2 subunits, respectively, were almost
identical. The TAP2 subunit when expressed alone had reduced affinity
for both nucleotide triphosphates and nucleotide diphosphates, compared
with TAP1.
Baculoviruses for Expression of TAP1, TAP2, TAP1(K544M),
TAP2(K509M), T2MT1C, T1MT2C, TAP1-eGFP, and
TAP2-eYFP--
Baculoviruses encoding wild type human TAP1 and TAP2
were obtained from the laboratory of Dr. Robert Tampé (22). We
have previously described the construction of baculoviruses encoding the TAP1 mutant (TAP1(K544M) and the TAP2 mutant (TAP2(K509M)) (5).
Construction of the chimeric TAP proteins T1MT2C (containing the TAP1
MSR and TAP2 NBD) and T2MT1C (containing the TAP2 MSR and TAP1 NBD) has
also previously been described (12).
For construction of TAP1 and TAP2 fluorescent protein fusions, human
TAP1 and TAP2 cDNA were obtained from Dr. John Trowsdale. Bridge
PCR was used to generate the TAP1-eGFP fusion, for which the eGFP
template was obtained from the pEGFP plasmid
(Clontech). The first PCR amplified the TAP1
portion of the fusion construct using a 5' primer with a
BglII site followed by a sequence complementary to the 5'
end of TAP1 and a 3' primer with the last 15 nucleotides of the TAP1
sequence and the first 15 nucleotides of the eGFP sequence. The second
PCR used a 5' primer that was complementary to the 3' primer used for
TAP1 amplification and a 3' primer complementary to the 3' end of the
eGFP sequence followed by a BglII site. Both of these PCR
products were gel-extracted and used as templates for a third PCR,
which used the 5' primer of the TAP1 PCR and the 3' primer of the eGFP
PCR. This bridge PCR product was gel-extracted, ligated into pPCRScript
(Stratagene), and sequenced. The TAP1-eGFP fusion was then ligated into
the Bgl II site of pAcUW51 (BD Pharmingen), which was used to generate
a TAP1-eGFP-encoding baculovirus using the BaculoGold Transfection kit
(BD Pharmingen).
For construction of the TAP2-eYFP fusion, TAP2 in pCR2.1 (5) was first
extracted by PCR using 5' and 3' primers that had BglII
sites. Two extra nucleotides were added between the last codon of TAP2
and the 3' BglII site to maintain the reading frame for YFP.
This PCR product was gel-purified and ligated into the vector pPCR
Script (Stratagene). The TAP2 was then excised from pPCRscript with
BglII and ligated into the vector pVL1393-YP (BD Pharmingen)
at the BamHI site to generate the TAP2 fusion with YFP. This
vector was then used to generate a TAP2-eYFP-encoding baculovirus.
Insect Cell Culture, Microsome Preparations, and Analyses of TAP
Expression--
Sf21 cells were cultured in Grace's insect
medium (Invitrogen), supplemented with 10% fetal bovine serum. The
cells were grown to confluence and infected with the desired
baculovirus combinations at appropriate multiplicities of infection
values (usually 5-30, depending on the individual baculovirus) and
incubated at 27 °C for ~60 h. Following these infections,
microsomal membrane fractions were generated as described (22). TAP
expression in the microsomes was verified by immunoblotting analyses
with the TAP1-specific antibody 148.3 (22) and the TAP2-specific
antibody (435.3) (23). Where comparison of the relative expression
levels of the different TAP constructs was required, we used the 1P3
antiserum (24), which recognizes all of the TAP constructs described here.
Peptide Translocation Experiments--
Iodinated peptide
translocation experiments were performed using the desired microsome
preparations as described previously (5). This procedure was based upon
well characterized assays (8, 22).
8-Azido-32P-Nucleotide Binding
Experiments--
Insect cell microsomes were washed three times in
assay buffer (40 mM Tris, 100 mM NaCl, 5 mM MgCl2, pH 7.0) and resuspended in half their
original volume in the same buffer. When
8-azido-[
For the ATP/ADP competition experiments, the same protocol as above was
used, except that varying amounts (0.25-600 µM) of unlabeled ATP or ADP were added to the tubes before adding the microsomes. The EC50 values were derived by plotting
labeling intensities corresponding to TAP1 and TAP2 as a function of
unlabeled ATP and ADP concentrations.
Expression, Translocation Function, and 8-Azido Nucleotide Binding
by Fluorescent Protein-tagged TAP Subunits or TAP1/TAP2
Complexes--
We created TAP1-eGFP and TAP2-eYFP fusion constructs
and expressed the proteins in insect cells using a baculovirus-based system. By expressing tagged TAP1 protein (TAP1-eGFP), tagged TAP2
protein (TAP2-eYFP), wild type TAP1, or wild type TAP2 in any
combination of a TAP1 and a TAP2, we were able to generate complexes
that were functional for peptide translocation. This was tested by
using insect cell microsomes expressing both tagged proteins
(TAP1-eGFP/TAP2-eYFP) or the individual subunits expressed singly in a
radiolabeled peptide-based translocation assay. Indeed, the tagged TAP
complex was able to translocate peptides, whereas the individual
subunits were not (Fig. 1B).
TAP complexes expressing one tagged subunit and one wild type subunit
(TAP1-eGFP/TAP2 and TAP1/TAP2-eYFP) were also competent for peptide
translocation (data not shown).
We next examined 8-azido-[ 8-Azido Nucleotide Binding to TAP Subunits Expressed Individually
and in Combination--
For all of the ATP binding analyses described
here on in the manuscript, we used 8-azido-[
The membranes were incubated with different concentrations of
photo-cross-linkable nucleotide analog
8-azido-[
TAP2-eYFP (or TAP2; data not shown), when expressed separately, had
reduced 8-azido-ATP binding affinity (KD = 19.3 ± 2.5 µM) compared with TAP1 (KD = 4.6 ± 1.9 µM) (or TAP1-eGFP; data not shown)
expressed alone (Fig. 2G and Table I). When TAP2 is complexed with excess
TAP1-eGFP, the affinity corresponding to the TAP2 labeling is increased
(KD = 2.7 ± 1 µM) (Fig.
