From the Division of Microbiology, Department of Genome Sciences, Kobe University Graduate School of Medicine, 7-5-1 Kusunoki-cho, Chuo-ku, Kobe, Hyogo 650-0017, Japan
Received for publication, November 5, 2002, and in revised form, January 17, 2003
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ABSTRACT |
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Kaposi's sarcoma associated-herpes virus
encodes two proteins, MIR (modulator of
immune recognition) 1 and 2, which are involved in the evasion of host immunity. MIR1 and 2 have been shown to function
as an E3 ubiquitin ligase for immune recognition-related molecules
(e.g. major histocompatibility complex class I, B7-2, and
ICAM-1) through the BKS (bovine herpesvirus 4, Kaposi's sarcoma associated-herpes virus, and
Swinepox virus) subclass of plant homeodomain (PHD) domain,
termed the BKS-PHD domain. Here we show that the human genome also
encodes a novel BKS-PHD domain-containing protein that functions as an
E3 ubiquitin ligase and whose putative substrate is the B7-2
co-stimulatory molecule. This novel E3 ubiquitin ligase was designated
as c-MIR (cellular MIR) based on its functional and structural similarity to MIR1 and 2. Forced expression of c-MIR
induced specific down-regulation of B7-2 surface expression through
ubiquitination, rapid endocytosis, and lysosomal degradation of the
target molecule. This specific targeting was dependent upon the binding
of c-MIR to B7-2. Replacing the BKS-PHD domain of MIR1 with the
corresponding domain of c-MIR did not alter MIR1 function. The
discovery of c-MIR, a novel E3 ubiquitin ligase, highlights the
possibility that viral immune regulatory proteins originated in the
host genome and presents unique functions of BKS-PHD domain-containing
proteins in mammals.
Previously, we and other groups found that Kaposi's sarcoma
associated-herpesvirus
(KSHV)1 MIR
(modulator of immune recognition),
MIR2 proteins, and murine The PHD domain has a similar structure to the RING (really
interesting new gene) domain, which
is a functionally critical domain for several E3 ubiquitin ligases such
as c-Cbl and Hakai (10-12). c-Cbl and Hakai induce the ubiquitination
of receptor or nonreceptor tyrosine kinase and E-cadherin, respectively
(10, 12, 13). c-Cbl and Hakai-mediated ubiquitination requires tyrosine
phosphorylation of targets and induces rapid endocytosis and lysosomal
degradation of targets (10, 12, 13). This is the case with MIR1 and
MIR2. The expression of MIR1 or MIR2 induces rapid endocytosis,
translocation to the trans-Golgi network, and lysosomal
degradation of MHC I (1, 9). The BKS-PHD domain of MIR1 has been shown
to be a functionally critical domain for the endocytosis of MHC I (9).
Recently, the PHD domain of mitogen-activated protein
kinase/extracellular signal-regulated kinase kinase kinase 1 has also
been shown to function as an E3 ubiquitin ligase and to induce ERK
degradation in a proteasome-dependent manner (14).
These findings gave rise to the question as to where the viral BKS-PHD
domain-containing proteins are derived from. The genomes of large DNA
viruses including herpesviruses carry many homologs of host cellular
proteins. Likewise, the KSHV genome encodes homologs of many human
genes (e.g. viral interleukin-6 (v-IL-6) and viral macrophage inflammatory protein (v-MIP)), which have been implicated in
viral pathogenesis (15), suggesting that viral BKS-PHD
domain-containing proteins might also have originated in the host
genome. To justify this hypothesis, we searched the human protein data
base of Celera Genomics (16). Bioinformatics and experimental analysis
led us to discover that the human genome contains a gene encoding a
novel BKS-PHD domain-containing protein that functions as an E3
ubiquitin ligase and whose putative substrate is the B7-2
co-stimulatory molecule. Based on its functional and structural
similarity to KSHV MIR1 and MIR2, we have designated this novel E3
ubiquitin ligase as c-MIR (cellular MIR).
Although overall sequence similarity between c-MIR and KSHV MIRs is
low, they share the same secondary structure. Forced expression of
c-MIR causes polyubiquitination, rapid endocytosis, and lysosomal
degradation of B7-2 specifically. This specific targeting is achieved
through binding to the transmembrane and/or cytoplasmic regions of
B7-2. Furthermore, the BKS-PHD domain of c-MIR functioned in the
context of KSHV MIR1. This finding highlights the possibility that
viral immune regulatory proteins originated in the host genome and
shows unique functions of mammalian BKS-PHD domain-containing proteins.
Data Base Searches and Cloning cDNA Encoding Human
c-MIR--
To look for the candidates of KSHV MIR1,2 homolog, the data
base of Celera Genomics was searched using the program BLAST-P, which
allowed a search for patterns in translated nucleic acid sequences data
bases (16). For cloning of the cDNA coding human c-MIR, total RNA
isolated from BJAB cells were reverse-transcribed using the SuperScript
RT kit (Invitrogen) according to the manufacturer's protocol. To
determine the 5' and 3' ends of the entire coding sequence of c-MIR
cDNA, 5' and 3' rapid amplification of cDNA ends were performed
using a GeneRacerTM kit (Invitrogen). A full-length
cDNA of c-MIR was obtained by PCR and subcloned into the pEF-1
vector and GFP co-expression vector pTracer EF-1 (Invitrogen) and
sequenced by a model 310 DNA sequencer (Applied Biosystems). The
putative secondary structure and transmembrane topology of c-MIR were
examined by Profile fed neural network system from HeiDelberg (PHD)
(17-19).
Detection of c-MIR mRNA from Human Tissues and Cultured
Cells--
For the detection of c-MIR mRNA from human tissues,
PCR-ready cDNA kits (Maxim Biotech) were subjected to PCR analysis
with the specific primers, 5'-CGCGAATTCGCCGCCATGAGCATGCCACTG-3'
(forward) and 5'-CGCTCTAGAGACGTGAATGATTTCTGCTCC-3' (reverse) to detect
the full-length human c-MIR mRNA according to the manufacturer's
protocol. For detection of mRNA from cultured cells, total RNA was
extracted by using the RNeasy mini kit (Qiagen), and 2 µg of total
RNA was reverse-transcribed. 400 ng of cDNA was subjected to PCR
analysis as described above. Each PCR product was subcloned into pEF-1 and verified by DNA sequencing.
Generation of Antibody--
Antibody directed against c-MIR was
produced by immunizing rabbits with a synthetic peptide,
DAISARVYRSKTKEKEREE, corresponding to amino acids of 15-33 of c-MIR.
The rabbits were immunized with keyhole limpet hemocyanin-coupled
peptides in complete adjuvant, followed by antisera preparation.
