Control of AMP deaminase 1 binding to myosin heavy
chain
Ichiro
Hisatome1,
Takayuki
Morisaki1,
Hiroshi
Kamma2,
Takako
Sugama1,
Hiroko
Morisaki1,
Akira
Ohtahara3, and
Edward W.
Holmes1
1 Departments of Medicine,
Genetics, and Biochemistry, University of Pennsylvania School of
Medicine, Philadelphia 19104-4283;
2 Howard Hughes Medical Institute
and Department of Biochemistry and Biophysics, University of
Pennsylvania School of Medicine, Philadelphia, Pennsylvania
19104-6148; and 3 The 1st
Department of Medicine, Tottori University Faculty of Medicine,
Yonago 683, Japan
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ABSTRACT |
AMP deaminase (AMPD) plays a central role in preserving the
adenylate energy charge in myocytes following exercise and in producing
intermediates for the citric acid cycle in muscle. Prior studies have
demonstrated that AMPD1 binds to myosin heavy chain (MHC)
in vitro; binding to the myofibril varies with the state of muscle
contraction in vivo, and binding of AMPD1 to MHC is required for
activation of this enzyme in myocytes. The present study has identified
three domains in AMPD1 that influence binding of this enzyme to MHC
using a cotransfection model that permits assessment of mutations
introduced into the AMPD1 peptide. One domain that encompasses residues
178-333 of this 727-amino acid peptide is essential for binding of
AMPD1 to MHC. This region of AMPD1 shares sequence similarity with
several regions of titin, another MHC binding protein. Two additional
domains regulate binding of this peptide to MHC in response to
intracellular and extracellular signals. A nucleotide binding site,
which is located at residues 660-674, controls binding of AMPD1 to
MHC in response to changes in intracellular ATP concentration. Deletion
analyses demonstrate that the amino-terminal 65 residues of AMPD1 play
a critical role in modulating the sensitivity to ATP-induced inhibition
of MHC binding. Alternative splicing of the AMPD1 gene product, which alters the sequence of residues 8-12, produces two AMPD1 isoforms that exhibit different MHC binding properties in the presence of ATP.
These findings are discussed in the context of the various roles
proposed for AMPD in energy production in the myocyte.
purine nucleotide cycle; adenylate energy charge; skeletal muscle
bioenergetics
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INTRODUCTION |
ADENOSINE MONOPHOSPHATE DEAMINASE (AMPD) plays a
central role in energy metabolism in eukaryotes (24). This enzyme
activity is found in all eukaryotes, from simple single cell organisms, such as yeast, to man, and it is present in all organs and tissues of
vertebrates. In higher eukaryotes, this enzyme is encoded by a
multigene family, the members of which exhibit tissue-specific and
stage-specific control of gene expression (19). The importance of this
enzyme activity in tissues routinely subjected to stress, such as
skeletal muscle, is illustrated by the myopathy that develops in
patients with reduced amounts of the AMPD1 gene product in skeletal
muscle, the predominant AMPD isoform in adult muscle (7, 17, 27).
AMPD is an allosteric enzyme, and in solution its activity is
controlled by a number of intracellular metabolites, principally purine
nucleotides and inorganic phosphate (2, 5, 20, 21, 33, 34). The
activity of this enzyme is governed in a physiological sense by the
adenylate energy charge of the cell (24), and it can be viewed as a
sensor of the energy needs of the cell. When AMPD is activated and AMP
is deaminated to IMP, the enzyme shifts the equilibrium of the
adenylate kinase reaction, thereby increasing the adenylate energy
charge (24). Activation of AMPD and the resultant increase in adenylate
charge help to preserve viability of the cell under stressful
conditions.
Recent studies have identified a novel mechanism for regulating the
activity of AMPD in myocytes. In resting muscle, more than 90% of this
enzyme is free in the sarcoplasm, and when free in the sarcoplasm the
enzyme is inactive. After vigorous muscle contraction, 50-60% of
this enzyme becomes bound to the myofibril (22, 23), and the activity
of the bound enzyme increases as a consequence of a decrease in
Michaelis-Menten constant for AMP and a decrease in inhibition by
nucleotides and inorganic phosphate (22, 23). These studies have
provided convincing evidence that the enzyme cannot be activated under
the conditions existent in maximally stressed myocytes when it is not
bound to the myofibril (22, 23).