2H and Table I). Mutation of the TAP2 Walker A lysine
residue (TAP2(K509M)) reduced the TAP2-associated signal and derived
affinity (KD > 20 µM when expressed
in complex with TAP1-eGFP) (Fig. 2, E, bottom
panel compared with top panel, and H and
Table I). Thus, the 8-azido-ATP binding affinity derived for TAP2 in
the wild type TAP1-eGFP/TAP2 complexes (KD = 2.7 ± 1 µM) must correspond to binding via residues
in the TAP2 Walker A motif. These studies indicated that the nucleotide
binding affinity of the isolated TAP2 subunit was lower than that of
the TAP1 counterpart but that TAP1/TAP2 complex formation induced changes that enhanced the nucleotide binding affinity at the TAP2 site.
When TAP1-eGFP is co-expressed with excess TAP2 (Fig. 2, C,
F, and I), the affinity corresponding to
TAP1-eGFP labeling was 2.1 ± 0.8 µM (Fig.
2I and Table I). In the corresponding
TAP1-eGFP/TAP2(K509M) complexes, a similar affinity was
derived corresponding to TAP1-eGFP labeling (KD = 2.8 ± 2.9 µM) (Fig. 2I and Table I). Thus, high affinity binding by TAP1-eGFP is visualized even in the
absence of high affinity nucleotide binding to TAP2. However, the
signals derived for TAP1-eGFP in the TAP1-eGFP/TAP2(K509M) mutant
complex are reduced compared with that derived for the wild type
complex (Fig. 2I), even though slightly higher levels of
TAP1-eGFP were present in the mutant complex (Fig. 2C,
lane 1 compared with lane 2).
Taken together, the observations described so far indicate that both
nucleotide-binding sites of TAP1/TAP2 complexes can bind 8-azido-ATP
with similar affinities. However, the signal intensities corresponding to TAP2 labeling are consistently lower than those observed for TAP1 labeling (for example, Fig. 2F, compare
TAP1-EGFP and TAP2 labeling in the upper panel), even when
the expression level of TAP1 is lower than TAP2 (Fig. 2C,
lane 1). Possible explanations for this observation are
considered under "Discussion."
The data shown in Fig. 2 were derived from analyses of
8-azido-[ TAP1/TAP2 NBD Interaction Appear to Contribute at Least
in Part to Enhanced Nucleotide Binding at the TAP2 Site upon
TAP1/TAP2 Complex Formation--
The marked affinity
enhancement at the TAP2 nucleotide-binding site upon TAP1/TAP2 complex
formation could result from a general stabilization of the TAP2
structure upon TAP1/TAP2 complex formation or, more specifically,
result from NBD-NBD interactions that allow a greater set of nucleotide
contacts at the TAP2 site of TAP1/TAP2 complexes compared with free
TAP2. To distinguish these possibilities, we compared affinities
corresponding to TAP1 and TAP2 labeling in TAP1/TAP2-eYFP complexes as
well as T1MT2C/TAP2-eYFP complexes. T1MT2C is a chimeric protein
containing the MSR of TAP1 and the NBD of TAP2 (12). We have previously
shown that this chimera forms complexes with TAP2 and that these
complexes are functional for peptide translocation, although with
reduced efficiency relative to wild type. The affinity corresponding to
TAP2-eYFP labeling was 1.8 ± 1.4 µM in
TAP1/TAP2-eYFP complexes, compared with 11.4 ± 2.4 µM in T1MT2C/TAP2-eYFP complexes (~6-fold different;
Fig. 3C, first and second
panels). As expected, the affinity corresponding to T1MT2C labeling was
also reduced in T1MT2C/TAP2-eYFP complexes (25 ± 14.2 µM) (Fig. 3). These observations suggested that TAP1/TAP2 NBD/NBD interactions are important for optimal nucleotide binding at
the TAP2 nucleotide-binding site and that TAP2/TAP2 NBD interactions or
TAP1/TAP2 MSR interactions are not fully sufficient for affinity enhancement at the TAP2 site.
We also examined 8-azido-ATP binding by TAP1-eGFP/T2MT1C complexes
(Fig. 3, B and C, third panel). T2MT1C
is a chimeric construct containing the MSR of TAP2 and the NBD of TAP1
(12). We previously showed strong binding of TAP1/T2MT1C complexes to
ATP and ADP agarose beads compared with TAP2/T1MT2C complexes, which
correlated with enhanced translocation efficiency of the TAP1/T2MT1C
complexes. Measurement of 8-azido-ATP binding to TAP1-eGFP/T2MT1C
complexes indicated an affinity of 0.5 ± 0.3 µM
corresponding to T2MT1C labeling and 0.4 ± 0.1 µM
corresponding to TAP1-eGFP labeling. Interestingly, at comparable
expression levels, the intensity of labeling of the T2MT1C construct
was significantly higher than that of TAP1-eGFP (Fig. 3, B
and C, third panel), a reversal of the trend
observed with TAP1-eGFP/TAP2 complexes (Figs. 2F and 3,
B and C, top panels). Furthermore
unlike TAP2, T2MT1C bound 8-azido-ATP with high affinity regardless of
being complexed with TAP1 (data not shown). Thus, whereas high affinity
nucleotide binding to a TAP1 NBD is observed both in the presence and
absence of a TAP2 NBD, high affinity nucleotide binding to a TAP2 NBD requires the presence of a TAP1 NBD.