Quantification by Real Time RT-PCR--
A one-step RT-PCR
analysis was performed using the QuantiTectTM SYBR Green
RT-PCR kit (Qiagen). 50 ng of total RNA was analyzed according to the
manufacturer's recommendation. The SYBR Green fluorescence was
measured after each elongation using an ABI PRISM 7000 sequence
detection system (Applied Biosystems). After PCR amplification, a
melting curve analysis was performed by increasing the temperature from
60 to 95 °C. To assess the purity of the amplicons of interest,
RT-PCR products were analyzed by gel electrophoresis. The intensity of
the SYBR Green fluorescent signal was converted into a relative number
of copies of interest based on the results of a series of standards
prepared by successive dilution of total RNA. The expression level of
glyceraldehye-3-phosphate dehydrogenase (GAPDH) was determined for
normalization of the data set. Each experiment was performed twice
using duplicate samples from independently generated cDNA templates.
Plasmid Construction--
B7-2 cDNA was obtained from total
RNA of BJAB cells by RT-PCR and subcloned into pEF-1. HLA-A2 and CD8
cDNA were kindly provided by Dr. G. Cohen (20). Each CD8 chimera
was constructed by overlapping PCR as described previously (2, 21).
Substitutions were engineered into each chimera by PCR-based
mutagenesis (Promega). For analysis by two-color flow cytometry, each
cDNA was subcloned into the GFP co-expression vector pTracer EF-1.
To introduce a FLAG epitope tag, each cDNA was subcloned into
p3XFLAG-CMV vector (Sigma). To construct plasmid DNAs for GST fusion
proteins, a fragment encoding the c-MIR BKS-PHD domain was amplified by
PCR and then subcloned into pGEX 4T-1 (Amersham Biosciences).
Cell Culture and Transfection--
BJAB, 293T, and A7 cells
(ATCC) were grown in RPMI with 10% fetal calf serum or Dulbecco's
modified Eagle's medium with 10% fetal calf serum and minimum
essential medium with 10% fetal calf serum. For transient
assays, expression plasmid DNAs were introduced by electroporation at
260 V and 975 microfarads in serum-free RPMI medium or by transfection
with FuGENE 6 reagent (Roche Molecular Biochemicals). To select stable
transformants, 2 mg/ml of G418 (Sigma) was added 48 h
post-transfection, and the cells were maintained in selective medium
for 3-6 weeks.
Monocytes and Dendritic Cells (DCs)--
Human peripheral blood
mononuclear cells (PBMCs) from healthy volunteers were separated from
peripheral blood by Ficoll-Hypaque centrifugation. The monocytes were
obtained from PBMCs using the MACS monocyte isolation kit (Miltenyi
Biotec), which depletes T cells, NK cells, B cells, and basophils from
PBMC using a mixture of hapten-conjugated CD3, CD7, CD19, CD45RA, CD56,
and anti-IgE antibodies. Immature DCs were generated by culturing
monocytes at a concentration of 1 × 106 cells/ml for
7 days; granulocyte-macrophage colony-stimulating factor (2000 units/ml; R&D Systems) and interleukin-4 (3000 units/ml; R&D Systems)
were added on days 0 and 4. The cell fraction remaining after monocyte
isolation was collected and used as a source of peripheral blood
lymphocytes (PBLs).
Metabolic Labeling, Immunoprecipitation, and PNGase-F
Digestion--
For metabolic labeling, the cells were washed three
times with phosphate-buffered saline (PBS), washed once with labeling medium (RPMI minus methionine and cysteine plus 10% dialyzed fetal calf serum), and then incubated with 5 ml of the same medium containing 50 µCi of [35S]methionine and
[35S]cysteine (PerkinElmer Life Sciences) for 6 h.
For pulse-chase analysis, the cells were labeled for 30 min and chased
for the indicated time. For immunoprecipitation, the cells were
harvested and lysed with lysis buffer (0.15 M NaCl, 1%
Nonidet P-40, and 50 mM HEPES buffer, pH 8.0) containing
protease inhibitors. Immunoprecipitation was performed with the
indicated antibody together with 30 µl of protein A/G-agarose beads
(Santa Cruz). For PNGase-F digestion, washed immunoprecipitates were
resuspended in 50 µl of 1× denaturing buffer (0.5% SDS, 1%
Immunofluorescence Microscopy--
The cells were fixed with 4%
paraformaldehyde PBS for 15 min and cold acetone for 15 min. Fixed
cells were stained with 1:100 diluted primary antibody in PBS for 30 min. After incubation, the cells were washed extensively with PBS and
incubated with a 1:1000 dilution of Alexa 488 or 568-conjugated
secondary antibody (Molecular Probes) in PBS for 30 min. Finally, the
cells were washed three times with PBS and mounted in mounting medium (Vector).
Flow Cytometry Analysis and Antibodies--
The cells (5 × 105) were washed with RPMI medium containing 2% fetal calf
serum and incubated with fluorescein isothiocyanate- or phycoerythrin
(PE)-conjugated monoclonal antibodies for 30 min at 4 °C. After
being washed, each sample was fixed with 2% paraformaldehyde solution,
and flow cytometry analysis was performed with a FACScan (Becton
Dickinson). W6/32 antibody for MHC I, RPA-T8 antibody for CD8, HA 58 antibody for ICAM-1, L307.4 for B7-1, and FUN-1 antibody for B7-2 used
for FACScan were obtained from PharMingen Becton Dickinson Company. The
M2 anti-FLAG antibody (Sigma), F7 anti-HA antibody (Santa Cruz), G-18
anti-His antibody (Santa Cruz), P4D1 anti-ubiquitin antibody (Santa
Cruz), and the anti-V5 antibody (Invitrogen) were used for
immunoprecipitation and/or immunoblot analysis.
Endocytosis Assay--
The experiments were performed as
described previously (2, 9). Briefly, c-MIR cells or control BJAB cells
were stained with PE-labeled anti-B7-2 antibody at 4 °C and then
incubated for various periods of time at 37 °C. The cells were then
washed in an acidic solution to remove uninternalized antibodies,
fixed, and subjected to flow cytometry. For confocal microscopy
analysis, fluorescein isothiocyanate-labeled B7-2 antibody was added to the culture medium of control BJAB cells or c-MIR cells and then incubated for 2 h at 37 °C. After being washed, these cells
were subjected to analysis with a confocal immunofluorescence
microscope (Bio-Rad).
Immunoprecipitation and Immunoblots--
Each plasmid DNA was
transfected into 293T cells using the FuGENE 6 reagent (Roche Molecular
Biochemicals). After 48 h, the transfected cells were harvested,
lysed with Nonidet P-40 buffer, immunoprecipitated as described above,
and subjected to serial immunoblots as indicated in each experiment.
Ubiquitination Assay--
For in vitro
auto-ubiquitination assay, the GST fusion proteins were produced as
follow. Expression of GST fusion proteins was induced by 0.1 mM isopropyl-1- Identification of a Functional Homolog of KSHV MIR1 and MIR2 in the
Human Genome--
We found a BKS-PHD domain-containing putative
protein, hCP36279, that shares the same secondary structure and
the same positioning of BKS-PHD domain with MIR1 and MIR2 in the data
base of Celera Genomics (16). Because amino-terminal sequences of
hCP36279 were missing in the public data base, 5' rapid amplification
of cDNA ends analysis was performed using cap-trapping methods. We determined the full sequence of an hCP36279 cDNA cloned from the BJAB B cell line (Fig. 1A).