Labeling of Nucleotide Binding-deficient TAP1 Is Dependent upon the
Presence of a Functional TAP2 Nucleotide-binding Site--
The above
observations raised the possibility that the presence of TAP1 residues
at the TAP2 nucleotide-binding site could be responsible for the
observed affinity enhancement at the TAP2 site upon TAP1/TAP2 complex
formation. To further explore this possibility and also investigate the
effect of TAP1 nucleotide binding upon TAP2 labeling, we examined
8-azido-ATP binding by a mutant TAP complex containing a defective TAP1
nucleotide-binding site (Fig. 4). We
previously showed that mutation of the Walker A lysine of TAP1
(TAP1(K544M)) resulted in significantly reduced binding of single
subunit TAP1 to ATP- and ADP-agarose beads (5). Surprisingly, however,
when TAP1(K544M) was expressed in combination with TAP2-eYFP, the
labeling intensities observed for TAP1(K544M) and TAP2-eYFP were almost
identical over the entire concentration range, and thus, nearly
identical affinities were derived (3.8 ± 3.5 and 4.0 ± 2.5 µM, corresponding to TAP1(K544M) and TAP2-eYFP labeling,
respectively; Fig. 4, A-C). The parent TAP1/TAP2-eYFP complexes yielded KDvalues of 1.5 ± 0.4 and
1.3 ± 0.5 µM, corresponding to labeling of TAP1 and
TAP2-eYFP, respectively (data not shown).
We also investigated labeling of TAP1(K544M) in the absence of a
functional TAP2 nucleotide-binding site. For these analyses, we
prepared microsomes containing TAP1(K544M)/TAP2 complexes and TAP1(K544M)/TAP2(K509M) complexes. Because only untagged versions (but
not fluorescent protein fusions) of the mutant TAP subunits were
available, only the combined signal on both the TAP1 and TAP2
components could be visualized with these complexes (Fig. 4,
D and E). At comparable expression levels of both
components (Fig. 4D), strong labeling was visualized for the
TAP1(K544M)/TAP2 combination, whereas signals for the
TAP1(K544M)/TAP2(K509M) combination were barely detectable (Fig.
4E, top and middle panels,
respectively). These results demonstrated that the TAP1(K544M)
component by itself did not bind very efficiently to
8-azido-[ TAP1 and TAP2 NBD sequences are ~60% identical. Previous
studies have shown that the two NBD interact differently with
nucleotide-agarose beads, with constructs containing TAP1 NBD binding
strongly, and with constructs containing TAP2 NBD interacting weakly
with nucleotide-agarose beads (12, 27). In the present studies, we
examined nucleotide binding to the TAP1 and TAP2 sites, when the
proteins were in complex. We constructed C-terminal fusions of TAP1 and
TAP2 with eGFP and eYFP, respectively. When either fusion protein was
paired with a wild type partner, the resulting complexes were
functional and could be electrophoretically separated from the partner
wild type subunit, allowing measurements of the binding of various 8-azido-adenosine nucleotides to TAP subunits when in complex.
The presence of the eGFP or eYFP fusion constructs did not influence
the derived affinity, because similar affinities were observed for TAP1
and TAP1-eGFP or for TAP2 and TAP2-eYFP, both when expressed
individually or as a complex. The nucleotide binding affinity of the
isolated TAP2 subunit was found to be reduced compared with the TAP1
counterpart. However, TAP1/TAP2 complex formation induced changes that
significantly enhanced the affinity of the TAP2 nucleotide-binding site
for both 8-azido-ATP and 8-azido-ADP. TAP1/TAP2 complex formation has a
less significant effect upon the affinity of the TAP1
nucleotide-binding site for 8-azido-ATP and 8-azido-ADP (Table I).
Using TAP1-eGFP/TAP2(K509M) complexes under conditions of TAP1 excess
or TAP2 excess (Fig. 2), we determined that the two nucleotide-binding
sites of TAP1/TAP2 complexes did in fact bind 8-azido-ATP with apparent
affinities that were, within the error of these measurements, quite
similar to each other. The EC50 values derived from
inhibition analyses with ATP (Fig. 1E) were quite similar to
the KD values derived from direct
8-azido-[ Functional differences between TAP1 and TAP2 have previously been
attributed to differences in nucleotide binding to the TAP1 and TAP2
sites of TAP1/TAP2 complexes. Based upon studies with rat TAP1 and
TAP2, it has been suggested that TAP1 binds ATP more efficiently than
does TAP2, whereas the binding of ADP by the two chains is essentially
equivalent (10). Our present studies demonstrate that the affinities at
the TAP1 and TAP2 sites for both nucleotides are in fact quite similar
when TAP1 and TAP2 are in complex (Table I). By contrast, analogous
studies with CFTR have indicated that the two NBD bind equivalently to
8-azido-ADP but differed in their interactions with 8-azido-ATP. The
binding curves for 8-azido-ATP showed the occurrence of simple
saturable binding at NBD1 but a sigmoidal profile at NBD2, suggesting
that the two NBDs of CFTR had distinct nucleotide binding properties (25). These observations suggest the possibility of mechanistic differences between the TAP and CFTR catalytic cycles.
The affinity enhancement at the TAP2 nucleotide-binding site upon
TAP1/TAP2 complex formation could result from a general stabilization
of the TAP2 structure upon TAP1/TAP2 complex formation or, more
specifically, could be due to NBD-NBD interactions that alter the
nucleotide interaction properties at the TAP2 site. The observations of
enhanced nucleotide binding by TAP2-eYFP in TAP1/TAP2-eYFP complexes
compared with T1MT2C/TAP2-eYFP complexes (Fig. 3) argue for a role for
NBD-NBD interactions in determining the binding affinity of the TAP2
nucleotide-binding site. In a model of the TAP2 nucleotide-binding site
based upon MJ0796 dimer structure (17), several residue differences are
predicted when the opposing NBD has a TAP2 sequence rather than a TAP1
sequence (Fig. 5). Apparently, the sum of
such differences accounts for the observed reduction in affinity (Fig.
3) when a TAP2-TAP1 NBD-NBD interface is altered to a TAP2-TAP2 NBD-NBD
interface.
-32P]ATP and
8-azido-[
-32P]ADP, TAP2 was found to have reduced
affinity for nucleotides compared with TAP1, when the two proteins were
separately expressed. Complex formation with TAP1 enhanced the binding
affinity of the TAP2 nucleotide-binding site for both nucleotides.
Binding analyses with mutant TAP complexes that are deficient in
nucleotide binding at one or both sites provided evidence for the
existence of two ATP-binding sites with relatively similar affinities
in TAP1/TAP2 complexes. TAP1/TAP2 NBD interactions appear to contribute
at least in part to enhanced nucleotide binding at the TAP2 site upon
TAP1/TAP2 complex formation. Binding analyses with mutant TAP complexes
also demonstrate that the extent of TAP1 labeling is dependent upon the
presence of a functional TAP2 nucleotide-binding site.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-32P]ATP was used, the following ATPase
inhibitors were added to the assay buffer: 5 mM
NaN3, 2 mM EGTA, and 1 mM ouabain.
8-Azido-[
-32P]ATP, 8-azido-[
-32P]ATP,
or 8-azido-[
-32P]ADP (Affinity Labeling Technologies)
and assay buffer were combined in a total volume of 10 µl, following
which 10 µl of the washed microsomes were added. After a 15-min
incubation on ice, the samples were transferred to wells of a 96-well
plate on ice, cross-linked immediately with a 254-nm UV lamp for 3 min,
and then transferred back into tubes. The samples were then washed
three times with assay buffer, resuspended in SDS-PAGE sample buffer,
and heated to 95 °C for 5 min. The proteins were separated on 10%
polyacrylamide gels and dried using a Bio-Rad model 483 gel drier. The
dried gels were exposed to a PhosphorImager plate overnight, and
labeled TAP proteins were visualized using a PhosphorImager SI
(Molecular Dynamics). Quantifications were obtained using ImageQuant
software. After background subtraction, the amounts of radioactivity
observed were plotted and analyzed using the Prism software package
(graph Pad software). The data were fitted to a one-site binding
equation, Y = Bmax*X/(KD + X), where Bmax is the maximal
binding, X is the concentration of 8-azido nucleotide, and
KD is the binding constant for
8-azido-32P-nucleotide. Typically, between 60 and 80% of
the input radioactivity was associated with the supernatants from the
first centrifugation step after cross-linking, with a higher percentage
being associated at the higher concentrations of 8-azido nucleotides.
Analogous binding experiments with crude membranes have previously been used to estimate the 8-azido nucleotide binding affinities of the NBD
of P-glycoprotein and CFTR, respectively (25, 26).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Peptide translocation and nucleotide binding
by fluorescent protein-tagged TAP constructs. A and
B, The TAP1-eGFP/TAP2-eYFP complex is able to mediate
peptide translocation in an ATP-dependent manner, whereas
the individual TAP subunits do not. A, immunoblotting
analyses of the microsomes used for the translocation analyses in
B. 148.3 is a TAP1-specific antibody, and 435.3 is a
TAP2-specific antibody. B, microsomes expressing the
indicated TAP constructs and combination were analyzed for their
ability to translocate an iodinated reporter peptide in the presence of
ATP (+ATP) or apyrase ( ATP). C and
D, 8-azido-[
-32P]ATP binding by microsomes
expressing the indicated TAP complexes or subunits. C,
immunoblotting analyses of the microsomes used for the nucleotide
binding analyses in D. D, the indicated
microsomes were incubated with 2 µM
8-azido-[
-32P]ATP for 15 min on ice and subsequently
cross-linked by UV irradiation. The proteins were separated by 10%
SDS-PAGE. The TAP bands were the major products detected by
phosphorimaging analyses. Additional lower molecular weight bands are
visualized (indicated by asterisks). E and
F, microsomes expressing TAP1-eGFP/TAP2 complexes were
incubated with 2 µM 8- azido-[
-32P]ATP
and the indicated concentrations of unlabeled ATP (E) or ADP
(F) for 15 min on ice and subsequently cross-linked by UV
irradiation. The proteins were separated by 10% SDS-PAGE, and the TAP
bands were quantitated after phosphorimaging analyses and plotted
as a function of ATP or ADP concentration to derive the indicated
EC50 values (average of two independent sets of
analyses).
-32P]nucleotide binding to
individual TAP subunits or TAP complexes in which one of the subunits was expressed as a fluorescent protein fusion construct. To examine 8-azido-[
-32P]nucleotide binding in the absence of
detergent, we incubated insect cell microsomes expressing one or both
TAP subunits or neither protein with 8-azido-[
-32P]ATP
for 15 min at 4 °C, followed by UV-induced covalent cross-linking of
bound 8-azido-[
-32P]nucleotides. The membranes were
washed to remove unbound nucleotide. The proteins were separated by
SDS-PAGE, and proteins that had bound
8-azido-[
-32P]nucleotides were visualized using
phosphorimaging analyses of dried gels. Compared with microsomes
derived from uninfected cells, additional labeled bands were visualized
in microsomes expressing one or both TAP subunits at the expected
molecular weights corresponding to the wild type subunits and/or that
corresponding to the fluorescent protein fusion constructs (Fig.
1D, lane 1 compared with other lanes). The use of
the fluorescent protein epitope tags allowed for adequate separation of
the TAP subunits by SDS-PAGE and for visualization of the labeling of
individual components of the TAP1/TAP2 complex by
8-azido-[
-32P]nucleotides (Fig. 1D,
lane 2). Labeling of TAP subunits was inhibitable by
unlabeled ATP and ADP, with derived EC50 values in
the low micromolar range (Fig. 1, E and F, respectively).
-32P]ATP
rather than 8-azido-[
-32P]ATP, because we found that
insect cell microsomal membrane preparations had measurable ATPase
activity, even at 4 °C in the presence of the ATPase inhibitors
ouabain, EGTA, and sodium azide (regardless of TAP expression).