Overall hCP36279 has 12 and 18% amino acid identity to MIR1 and MIR2,
respectively. However, within their BKS-PHD domains, hCP36279 exhibits
36 and 42% amino acid identity compared with MIR1 and MIR2,
respectively (Fig. 1B). hCP36279 was predicted to have two
intracytoplasmic regions, helical transmembrane regions separated by an
extra-cytoplasmic domain, and a BKS-PHD domain located in the putative
amino-terminal intracytoplasmic region. The predicted structure of
hCP36279 is depicted schematically, together with those of MIR1 and
MIR2, in Fig. 1C. To examine the functional similarity of
this protein to MIR1 and MIR2, His-tagged protein was transiently
expressed in BJAB cells by electroporation, and the surface expression
of immune recognition-related molecules was analyzed by two-color flow
cytometry. As shown in Fig.
2A, B7-2 surface expression
was significantly down-regulated, but not MHC I and ICAM-1. The
FLAG-tagged version also exhibited the same function (data not shown).
Based on these findings, this protein has been designated as c-MIR. To
confirm these findings, BJAB cell lines were engineered to have stable
and excess expression of c-MIR (c-MIR cells) and examined for surface
expression of immune recognition-related molecules by flow cytometry.
In this experiment, the examination of B7-1 was included to confirm
the target specificity of c-MIR. The stable expression of exogenous c-MIR (His-tagged c-MIR) was confirmed by immunoprecipitation from the
metabolically labeled cell lysate with an anti-His antibody (Fig.
2B, left panel). Real time RT-PCR analysis showed
that the c-MIR expression level of c-MIR cells was 57-fold higher than that of control BJAB cells (Fig. 2B, right
panel). Flow cytometry analysis showed that the surface expression
of B7-2 was specifically down-regulated on c-MIR cells (Fig.
2C).
Detection of c-MIR mRNA and Protein in Human Monocyte-derived
DCs--
As shown in Fig. 2, c-MIR is able to down-regulate the
surface expression of B7-2 in BJAB cells. This finding suggests that c-MIR might be a functional molecule in antigen-presenting cells, because B7-2 is a co-stimulatory molecule for antigen presentation. To
test this hypothesis, the expression profile of c-MIR mRNA in
various tissues was examined by RT-PCR. The primer set used in this
experiment is able to detect full-length c-MIR mRNA (873 bp). As
shown in Fig. 3A, full-length
c-MIR mRNA was detected in neonatal brain, lymph node, spleen, and
placenta. The nucleotide sequence of all PCR products was exactly the
same as that of cDNA derived from BJAB cells. In addition, a band
slightly bigger than 873 bp was detected in neonatal brain. DNA
sequencing analysis revealed that this transcript had additional
sequences within full-length c-MIR mRNA, suggesting that it is an
immature transcript of c-MIR.
Because full-length c-MIR mRNA was detected in lymphoid tissues, we
sought to determine whether DCs, which are potent antigen-presenting cells, express c-MIR mRNA or not. Immature DCs were differentiated from monocytes that had been purified from PMBCs by negative selection with anti-CD3, anti-CD7, anti-CD19, anti-CD45RA, anti-CD56, and anti-IgE. 95% of the cells obtained in this way were
CD14 c-MIR Induces Rapid Endocytosis and Lysosomal Degradation of
B7-2--
To reveal the molecular mechanism of B7-2 down-regulation,
protein synthesis, degradation, and the trafficking of B7-2 were examined. c-MIR cells were pulse-labeled with
[35S]methionine and [35S]cysteine and
chased for the indicated periods of time. At the end of the chase
periods, pulse-labeled proteins were immunoprecipitated with anti-B7-2
or MHC I antibody and analyzed by SDS-PAGE. Because of the high degree
of glycosylation of B7-2, fully matured B7-2 molecules (marked with
four small closed triangles in Fig.
4A) obviously migrated slower
than the immature form (marked with asterisks in Fig.
4A). In contrast, fully glycosylated MHC I migrated less
slowly than the immature form, probably because of the lower degree of
glycosylation. In control cells (Cont), the amount of fully
matured B7-2 did not decrease significantly up to 6 h. In contrast, in c-MIR cells (c-MIR), at 3 and 6 h, the
amount of fully matured B7-2 was significantly reduced compared with
control cells (Fig. 4A). To confirm an enhanced degradation
of B7-2, B7-2 was treated with PNGase-F to remove glycans, and the
treated protein was analyzed by SDS-PAGE. As shown in Fig.
4A, this experiment clearly demonstrated an enhanced
degradation of B7-2 by c-MIR. In addition, at 0 h, there was not a
significant difference of the intensity of bands between control cells
and c-MIR cells, indicating that the down-regulation of B7-2 surface
expression is not due to the inhibition of the protein synthesis.
Consistent with the observation that c-MIR did not down-regulate MHC I
surface expression, degradation of MHC I was not significantly enhanced by c-MIR (Fig. 4A). Rapid degradation of the target molecule
was also observed in MIR1 or MIR2-expressing BJAB cells, and this degradation was shown to take place in lysosome (1, 9). Because of the
structural and functional similarity of c-MIR to MIR1 and MIR2, the
possibility of lysosomal degradation of B7-2 was examined. Treatment of
c-MIR cells with bafilomycin A1, which raises endolysosomal pH through
the inhibition of the vacuolar H+-ATPase, increased the
steady state level of B7-2 protein as judged by immunoblot analysis of
whole cell lysate (Fig. 4B). Furthermore, pulse-chase
analysis was performed to confirm the inhibition of B7-2 degradation by
bafilomycin A1 (Fig. 4C). Control cells and c-MIR cells were
pretreated with bafilomycin A1 for 6 h, pulse-labeled with
[35S]methionine and [35S]cysteine, chased,
and analyzed as in Fig. 4A. Immunoprecipitated B7-2 proteins
were treated with PNGase-F and analyzed by SDS-PAGE. As shown in Fig.
4C, pretreatment with bafilomycin A1 inhibited degradation
of B7-2. These results demonstrate that c-MIR targets B7-2 for
lysosomal degradation.
Next, the possibility of enhanced endocytosis of B7-2 was examined;
MIR1 and MIR2 do not affect the trafficking of target proteins to the
plasma membrane but instead function by altering the endocytic pathway
(1, 9, 21). BJAB cells (Cont) and c-MIR cells
(c-MIR) were stained with PE-conjugated anti-B7-2 antibody
at 4 °C, washed with PBS to remove unbound antibodies, and incubated
in complete RPMI medium at 37 °C for 10 and 30 min. After the
incubation, uninternalized antibodies were removed with acidic
solution, and internalized fluorescence signal was measured by flow
cytometry. Even after incubation for 10 min, significant internalized
fluorescence was observed in c-MIR cells (shown as the shaded
histogram in panel 10 of c-MIR cells in Fig.