Although 8-azido-[
-32P]ATP would be expected to be
hydrolyzed as efficiently as 8-azido-[
-32P]ATP,
labeling by 8-azido-[
-32P]ATP would report just on
nucleotide triphosphate binding to TAP complexes. Microsomal membrane
fractions were prepared of insect cells infected with viruses encoding
single TAP subunits (Fig. 2A),
or TAP1/TAP2 combinations (Fig. 2, B and C). For
examining nucleotide binding by TAP2 when in complex, the TAP1
component was expressed in excess (Fig. 2B), and for
examining nucleotide binding by TAP1 when in complex, the TAP2
component was expressed in excess (Fig. 2C).
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Fig. 2.
Complex formation with TAP1 markedly
increases the 8-azido-ATP binding affinity of the TAP2
nucleotide-binding site. Insect cell microsomal membranes
expressing TAP1 alone or TAP2-eYFP alone (A), the
TAP1-eGFP/TAP2 or TAP1-eGFP/TAP2(K509M) combinations with the TAP1-eGFP
component in excess (B), or the TAP1-eGFP/TAP2 or
TAP1-eGFP/TAP2(K509M) combinations with the TAP2 or
TAP2(K509M) components in excess (C) were
incubated with different concentration of
8-azido-[ -32P]ATP for 15 min on ice and subsequently
cross-linked by UV irradiation. The proteins were then separated by
10% SDS-PAGE, and radiolabeling of TAP subunits was visualized by
phosphorimaging analyses. A-C, immunoblotting analyses to
visualize one or both TAP proteins. The 1P3 antibody recognized both
TAP1 and TAP2. D-F, phosphorimaging analyses of labeled TAP
proteins. G-I, quantification of labeled TAP bands and
fitting to a one site binding model. The KD values
corresponding to labeling of the indicated TAP subunit(s) were derived
from multiple independent analyses, summarized in Table I.
-32P]ATP, followed by UV cross-linking and
analyses of TAP labeling by SDS-PAGE and phosphorimaging analyses (Fig.
2, D-F). The extent of labeling of the TAP subunits was
quantified using ImageQuant software, plotted as a function of the
input 8-azido-[
-32P]ATP concentration, and then fitted
to a one-site binding model to derive KD values
corresponding to TAP1 and TAP2 labeling, when expressed individually or
in combination (Fig. 2, G-I).
Apparent affinities of the wild type or mutant TAP subunits for
8-azido-[-32P]ATP and 8-azido-[
-32P]ADP when
expressed individually or in combination with an excess of partner
TAP subunit.
-32P]ATP binding to TAP1/TAP2 complexes. A
similar set of analyses was also carried out using
8-azido-[
-32P]ADP, and analogous results were obtained
(Table I). TAP2-eYFP had reduced affinity for
8-azido-[
-32P]ADP compared with TAP1 when the two
subunits were separately expressed. Complex formation with TAP1
increased the binding affinity corresponding to TAP2 labeling by 9-fold
(Table I), whereas complex formation with TAP2 increased the binding
affinity corresponding to TAP1 labeling by only 2-fold (Table I).
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Fig. 3.
8-Azido-ATP binding affinities at the two
nucleotide-binding sites of TAP1/TAP2-eYFP complexes or chimeric TAP
complexes containing two TAP1 NBD or two TAP2 NBD. Insect cell
microsomal membranes expressing TAP1/TAP2-eYFP, T1MT2C/TAP2-eYFP (two
TAP2 NBDs), or TAP1-eGFP/T2MT1C (two TAP1 NBDs) were incubated with
different concentration of 8-azido-[ -32P]ATP for 15 min on ice and subsequently cross-linked by UV irradiation. The
proteins were then separated by 10% SDS-PAGE, and radiolabeling of TAP
subunits was visualized by phosphorimaging analyses. A,
immunoblotting analyses to visualize components of each complex.
B, phosphorimaging analyses of labeled TAP proteins.
C, quantification of labeled TAP bands and fitting to a
one-site binding model. The indicated KD values are
the averages of two or three independent sets of analyses.
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Fig. 4.
Labeling of nucleotide binding-deficient TAP1
is dependent on the presence of a functional TAP2 nucleotide-binding
site. Labeling procedures were similar to that described in the
legend to Fig. 2. A and D, immunoblotting
analyses of microsomes used in binding analyses shown in B
and E, respectively. B, phosphorimaging analyses
of 8-azido-[ -32P]ATP binding by the TAP1(K544M) and
TAP2-eYFP components of TAP1(K544M)/TAP2-eYFP complexes. C,
quantification of signals from B and estimation of
KD values. The indicated KD
values are the averages of three independent sets of analyses. The
KD values derived for a corresponding set of
analyses with wild type TAP1/TAP2-eYFP complexes were 1.5 ± 0.4 µM (for TAP1) and 1.3 ± 0.5 µM (for
TAP2-eYFP). E, phosphorimaging analyses of
8-azido-[
-32P]ATP binding to microsomes containing
TAP1(K544M)/TAP2 (top panel), TAP1(K544M)/TAP2(K509M)
(middle panel), or TAP1(K544M)/TAP2-eYFP (bottom
panel). Signals corresponding to TAP1 (K544M) could be observed
when in complex with TAP2-eYFP but not when in complex with
TAP2(K509M). The absence of a signal was not due to the expression
level, as TAP1(K544M) was expressed at higher levels in the microsomes
with the TAP2(K509M) combination compared with the TAP2-eYFP
combination (see D).
-32P]ATP, consistent with our previous report
that the TAP1(K544M) mutant was impaired in binding to ATP-agarose and
ADP-agarose beads relative to wild type TAP1 (5). Most interestingly,
although distinct labeled bands corresponding to TAP1(K544M) were
observed in TAP1(K544M)/TAP2-eYFP complexes (Fig. 4E,
bottom panel), a corresponding distinct signal was not
visualized in the TAP1(K544M)/TAP2(K509M) combination (Fig.
4E, middle panel), despite the higher expression of TAP1(K544M) in the latter complexes. These observations, taken together with the result that the affinity corresponding to TAP1(K544M) labeling complexes was very similar to that derived for TAP2-eYFP labeling in TAP1(K544M)/TAP2-eYFP complexes (Fig. 4C),
suggested the possibility of labeling of TAP1(K544M) residues that are
in the vicinity of the TAP2 nucleotide-binding site.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-32P]ATP binding analyses (Table I). The
EC50 values derived from inhibition analyses with ADP (Fig.
1F) were higher than the KD value derived
from the direct 8-azido-[
-32P]ADP binding analyses
(Table I). Thus, it is possible that 8-azido-ADP does bind to TAP
complexes with higher affinity than ADP, as reported for other ABC transporters.
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Fig. 5.
Proposed architectures of the TAP2
nucleotide-binding site of TAP1/TAP2 complexes (A)
compared with T1MT2C/TAP2 complexes (two TAP2 NBD)
(B), based upon the structure of the MJ0796 dimer
(17). Residues predicted to be involved in nucleotide contacts
from the Walker A end and the Consensus C end are indicated for each
site, and the residues involved in protein-protein contacts are
indicated in the extreme left of each panel. The C-8 carbon
of a TAP2-bound nucleotide is predicted to be in close proximity to
Tyr477 of TAP2 as well as Gln642 of TAP1.
Because cross-linking by an azido group is not residue-specific, an
8-azido-ATP bound at the TAP2 site could potentially be cross-linked in
the vicinity of either of these residues, which could explain the
results with TAP1(K544M)/TAP2-eYFP complexes (Fig. 4).
Mutation of the TAP2 Walker A lysine residue (TAP2(K509M)) indeed influenced nucleotide binding at the TAP2 site (Fig. 2). This effect of the TAP2 Walker A mutation was not apparent in previous analyses of the binding to nucleotide agarose beads of single subunit wild type and mutant TAP2 (5, 11), presumably because of the low binding affinities of both wild type and mutant TAP2. The corresponding mutation in TAP1 (TAP1(K544M)) had a marked effect upon the binding of single subunit TAP1 to nucleotide agarose beads (5, 7). Surprisingly, we found that when the TAP1 Walker A mutant TAP1(K544M) was expressed in combination with TAP2-eYFP, the signal intensities as well as the affinities corresponding to TAP1 and TAP2 labeling were nearly identical (Fig. 4). This was apparent in analyses with both 8-azido-ATP (Fig. 4) and 8-azido-ADP (data not shown). However, co-expression of TAP1(K544M) with TAP2(K509M) resulted in a nucleotide-binding deficient complex (Ref. 7 and Fig. 4). These observations demonstrate a profound influence of nucleotide binding by TAP2 upon TAP1 labeling under conformational conditions that are prevalent in TAP1(K544M)/TAP2 complexes.
What mechanisms could be responsible for enhanced TAP1 labeling in
TAP1(K544M)/TAP2-eYFP complexes compared with TAP1(K544M)/TAP2(K509M) complexes? In Rad50 and MJ0796-like model of NBD-NBD interactions, ATP
binding was required to promote NBD dimerization (Fig.
6A) (15, 17). If TAP NBD
interactions resemble Rad50 and MJ0796, it is conceivable that
disruption of ATP binding by Walker A mutations at the TAP1 site or
TAP2 site could result in disrupted NBD-NBD contacts at the mutated
nucleotide-binding site. In the case of the mutant TAP1(K544M)/TAP2
complexes, a concurrent "closed" conformation at the TAP2
nucleotide-binding site could result in labeling of TAP1(K544M)
residues (in the vicinity of the consensus C motif) that are in close
proximity to the TAP2 nucleotide-binding site (Fig. 6B). An
alternative possibility is that nucleotide binding at the TAP2 site
could enhance nucleotide binding at the TAP1 site by increasing the
affinity or accessibility of the TAP1 nucleotide-binding site (Fig.
6B). When we compared
8-azido-[-32P]ATP binding by TAP1- eGFP in
TAP1-eGFP/TAP2 complexes and TAP1-eGFP/TAP2(K509M) complexes, we found
that the TAP1-eGFP labeling intensity was enhanced when in complex with
wild type TAP2 compared with TAP2(K509M). Importantly, however, the
binding affinity was unchanged (Fig. 2I). These results
indicated that nucleotide binding at the TAP2 site could enhance
nucleotide binding at the TAP1 site by inducing a more "open"
(nucleotide-accessible) conformation at the TAP1 nucleotide-binding
site (but not by an affinity alteration at the TAP1 site). We observed
that the derived affinity corresponding to TAP1(K544M) labeling in
TAP1(K544M)/TAP2-eYFP complexes was nearly identical to that
corresponding to TAP2-eYFP labeling (Fig. 4C) and
significantly higher than that measured in TAP1(K544M)/TAP2(K509M) (Fig. 4E, middle panel; KD
cannot be estimated because significant labeling was not visualized).
Thus, the TAP2(K509M) mutation appears to have distinct effects on
labeling of TAP1 compared with TAP1(K544M). A likely explanation for
this apparent discrepancy is that there are two possible high affinity
nucleotide-binding sites in TAP1: one in the vicinity of its Walker A
sequence and a second in the vicinity of its consensus C sequence.
Labeling of the latter site is dependent upon the presence of a
functional TAP2 nucleotide-binding site, whereas the labeling of the
first site is independent of a functional TAP2 site. Disruption of the first site by the K544M mutation renders high affinity TAP1(K544M) labeling dependent upon the presence a functional TAP2 site. In a model
of TAP1/TAP2 based upon the MJ0796 structure (17), the C-8 carbon of a
TAP2-bound nucleotide is predicted to be in close proximity to
Tyr477 of TAP2 as well as Gln642 of TAP1 (Fig.