5A), but not in the control
cells were observed (shown as the shaded histogram in
panel 10 of the control cells in Fig. 5A). After
incubation for 30 min, there was no obvious increase in internalized
fluorescence in c-MIR cells, and significant internalized signals were
not yet observed in the control cells (Fig. 5A). These
results suggest that c-MIR induces rapid endocytosis of B7-2 that is
probably accomplished within 10 min. The same analysis showed that the endocytosis of MHC I was not enhanced in c-MIR cells (data not shown).
To confirm c-MIR-induced rapid endocytosis of B7-2, we visualized
endocytosed B7-2 molecules by confocal microscopy. The c-MIR cells and
control cells were incubated with fluorescein isothiocyanate-labeled
anti-B7-2 antibody at 37 °C and examined for the localization of
B7-2 molecules. This examination clearly showed internalized B7-2
molecules in the c-MIR cells but not in the control cells (Fig.
5B).
Transmembrane and/or Cytoplasmic Regions of B7-2 Are Involved in
Specific Targeting by c-MIR--
Previously, we demonstrated that the
transmembrane and cytoplasmic regions of target molecules are
sufficient for MIR1 and MIR2-mediated down-regulation by employing CD8
chimeras (2, 21). To see whether the same regions of B7-2 are
sufficient for c-MIR-mediated down-regulation, CD8 chimeras were
constructed with HLA-A2 and B7-2 molecules. CD8/A2 and CD8/B7-2
chimeras contain the transmembrane and cytoplasmic regions of HLA-A2
and B7-2, respectively, fused to the carboxyl terminus of the
extracellular region of CD8 c-MIR Functions as an E3 Ubiquitin Ligase--
c-MIR contains a
BKS-PHD domain similar to MIR1 and MIR2. The BKS-PHD domain of MIR1 and
MIR2 was shown to be a functional domain for E3 ubiquitin ligase (6,
7). To verify the E3 ubiquitin ligase activity of the BKS-PHD domain of
c-MIR, the BKS-PHD domain of MIR1 was replaced with that of c-MIR, and
the resulting chimeric protein, termed MIR1/c-MIR, was subjected to flow cytometry (Fig. 7A).
Another chimera containing a mutated BKS-PHD domain of c-MIR, termed
MIR1/c-MIR(Cys Ubiquitination Is Necessary for c-MIR-mediated Down-regulation of
B7-2 Surface Expression--
We earlier showed c-MIR as a modulator of
B7-2 surface expression and a novel E3 ubiquitin ligase for B7-2.
However, it was still unknown whether or not ubiquitination was
really necessary for down-regulation of B7-2 surface expression.
To clarify this point, c-MIR(Cys Specific Ubiquitination and Down-regulation of B7-2 through
Molecular Interaction--
The possible molecular interaction of c-MIR
and B7-2 was then examined, because E3 ubiquitin ligase has been shown
to function through binding to targets (22, 23). c-MIR and CD8-B7 were expressed together or individually in 293T cells, and these cell lysates were analyzed by immunoprecipitation and immunoblot analysis as
indicated. Only when c-MIR and CD8-B7 were expressed together, a
specific band corresponding to c-MIR was detected, demonstrating the
molecular interaction of both proteins (Fig.
9A). It has been proposed that
the transmembrane regions of MIR1 and MIR2 confer specific targeting
through binding to the target protein transmembrane region (6, 24). If
so, c-MIR(Cys
Next, to confirm the molecular interaction, we examined where they
associate with each other by immunofluorescence microscopy. For this
purpose, we used A7 cells in which the relatively large cytoplasmic
content facilitates the examination of subcellular localization. c-MIR
or c-MIR(Cys
To see whether specific down-regulation of B7-2 is due to molecular
interaction, CD8/A2 used in Fig. 6 was employed for further examination
because the surface expression of CD8/A2 chimera was not down-regulated
in c-MIR cells (Fig. 6B). CD8/A2 was modified to encode a
FLAG epitope tag at its amino terminus, termed CD8-A2, and CD8-A2 or
CD8-B7 was co-expressed with c-MIR(Cys We have identified a functional homolog of KSHV MIR1 and MIR2 in
the human genome. This functional human homolog has been designated as
c-MIR. c-MIR targets the B7-2 co-stimulatory molecule to lysosomal
degradation through the enhancement of endocytosis, eventually leading
to down-regulation of B7-2 surface expression. Moreover, the binding of
c-MIR to B7-2 and the following ubiquitination of the B7-2 cytoplasmic
tail are necessary for down-regulation of B7-2 surface expression.
MIR1, MIR2, and c-MIR share the same secondary structure, a similar
BKS-PHD domain, and the same positioning of the BKS-PHD domain,
suggesting that these molecules belong to the same class of E3
ubiquitin ligase. Although the physiological role of c-MIR is as yet
unknown, our findings suggest some attractive hypotheses that are
discussed below.
The first hypothesis is that c-MIR might be a novel regulator for
antigen presentation because of its unique targeting property. The B7-2
co-stimulatory molecule supports MHC class II-mediated antigen
presentation to effector T cells on the surface of antigen presenting
cells (e.g. dendritic cells and B cells) and has been shown
to be one of the modulators for immune synapse formation (25, 26)
between T cells and antigen presenting cells. The formation of an
immunological synapse has been demonstrated to potentiate T cell
activation (27, 28). Furthermore, co-stimulatory signaling has been
implicated in the progression of autoimmune diseases (29-31). Hence,
c-MIR-mediated modulation of B7-2 surface expression is expected to
regulate the status of T cell activity and immunity. To establish an
animal model and test this attractive hypothesis, we cloned mouse c-MIR
from murine B cell lines. Mouse c-MIR showed exactly the same
function,2 suggesting that
mice also have a similar system. This mRNA was detected by RT-PCR
in most tissues except in the thymus.2 This preliminary
finding may still be consistent with our hypothesis, because B7-2 is
expressed abundantly in the thymus. If c-MIR is expressed abundantly in
the thymus, B7-2 would be expressed inefficiently. A detailed
examination of the profile of tissue distribution and the expression
status of c-MIR in either a physiological or pathological setting and
genetically modified mice are necessary to support our hypothesis.
Another hypothesis is related to the origin of MIR1 and MIR2. The KSHV
genome encodes many homologs of human genes such as viral
interleukin-6, viral macrophage inflammatory protein, and vBcl-2
(15). In Fig. 7B, we showed that the BKS-PHD domain of MIR1
could be replaced with that of c-MIR without alteration of the function
of MIR1. These findings suggest that at least the functional domains of
MIR1 and MIR2 might have originated in the host genome. Moreover, the
homologs of MIR1 and MIR2 have been identified in simian and bovine
At present, several classes of E3 ubiquitin ligase have been discovered
(23). Catalytic domains of E3 ubiquitin ligase are classified into two
major groups: HECT (homologous to E6-AP
carboxyl terminus) or RING domains. HECT and
RING domains are critical for their E3 ubiquitin ligase activities,
because they recruit E2 and transfer ubiquitin to target molecules. In
this study, we showed that a mammalian BKS-PHD domain, which is a
subclass of the PHD domain whose viral version is represented by the
KSHV MIR proteins, was essential for ubiquitination in vivo
and able to function in cooperation with UbcH5a/E2 in vitro
(Fig. 7, C and D). Further, this mammalian
BKS-PHD domain was able to function in the context of KSHV MIR1 (Fig.
7B). Because the sequence of the BKS-PHD domain resembles
the RING domain consensus sequence CX2CX9-39CX1-3HX2-3(C/H)X2CX4-48CX2C, we would like to propose that the BKS-PHD domain is a novel subclass of
RING domain in mammals as well as in viruses. In this regard, the E3
ubiquitin ligase activity of the mammalian PHD domain has been reported
in mitogen-activated protein kinase/extracellular signal-regulated
kinase kinase kinase 1 (14). There are several mammalian BKS-PHD
domain-containing proteins in the public protein data base whose
functions are still unknown. It will be important to analyze these
hypothetical proteins and their targets for E3 ubiquitin ligase activity.
Our findings show that the mechanism of c-MIR-mediated down-regulation
is strikingly similar to that of KSHV MIR1 and MIR2. c-MIR, like MIR1
and MIR2, ubiquitinates a target molecule and leads to rapid
endocytosis and translocation to lysosome, followed by rapid
degradation of a target molecule. So far, they only differ with regard
to their specific target molecules. It will be important to understand
how they specifically target different molecules. In this connection,
it has been reported that an MIR1 chimera containing the transmembrane
domain of MIR2 is able to down-regulate the MIR2 targets B7-2 and
ICAM-1 (24), suggesting that different targeting is achieved by the
binding property of the transmembrane domains of MIR1 and MIR2.
Although both MIR2 and c-MIR were able to target B7-2 (21, 34), there
was no significant homology between their transmembrane domains.
Further detailed analysis is necessary to reveal how these E3 ubiquitin
ligases regulate targeting specificity.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-herpesvirus 68 K3 protein down-regulate
the surface expression of MHC class I (MHC I) on several cell lines
(1-4). Recently, these viral proteins have been shown to ubiquitinate
MHC I (5-7). Notably, MIR1 and MIR2 have been shown to function as
E3 ubiquitin ligase and the BKS (bovine
herpesvirus 4, KSHV, and Swinepox virus) subclass of plant homeodomain (PHD) domain, termed the BKS-PHD domain,
was identified as a catalytic domain of their E3 ubiquitin ligase
activity (6). The BKS-PHD domain was designated based on its existence
in bovine herpesvirus, KSHV, and Swinepox virus genome (8). This domain
was classified as a subclass of the PHD domain because of an interval
difference between the third and fourth cysteine residues in a
zinc-binding motif. The PHD domain has been found in many proteins
involved in chromatin-mediated transcriptional regulation, but their
functions remain unknown. Although KSHV MIRs and murine
-herpesvirus
68 K3 share the same structure and the same positioning of BKS-PHD,
they utilize distinct pathways for degradation of MHC I; KSHV MIRs lead
target molecules to lysosomal degradation (1, 9), whereas murine
-herpesvirus 68 K3 causes proteasomal degradation (3, 5).
Nevertheless, the BKS-PHD domain has been shown to be a critical domain
for ubiquitination and degradation of MHC I in all cases.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-mercaptoethanol), heated for 10 min at 100 °C, and incubated for
1 h at 37 °C with 1 µl of PNGase-F (New England Biolabs).
-D-thio-galactopyranoside for 3-6 h. Bacterial pellets were sonicated in PBS containing protease inhibitors and 0.1% Triton X-100. After being cleared by
centrifugation, bacterial lysates were incubated with
glutathione-Sepharose beads (Amersham Biosciences). 10 µg of
precipitated GST fusion proteins were mixed with 1 µg of UbcH5a
(Boston Biochem), 2 µg of His-tagged ubiquitin (Calbiochem), and 40 ng of E1 (Calbiochem) in the auto-ubiquitination buffer (40 mM Tris-HCl, pH 7.5, 5 mM MgCl2, 2 mM ATP, 2 mM dithiothreitol, 25 µM MG132), incubated at 35 °C for 3 h, and
subjected to immunoblot with anti-ubiquitin antibody.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
hCP36279 is a structural homolog of KSHV MIR1
and MIR2. A, the amino acid sequence of hCP36279 is
shown. Two transmembrane domains (shown as TM1 and
TM2) were predicted by the PHDhtm program.
Underlining indicates consensus residues of the BKS-PHD
domain. B, the alignment of protein sequences from hCP36279,
MIR1, and MIR2 within their BKS-PHD domains. The putative zinc-binding
residues are marked by asterisks. C, a putative
molecular structure and transmembrane topology of hCP36279 are shown
together with MIR1 and MIR2. The BKS-PHD domain is shown as
PHD. The putative secondary structure and transmembrane
topology of c-MIR were determined by using Profile fed neural network
system from HeiDelberg (17-19).
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Fig. 2.
hCP36279, renamed as c-MIR, is a functional
homolog of KSHV MIR1 and MIR2. A, BJAB cells were
transfected with the GFP-c-MIR vector by electroporation. The surface
expression of each molecule and the expression of GFP were analyzed
24 h post-transfection by two-color flow cytometry as in previous
experiments (2, 9). In brief, the cells (5 × 105)
were washed with RPMI medium containing 2% fetal calf serum and
incubated with the indicated PE-conjugated monoclonal antibodies for 30 min at 4 °C. After being washed, each sample was fixed with 2%
paraformaldehyde solution, and flow cytometry analysis was performed
with a FACScan (Becton Dickinson). The y axis shows the
level of surface expression of MHC I, ICAM-1, and B7-2, and the
x axis shows the level of GFP expression. B, the
left panel shows the stable expression of exogenous c-MIR
protein in c-MIR cells. c-MIR cells and control BJAB cells were labeled
with [35S]methionine and [35S]cysteine for
6 h, and metabolically labeled cell lysates were
immunoprecipitated with anti-His polyclonal antibody.
Immunoprecipitates were analyzed by SDS-PAGE. In the right
panel, the level of c-MIR mRNA was determined by using
quantitative RT-PCR in control BJAB cells and c-MIR cells. The
expression level of c-MIR was normalized to GAPDH expression and is
indicated relative to the expression level in control BJAB cells. All
of the reactions were performed in duplicate, and the standard
deviation is indicated. C, BJAB cells stably overexpressing
c-MIR (c-MIR) and control BJAB cells were stained with the
indicated antibodies as described for A, and the surface
expression of the indicated molecules was analyzed by flow cytometry.
In merged panels, the bold lines and shaded
histograms show the results of c-MIR cells and control BJAB cells,
respectively. The results of the staining with isotype control antibody
are shown as dotted lines.
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Fig. 3.