5A). Because cross-linking by an azido group is not
residue-specific, an 8-azido-ATP bound at the TAP2 site could potentially be cross-linked in the vicinity of either of these residues, which could explain the results with TAP1(K544M)/TAP2-eYFP complexes (Fig. 4).
|
The above arguments raise the possibility of TAP1 residues being in close proximity to the TAP2 nucleotide-binding site of TAP1/TAP2 complexes, under some conformational conditions, such as that trapped in TAP1(K544M)/TAP2-eYFP complexes. High affinity labeling of TAP1(K544M) residues in TAP1(K544M)/TAP2-eYFP complexes (Fig. 4), but not of TAP2(K509M) residues in TAP1-eGFP/TAP2(K509M) complexes (Fig. 2), might arise because of conformational differences between the two mutant complexes. Alternatively, structural differences between the TAP1 and TAP2 nucleotide-binding sites, such as differences in the consensus C residues (Fig. 6), could influence the extent of NBD-NBD interaction at each nucleotide-binding site. It remains to be defined whether other conformational conditions exist in which TAP2 residues are in close proximity to the TAP1 nucleotide-binding site. Additionally, it remains to be established whether TAP1 residues are in close proximity to the TAP2 nucleotide-binding site, in resting state wild type TAP1/TAP2 complexes. Reanalyses of the TAP1 binding data in Fig. 2F (for TAP1-eGFP/TAP2) using a two-site binding model (rather than a single site binding model) did not yield better R2 values. Furthermore, consistent KD 2 values were not derived in two-site binding analyses of three independent experimental data sets. The analysis is somewhat complicated by the result that the TAP1 and TAP2 sites have nearly identical affinities in TAP1/TAP2 complexes (Fig. 2) and thus do not allow us to distinguish between two-site or single-site binding models for TAP1 in TAP1-eGFP/TAP2 complexes. Additional experiments will be required to demonstrate the exact proximity of TAP1 consensus C (and other residues) to the TAP2 nucleotide-binding site, and vice versa, under different conformational conditions.
Although the affinities derived for the TAP1 and TAP2 nucleotide-binding sites of TAP1/TAP2 complexes were very similar, we observed that the signal intensities were generally higher for TAP1 labeling compared with TAP2 labeling, even if the TAP2 expression was higher (for example Fig. 2F). Many factors could influence the observed labeling intensity, including the 8-azido-ATP coupling efficiency (which could vary as a function of the exact chemical environment of a given nucleotide-binding site), and the relative populations of active proteins (stability). It is also possible, as suggested in Fig. 6B, that different conformational states of TAP exist in which the two nucleotide-binding sites vary in their nucleotide accessibilities. In the resting state of TAP1/TAP2 complexes, the TAP2 site could be in a more closed (nucleotide-inaccessible) conformation compared with the TAP1 site, which would result in lower efficiency TAP2 labeling compared with TAP1 labeling. Although our studies clearly demonstrate that the inherent binding affinities of the two sites are very similar in TAP1/TAP2 complexes, more information is required pertaining to the accessibility of each site in resting state and transition state conformations.
The analyses undertaken here also allow for a reassessment of the effects of TAP1(K544M) and TAP2(K509M) mutations upon peptide binding to TAP1/TAP2 complexes. We show here as previously suggested (7) that both mutations significantly reduce the nucleotide binding affinities at the corresponding nucleotide-binding sites. Both mutant complexes were found to bind TAP-specific peptides with high affinity at room temperature (5); however, whereas the binding affinity of the TAP1(K544M)/TAP2 complex (KD = 17.4 ± 4.8 nM) was very similar to wild type (KD = 19.4 ± 4.8 nM), the affinity of the TAP1/TAP2(K509M) was ~2-fold reduced (KD = 39.2 ± 5.9 nM). Thus, a 10-fold or greater reduction in the nucleotide binding affinity at the TAP2 site has a small effect on the peptide binding affinity, whereas nucleotide binding at the TAP1 site does not appear to influence peptide binding. Other studies found that a defective TAP2 nucleotide-binding site resulted in a loss of peptide binding at 37 °C (but not at low temperatures), which was interpreted as indicating that nucleotide binding to TAP2 was required for peptide binding (7). Our observation is that nucleotide binding to TAP1/TAP2 complexes significantly enhances the thermostability of the TAP peptide-binding site (28) but that nucleotide binding per se is nonessential for peptide binding, because mutant TAP complexes are capable of peptide binding with high affinity at reduced temperatures (5). Such a stability model is sufficient to explain why mutant TAP complexes containing defective TAP2 nucleotide-binding sites are defective for peptide binding when expressed in mammalian cells that are cultured at 37 °C (10, 11).
In summary, we demonstrate here (i) that TAP1 and TAP2 NBD differ
markedly in their nucleotide binding properties when expressed as
individual subunits but not when expressed as a complex, (ii) that
TAP1/TAP2 NBD-NBD interactions are critical for optimal nucleotide binding to the TAP2 site of TAP1/TAP2 complexes, and (iii) that the
extent of TAP1 labeling is dependent upon the presence of a functional
TAP2 nucleotide-binding site. Although these studies establish that the
TAP1 and TAP2 nucleotide-binding sites of TAP1/TAP2 complexes are both
capable of binding nucleotides with similar affinities, we cannot
presently assess whether the two sites differ in nucleotide
accessibility in resting state TAP1/TAP2 complexes. Furthermore,
whether both sites are capable of ATP hydrolysis also remains to be
established. Our findings that chimeric TAP1/TAP2 complexes containing
two TAP1 NBD were functional for peptide translocation (with reduced
efficiency relative to wild type TAP1/TAP2 complexes) indicated that a
TAP1 NBD is capable of ATP hydrolysis, at least when the opposing NBD
has the TAP1 sequence (12). Likewise, TAP complexes containing two TAP2
NBD were also functional for peptide translocation, although with low
efficiency. These studies have suggested that both nucleotide-binding
sites of TAP1/TAP2 complexes can hydrolyze ATP. However, this remains
to be unambiguously established using wild type TAP1/TAP2 complexes.