Expression of c-MIR. A, 20 ng
of cDNA from neonatal brain, lymph node, heart, lung, liver,
spleen, kidney, and placenta were PCR-amplified using the primers
described under "Experimental Procedures" to detect full-length
c-MIR mRNA (upper panel). Also the expression of GAPDH
mRNA was measured (lower panel). B, DCs,
monocytes and PBLs were prepared from human PBMCs as described under
"Experimental Procedures." Total RNA from these cells was
reverse-transcribed and PCR-amplified as performed in A
(upper panel). The lower panel shows the
expression of GAPDH mRNA. C, DCs, c-MIR-expressed 293T
(293T+c-MIR), and 293T were labeled with
[35S]methionine and [35S]cysteine for
6 h. Each labeled cell extract was immunoprecipitated with
anti-c-MIR polyclonal antibody ( -c-MIR) or preimmune
serum (Cont). Each precipitated protein sample was analyzed
by SDS-PAGE. The bands corresponding to the authentic c-MIR
protein are marked with asterisks.
CD11c+ (data not shown). The remaining
cell fraction from monocyte isolation was collected as PBLs. Immature
DCs, monocytes, and PBLs were subjected to RT-PCR analysis. As shown in
Fig. 3B, c-MIR mRNA was detected in monocytes and DCs
but not in PBLs. The intensity of PCR band of DCs was stronger than
that of monocytes, suggesting that DCs express c-MIR mRNA more than
monocytes. To confirm that DCs express the authentic c-MIR protein,
rabbit polyclonal antibody directed against c-MIR was generated. 1 × 107 immature DCs were labeled with
[35S]methionine and [35S]cysteine for 6h,
and metabolically labeled cell lysates were immunoprecipitated with
anti-c-MIR polyclonal antibody (
-c-MIR in Fig.
3C) or rabbit preimmune serum (Cont in Fig.
3C). To verify the full size of the authentic c-MIR protein,
the epitope tag sequences (e.g. V5 and His tag) were deleted
from the c-MIR expression plasmid, and the resulting expression plasmid
was transfected into 293T cells. Transfected 293T cells
(293T+c-MIR in Fig. 3C) or nontransfected 293T
cells (293T in Fig. 3C) were labeled with [35S]methionine and [35S]cysteine and
immunoprecipitated with anti-c-MIR antibody or preimmune serum. As
shown in Fig. 3C, this antibody was able to precipitate the
proteins of the same molecular weight (marked with asterisks
in Fig. 3C) from cell lysates of DCs and transfected 293T
cells, but not from nontransfected 293T cell lysates. On the other
hand, with preimmune serum, a band corresponding to c-MIR was not
detected from any cell lysates. These results indicate that immature
DCs express the authentic c-MIR protein. Taken together, these findings
suggest that c-MIR might be a functional molecule in DCs.
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Fig. 4.
c-MIR targets B7-2 to lysosomal
degradation. A, BJAB cells (Cont) and c-MIR
cells (c-MIR) were pulse-labeled with
[35S]methionine and [35S]cysteine for 30 min and chased for 1-6 h. At the end of the chase periods, the cells
were lysed and immunoprecipitated with anti-B7-2 or anti-MHC I
antibody. Each precipitated protein sample was analyzed by SDS-PAGE
after treatment with PNGase-F (+) or not ( ). B, c-MIR
cells were treated with 10 µM bafilomycin A1, an
inhibitor of lysosomal degradation, for 2-4 h. At the end of
treatment, the cells were subjected to immunoblot analysis with an
anti-B7-2 antibody (upper panel) or anti-actin antibody
(lower panel). C, c-MIR cells were treated with
10 µM bafilomycin A1 (shown as +) or Me2SO as
control (shown as
) for 6 h and subjected to pulse-chase
analysis as in Fig. 3A. At the end of the chase periods, the
cells were lysed and immunoprecipitated with anti-B7-2, followed by
SDS-PAGE analysis after treatment with PNGase-F.
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Fig. 5.
c-MIR induces rapid endocytosis of B7-2.
A, BJAB cells (Cont) and c-MIR cells
(c-MIR) were stained with PE-conjugated anti-B7-2 antibody
at 4 °C, washed with PBS to remove unbound antibodies, and incubated
in complete RPMI medium at 37 °C for 10 and 30 min. After the
incubation, uninternalized antibodies were removed with acidic
solution, and internalized fluorescence was measured by flow cytometry.
These results are shown as shaded histograms in panels
labeled with 10 and 30, respectively, to their left. To
determine the background level of fluorescence, just after being
stained with PE-conjugated anti-B7-2 antibody, cell surface-associated
antibodies were removed with acidic solution, and a background level
was measured by flow cytometry. These results are shown as bold
lines in panels labeled 10 or 30. The panels
labeled Pre show the level of B7-2 surface expression before
stripping of surface-associated antibodies. The results of the staining
with isotype control antibody are shown as dotted lines.
B, BJAB cells (Cont) and c-MIR cells
(c-MIR) were incubated with fluorescein
isothiocyanate-labeled anti-B7-2 antibody at 37 °C for 2 h, and
then internalized B7-2 molecules were observed with confocal
microscopy.
(Fig.
6A). These CD8 chimeras were
expressed in control BJAB or c-MIR cells, and CD8 surface expression
was examined by flow cytometry. Because there is no CD8 expression on
the surface of BJAB cells, this approach gives a clear signal for
detecting down-regulation of the target molecule. Both CD8 chimeras
were expressed efficiently on the surface of control BJAB cells (Fig. 5). On the other hand, whereas CD8/B7-2 chimera was not expressed efficiently on the surface of c-MIR cells, the CD8/A2 chimera was (Fig.
6B). This result confirms the specific targeting by c-MIR
and shows that transmembrane and/or cytoplasmic regions of the target
molecule are involved in this specificity.
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Fig. 6.
Transmembrane and/or cytoplasmic regions of
B7-2 are involved in c-MIR-mediated down-regulation. A,
schematic representation of the structure of CD8 chimeras used in this
experiment. Extracellular domain (Extra), transmembrane
domain (Tm), cytoplasmic domain (Cyto) are
indicated. B, BJAB cells (Cont) and c-MIR cells
(c-MIR) were transiently transfected with GFP-CD8/A2 or
GFP-CD8/B7-2 vector by electroporation. After 24 h, the
transfected cells were stained with PE-labeled anti-CD8 antibody, and
the surface expression of CD8 chimeras and the expression of GFP were
analyzed by two-color flow cytometry. The y axis shows the
surface expression of CD8, and the x axis shows the
expression of GFP.
Ser), was also included (Fig. 7A). In
this mutant, cysteines at positions 80, 83, 123, and 125, which are
putative zinc-binding residues, were mutated to serine. These chimeras
and MIR1 were expressed in A7 cells, and MHC I surface expression was
examined by two-color flow cytometry. As expected, MIR1/c-MIR
down-regulated MHC I surface expression efficiently (Fig.
7B). This down-regulation was also dependent on BKS-PHD
domain; MIR1/c-MIR(Cys
Ser) was not able to down-regulate the
surface expression of MHC I. These results strongly suggest that c-MIR
has E3 ubiquitin ligase activity. To confirm this, an in
vitro auto-ubiquitination assay was performed. The wild or mutant
type of the BKS-PHD domain of c-MIR was fused to the carboxyl terminus
of GST, and purified GST fusion proteins were subjected to
auto-ubiquitination assay in a mixture of ATP, free ubiquitin, E1, and
E2 (UbcH5a). A clear polyubiquitinated form of the wild type BKS-PHD
domain-containing GST fusion protein (GST-c-MIR) was observed, but GST
alone (GST) or mutated BKS-PHD domain-containing GST fusion protein
(GST-c-MIR(Cys
Ser)) did not have this form (Fig. 7C).