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ACKNOWLEDGEMENTS |
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We thank Richard Cantley for generating the TAP2-eYFP construct. We are grateful to Dr. Richard Neubig for critical comments and many valuable suggestions. We thank Dr. Robert Tampé for the baculoviruses encoding TAP1 and TAP2 and the 148.3 antibody and Dr. Frank Momburg for the 1P3 antiserum. We thank the University of Michigan Biomedical Research Core facilities for peptide synthesis and purification, the University of Michigan hybridoma core for 435.3 ascites fluids, and the University of Michigan Reproductive Sciences Program for peptide iodination.
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FOOTNOTES |
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* This work was supported by National Institutes of Health Grant AI44115-03 (to M. R.) and by National Institutes of Health Rheumatic Disease Core Center Grant AR48310-02 (to the University of Michigan).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.
These authors contributed equally to this work.
§ 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, December 25, 2002, DOI 10.1074/jbc.M208930200
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ABBREVIATIONS |
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The abbreviations used are: TAP, transporter associated with antigen processing; TAP1, TAP subunit 1; TAP2, TAP subunit 2; TAP1-eGFP, TAP1 and enhanced green fluorescence protein fusion construct; TAP2-eYFP, TAP2 and enhanced yellow fluorescence protein fusion construct; NBD nucleotide-binding domain(s), MSR, membrane-spanning region; ABC, ATP-binding cassette; T1MT2C, a chimeric protein containing the MSR of TAP1 and the NBD of TAP2; T2MT1C, a chimeric protein containing the MSR of TAP2 and the NBD of TAP1; CFTR, cystic fibrosis transmembrane conductance regulator.
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REFERENCES |
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---|
1. | Pamer, E., and Cresswell, P. (1998) Annu. Rev. Immunol. 16, 323-358[CrossRef][Medline] [Order article via Infotrieve] |
2. | Abele, R., and Tampe, R. (1999) Biochim. Biophys. Acta 1461, 405-419[Medline] [Order article via Infotrieve] |
3. | Momburg, F., and Hammerling, G. J. (1998) Adv. Immunol. 68, 191-256[Medline] [Order article via Infotrieve] |
4. |
Androlewicz, M. J.,
Ortmann, B.,
van Endert, P. M.,
Spies, T.,
and Cresswell, P.
(1994)
Proc. Natl. Acad. Sci. U. S. A.
91,
12716-12720 |
5. |
Lapinski, P. E.,
Neubig, R. R.,
and Raghavan, M.
(2001)
J. Biol. Chem.
276,
7526-7533 |
6. |
van Endert, P. M.
(1999)
J. Biol. Chem.
274,
14632-14638 |
7. |
Saveanu, L.,
Daniel, S.,
and van Endert, P. M.
(2001)
J. Biol. Chem.
276,
22107-22113 |
8. | Neefjes, J. J., Momburg, F., and Hammerling, G. J. (1993) Science 261, 769-771[Medline] [Order article via Infotrieve] |
9. |
Gorbulev, S.,
Abele, R.,
and Tampe, R.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
3732-3737 |
10. | Alberts, P., Daumke, O., Deverson, E. V., Howard, J. C., and Knittler, M. R. (2001) Curr. Biol. 11, 242-251[CrossRef][Medline] [Order article via Infotrieve] |
11. |
Karttunen, J. T.,
Lehner, P. J.,
Gupta, S. S.,
Hewitt, E. W.,
and Cresswell, P.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
7431-7436 |
12. |
Arora, S.,
Lapinski, P. E.,
and Raghavan, M.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
7241-7246 |
13. |
Lapinski, P. E.,
Miller, G. G.,
Tampe, R.,
and Raghavan, M.
(2000)
J. Biol. Chem.
275,
6831-6840 |
14. | Jones, P. M., and George, A. M. (1999) FEMS Microbiol. Lett. 179, 187-202[CrossRef][Medline] [Order article via Infotrieve] |
15. | Hopfner, K. P., Karcher, A., Shin, D. S., Craig, L., Arthur, L. M., Carney, J. P., and Tainer, J. A. (2000) Cell 101, 789-800[Medline] [Order article via Infotrieve] |
16. |
Locher, K. P.,
Lee, A. T.,
and Rees, D. C.
(2002)
Science
296,
1091-1098 |
17. | Smith, P. C., Karpowich, N., Millen, L., Moody, J. E., Rosen, J., Thomas, P. J., and Hunt, J. F. (2002) Mol. Cell 10, 139-149[Medline] [Order article via Infotrieve] |
18. |
Fetsch, E. E.,
and Davidson, A. L.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
9685-9690 |
19. |
Yuan, Y. R.,
Blecker, S.,
Martsinkevich, O.,
Millen, L.,
Thomas, P. J.,
and Hunt, J. F.
(2001)
J. Biol. Chem.
276,
32313-32321 |
20. | 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] |
21. |
Gaudet, R.,
and Wiley, D. C.
(2001)
EMBO J.
20,
4964-4972 |
22. | Meyer, T. H., van Endert, P. M., Uebel, S., Ehring, B., and Tampe, R. (1994) FEBS Lett. 351, 443-447[CrossRef][Medline] [Order article via Infotrieve] |
23. | van Endert, P. M., Tampe, R., Meyer, T. H., Tisch, R., Bach, J. F., and McDevitt, H. O. (1994) Immunity 1, 491-500[Medline] [Order article via Infotrieve] |
24. | Nijenhuis, M., and Hammerling, G. J. (1996) J. Immunol. 157, 5467-5477[Abstract] |
25. |
Aleksandrov, L.,
Aleksandrov, A. A.,
Chang, X. B.,
and Riordan, J. R.
(2002)
J. Biol. Chem.
277,
15419-15425 |
26. |
Sauna, Z. E.,
Smith, M. M.,
Muller, M.,
and Ambudkar, S. V.
(2001)
J. Biol. Chem.
276,
21199-21208 |
27. |
Daumke, O.,
and Knittler, M. R.
(2001)
Eur. J. Biochem.
268,
4776-4786 |
28. |
Raghuraman, G.,
Lapinski, P. E.,
and Raghavan, M.
(2002)
J. Biol. Chem.
277,
41786-41794 |