GST-c-MIR(Cys
Ser) has the same mutation as
MIR1/c-MIR(Cys
Ser) in the BKS-PHD domain. This result confirmed
the E3 ubiquitin ligase activity of the BKS-PHD domain of c-MIR. To
test whether B7-2 can be ubiquitinated by c-MIR, B7-2 was co-expressed
with HA-tagged ubiquitin and either wild type c-MIR (c-MIR) or mutant
type c-MIR whose BKS-PHD domain was mutated (termed
c-MIR(Cys
Ser)) as done in the case of MIR1/c-MIR(Cys
Ser).
These cell lysates were precipitated with anti-B7-2 antibody and
subjected to immunoblot analysis with anti-HA or B7-2 antibody. With
co-expression of c-MIR, a clear polyubiquitinated form of B7-2 was
detected by HA and B7-2 blots but not without expression of c-MIR
(Cont) or with co-expression of c-MIR(Cys
Ser) (Fig. 7D). Taken together, these findings confirm that c-MIR
functions as a E3 ubiquitin ligase for the B7-2 co-stimulatory molecule through its BKS-PHD domain.
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Fig. 7.
c-MIR is an E3 ubiquitin ligase for
B7-2. A, schematic representation of the structure of
MIR-1 and its chimeras used in this experiment. The BKS-PHD domain of
MIR1 (8-57 amino acids) was replaced by that of c-MIR (78-137 amino
acids) to construct MIR1/c-MIR. In addition, cysteines 80, 83, 123, and
126 in the BKS-PHD domain of c-MIR were mutated to serine and also used
to replace the c-MIR BKS-PHD domain as above. This chimera was named
MIR1/c-NIR(Cys Ser). Transmembrane domain (Tm) and
BKS-PHD domain are indicated. CC CC HC CC is the consensus
amino acid sequence of the BKS-PHD domain. B, GFP-MIR1,
GFP-MIR1/c-MIR and GFP-MIR1/c-MIR(Cys
Ser) vectors were
transfected in A7 cells, and surface expression of MHC I was analyzed
as in Fig. 2A. C, the BKS-PHD domain of c-MIR
(78-137 amino acids) was fused to GST and subjected to in
vitro auto-ubiquitination assay. Mutant BKS-PHD domain that has
the same mutations as MIR1/c-MIR(Cys
Ser) was also included. GST
alone (GST), the GST-BKS-PHD domain of c-MIR
(GST-c-MIR), and the GST-mutant BKS-PHD domain of c-MIR
(GST-c-MIR(cys>ser)) were incubated in ubiquitination
buffer (details under "Experimental Procedures") for 3 h. The
incubated samples were probed with anti-ubiquitin antibody
(
-Ubi). D, 293T cells were co-transfected with
HA-tagged ubiquitin expression plasmid, pEF-B7-2, and one of the
following plasmid DNAs: pEF (Cont), pEF-c-MIR
(c-MIR), and pEF-c-MIR(Cys
Ser)
(c-MIR(cys>ser)). c-MIR(Cys
Ser) has the same
mutations as MIR1/c-MIR(Cys
Ser) shown in A. Each
co-expressed protein was immunoprecipitated with anti-B7-2
(top and middle panels) and probed with anti-HA
antibody (top panel) or anti-B7-2 antibody (middle
panel). Whole cell lysates were probed with anti-His antibody to
show the comparable expression of c-MIR or c-MIR(Cys
Ser) in each
cell lysate (bottom panel). IP,
immunoprecipitation; IB, immunoblot.
Ser), a ubiquitination dead
mutant, was transiently expressed in BJAB cells by electroporation and
examined by two-color flow cytometry. As shown in Fig.
8B, c-MIR(Cys
Ser) did
not down-regulate the surface expression of B7-2 at all. We
subsequently constructed a mutant B7-2 molecule that was no longer
ubiquitinated by c-MIR. It has been shown that MIR2 mediates the
ubiquitination of lysine residues located at the cytoplasmic tail of
targets (6). This report led us to test whether this is the case with c-MIR. For this examination, the CD8 chimera used in Fig. 6 was employed. All of the lysine residues located at the B7-2 cytoplasmic region of the CD8/B7-2 chimera were mutated to arginine by PCR-based mutagenesis and modified to encode a FLAG epitope tag at its amino terminus. This chimera was termed CD8-B7(Lys
Arg). Furthermore, the CD8/B7-2 chimera was modified to encode a FLAG epitope tag at its
amino terminus, and the resulting modified chimeric protein was termed
CD8-B7. Each FLAG-tagged CD8 chimera was co-expressed with HA-tagged
ubiquitin and c-MIR in 293T cells and subjected to the same experiment
as performed in Fig. 7D. HA and FLAG blots clearly showed
that CD8-B7(Lys
Arg) was no longer ubiquitinated by c-MIR (Fig.
8C). To see whether ubiquitination is linked to down-regulation of target surface expression, CD8-B7(Lys
Arg) and
CD8-B7 were expressed in BJAB cells (control cells) or c-MIR cells, and
the surface expression level of CD8 was compared. Flow cytometry
analysis showed that the expression of CD8-B7(Lys
Arg) was not
inhibited in c-MIR cells (Fig. 8D), suggesting that the
ubiquitination of cytoplasmic lysine residues of B7-2 is necessary for
down-regulation.
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Fig. 8.
Ubiquitination is necessary for the
down-regulation of B7-2 surface expression. A,
schematic representation of the structure of c-MIR and its mutant used
in this experiment. B, BJAB cells were transfected with
GFP-c-MIR or GFP-c-MIR(Cys Ser) vector, and the surface
expression of B7-2 and the expression of GFP were analyzed by two-color
flow cytometry. C, all of the lysine residues in the
cytoplasmic tail of CD8/B7-2 were mutated to arginine by PCR-based
mutagenesis to construct CD8/B7(Lys
Arg). CD8/B7-2 and
CD8/B7(Lys
Arg) were modified to encode a FLAG epitope tag at
their amino termini and termed CD8-B7 and CD8-B7(Lys
Arg),
respectively. CD8-B7-2 and CD8-B7(Lys
Arg) were co-expressed in
293T cells with HA-tagged ubiquitin and c-MIR, and their ubiquitination
status was analyzed as in Fig. 7D. Each expressed protein
was immunoprecipitated with anti-FLAG antibody and probed with anti-HA
(left upper panel) or with anti-FLAG antibody (right
upper and lower panels). The right upper and
lower panels show the results of the same membrane, which
was probed with anti-FLAG antibody. The upper part was
exposed for a longer time than the lower part because of the
weak signal of ubiquitinated FLAG-CD8 chimera (shown as ub-FLAG-CD8
chimera). To show the same expression of c-MIR, each whole cell lysate
was probed with anti-His antibody (left lower panel).
D, BJAB cells (Cont) and c-MIR cells
(c-MIR) were transfected with GFP-CD8-B7 or
GFP-CD8-B7(Lys
Arg) vector, and the surface expression of CD8 and
the expression of GFP were analyzed by two-color flow cytometry.
IP, immunoprecipitation; IB, immunoblot.
Ser), which has intact transmembrane domains, is
expected to be able to associate with the target. To test this
hypothesis, c-MIR(Cys
Ser) was subjected to the same assay as
performed in Fig. 9A. As shown in Fig. 9B, a
clear band corresponding to c-MIR(Cys
Ser) was detected only when
both c-MIR(Cys
Ser) and CD8-B7 were expressed. These results indicate that both c-MIR and c-MIR(Cys
Ser) were able to bind to
CD8-B7.
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Fig. 9.
Molecular interaction of c-MIR and
CD8-B7. A, c-MIR and CD8-B7 were expressed in 293T
cells either together or individually as indicated. The amount of each
transfected DNA was adjusted by adding empty vector DNA. 48 h
after transfection, the expressed proteins were immunoprecipitated with
anti-FLAG antibody and probed with anti-V5 antibody that recognizes
c-MIR (top panel). To show equal expression of CD8 chimeras,
the same precipitated samples were probed with anti-FLAG antibody
(middle panel). Whole cell lysate was probed with anti-V5
antibody to show equal expression of c-MIR. B, the same
analysis was performed by using c-MIR(Cys Ser). C,
CD8-B7 was co-expressed with c-MIR or c-MIR(Cys
Ser) in A7 cells.
CD8-B7 was visualized by staining with anti-CD8 monoclonal antibody and
Alexa 568-conjugated secondary antibody. c-MIR and c-MIR(Cys
Ser)
were visualized by staining with anti-His polyclonal antibody and Alexa
488-conjugated secondary antibody. IP, immunoprecipitation;
IB, immunoblot.
Ser) was co-expressed in A7 cells with CD8-B7 and
stained with anti-His antibody for c-MIR and c-MIR(Cys
Ser) and
with anti-CD8 antibody for CD8-B7. As shown in Fig. 9C, both
c-MIR and c-MIR(Cys
Ser) were co-localized with CD-B7. c-MIR was
localized mainly in the perinuclear region. On the other hand,
c-MIR(Cys
Ser) was localized in both the plasma membrane and
perinuclear regions. These results confirmed the molecular interaction
of B7-2 with c-MIR or c-MIR(Cys
Ser). During these experiments,
we found that a complex of c-MIR(Cys
Ser) and CD8-B7 was more
easily detected than that of wt-c-MIR and CD8-B7, probably because of
the instability of wt-c-MIR/CD8-B7 complex. Consistent with this
explanation, the amount of the high molecular form of CD8-B7, a
probably matured form, was drastically reduced by co-expression of
c-MIR compared with the single expression of CD8-B7 (Fig.
9A), and the intensity of plasma membrane staining of CD8-B7
was drastically reduced by co-expression of c-MIR (Fig. 9C).
Ser) in 293T cells, and
the possible interaction was examined by immunoprecipitation and
immunoblotting analysis. In this study, we employed
c-MIR(Cys
Ser) because of easy detection of c-MIR-CD-8-B7
complexes. As shown in Fig.
10A, c-MIR(Cys
Ser)
was efficiently co-precipitated with CD8-B7, but not with CD8-A2,
suggesting that molecular interaction is necessary for specific
down-regulation by c-MIR. To clarify whether the molecular interaction
is linked to specific ubiquitination of targets, CD8-A2 and CD8-B7 were
subjected to the same analysis as performed in Fig. 8C. As
shown in Fig. 10B, CD8-B7, but not CD8-A2, was efficiently
ubiquitinated. These results demonstrate that a molecular interaction
is necessary for c-MIR-mediated ubiquitination and the following
down-regulation of the target molecule.
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Fig. 10.
Specific ubiquitination through binding to
the target. A, CD8/A2 used in Fig. 5 was tagged with FLAG
epitope at its amino terminus. FLAG-CD8-A2 (CD8-A2) and FLAG-CD8-B7-2
(CD8-B7) were co-expressed with c-MIR(Cys Ser) in 293T cells.
Cell lysate was immunoprecipitated with anti-HA or FLAG antibody as
indicated and probed with anti-V5 (top panel) or anti-FLAG
antibody (middle panel). Each whole cell lysate was probed
with anti-V5 antibody (bottom panel). B, CD8-A2
or CD8-B7 was co-expressed in A7 cells with HA-tagged ubiquitin and
c-MIR, and the ubiquitination status of each molecule was analyzed as
in Fig. 8C. IP, immunoprecipitation;
IB, immunoblot.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-herpesviruses and in swinepox virus (8), suggesting that these
proteins might be derived from a common host predating divergence of
these species. This hypothesis might be extended to other viral immune
regulatory proteins (e.g. US2,11 of human cytomegalovirus).
Herpesviruses have many immune regulatory proteins that down-regulate
MHC I surface expression (32, 33). So far, there are no reports describing functional host homologs of these proteins. In the case of
MIR1 and MIR2, the existence of a unique functional domain, the BKS-PHD
domain, made it easy to identify functional homolog candidates. For
other viral immune regulatory proteins, however, this might prove
difficult because there are no well known domains in these proteins so far.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank J. U. Jung for sharing
materials, R. E. Means for technical advice, D. Bohmann for the
HA-Ubi expression plasmid, G. B. Cohen for CD8 cDNA and
HLA-A2 cDNA, and W. E. Johnson and R. H. Florese for
critical reading of this manuscript.
![]() |
FOOTNOTES |
---|
* This work was supported in part by grants-in-aid for Scientific Research from the Ministry of Education, Science, Sports, and Culture of Japan (to S. I.) and also by grants from the Uehara Memorial Foundation and the Hyogo Science and Technology Association (to S. I.).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 all correspondence should be addressed. Tel.: 81-78-382-5501; Fax: 81-78-382-5519; E-mail: ishido@med.kobe-u.ac.jp.
Published, JBC Papers in Press, February 11, 2003, DOI 10.1074/jbc.M211285200
2 E. Goto, S. Ishido, Y. Sato, S. Ohgimoto, K. Ohgimoto, M. Nagano-Fujii, and H. Hotta, unpublished data.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: KSHV, Kaposi's sarcoma-associated herpesvirus; PHD, plant homeodomain; MHC, major histocompatibility complex; HLA, human leukocyte antigen; PNGase, peptide N-glycanase; HA, hemagglutinin; GST, glutathione S-transferase; ICAM-1, intercellular adhesion molecule 1; E1, ubiquitin-activating enzyme; E2, ubiquitin carrier protein; E3, ubiquitin-protein isopeptide ligase; RT, reverse transcriptase; GAPDH, glyceraldehye-3-phosphate dehydrogenase; GFP, green fluorescent protein; DC, dendritic cell; PBMC, peripheral blood mononuclear cell; PBL, peripheral blood lymphocyte; PBS, phosphate-buffered saline; PE, phycoerythrin.
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REFERENCES |
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