The present study was undertaken to define the properties of the AMPD1
peptide that control binding of this isoform to the myofibril. AMPD1
has been demonstrated to bind to myosin heavy chain (MHC) in vitro, in
particular the S2 subregion of MHC (1, 3, 9, 10). Presumably, this is
the target of binding for this enzyme in the myofibril, but other
macromolecules have also been proposed as binding sites for AMPD1 in
the myofibril (11). Experiments reported here have taken advantage of a
model in which expression of MHC in nonmyocytes leads to formation of
filamentous structures that can be identified immunohistochemically
(29, 30). Through cotransfection of epitope-tagged AMPD1 expression vectors containing deletion or point mutations in the AMPD1 peptide, this system has been exploited to assess binding of wild-type and
mutant AMPD peptides to the myosin filament in vivo. This model has the
advantage that other components of the myofibril are not present,
excluding AMPD1 binding to other proteins that are present in the
sarcomere. This approach is superior to in vitro binding experiments,
since cell lysis and extraction of AMPD1 uniformly lead to proteolysis
with a resultant change in binding properties of the enzyme (13).
Results of the studies reported here demonstrate that at least three
regions of the AMPD1 peptide influence binding of this enzyme to MHC.
The central region of the native 727-amino acid peptide contains a
myosin binding site (residues 178-333). The ATP binding site
(residues 660-674) located in the carboxy terminus regulates
binding in response to changes in the intracellular concentration of
this nucleotide. The amino-terminal 65 residues of the AMPD1 peptide
are essential to the intact molecule for mediation of
ATP-induced inhibition of binding to MHC. The AMPD1 primary
transcript is subject to tissue-specific and stage-specific alternative
splicing in this region of the AMPD1 peptide, generating isoforms that
contain exon 2-encoded sequences (E2+) or that exclude sequences
encoded by exon 2 (E2
) (16). Thus alternative splicing of the
AMPD1 primary transcript can eliminate four amino acid residues from
the amino-terminal region of the AMPD1 peptide (16, 18), and this
physiological deletion makes the AMPD1 peptide less sensitive to
ATP-induced inhibition of binding to MHC under certain conditions.
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MATERIALS AND METHODS |
Plasmid constructs.
pRC/CMV (Invitrogen) was used as the expression vector for both the
AMPD1 and MHC cDNAs. The rat and human AMPD1 cDNAs used in this study
have been described previously (25, 26). A full length of the rat
cardiac MHC-
cDNA was provided by Dr. Lee Sweeney of the Physiology
Department at the University of Pennsylvania. Deletions were made in
the rat AMPD1 cDNA using standard restriction enzyme digestion
techniques, and point mutations were introduced into the human AMPD1
cDNA as previously described (8). Decisions for selection of
restriction sites to create the various deletion mutants were based on
availability of convenient cloning sites that removed reasonable
stretches of amino acid residues from the amino and carboxy termini.
Rat/human chimeras were made by ligating the 5' end of rat AMPD1
cDNA to the 3' end of human AMPD1 cDNA. These chimeras were
created to take advantage of the availability of ATP and
catalytic site point mutations created previously in human AMPD1 (8). A
doublestranded oligonucleotide encoding the human
c-myc epitope, MEQKLISEEDL,
was added to the amino terminus of all AMPD1 constructs. This epitope
is recognized by an anti-Myc monoclonal antibody (MAb), 9E10 (6). It
has the advantage of giving a low immunofluorescent background in
untransfected cells or cells transfected with constructs that do not
contain this epitope. All constructs were sequenced to confirm their
primary structure.
Cell culture and transfection conditions.
Human HeLa cells and Monkey kidney COS-1 cells were routinely grown in
DMEM (GIBCO) supplemented with 10% fetal bovine serum (FBS; GIBCO).
Cells grown on glass coverslips in 30-mm dishes were transfected with 5 µg of a single expression plasmid or 5 µg of both plasmids for
cotransfection studies, using a calcium phosphate coprecipitation
technique (28). Plasmid DNA was added to HEPES-buffered saline composed
of 140 mM NaCl, 1.5 mM
Na2HPO4, and 25 mM HEPES, pH adjusted to 7.05 with HCl, and precipitated by
adding CaCl2 with constant
stirring. After the COS cells were incubated with the coprecipitate for
4 h and washed with PBS, cells were cultured for 44 h before analysis
by immunofluorescence microscopy. After the HeLa cells were incubated
with the coprecipitate for 24 h and washed with PBS, cells were
cultured for 24 h before analysis by immunofluorescence microscopy.
Cell permeabilization.
HeLa cells were permeabilized using a Trans Port Kit from GIBCO using
conditions recommended by the manufacturer (GIBCO). Permeabilized cells
were incubated for 60 min with cytosolic solution (in mM: 140 KCl, 0.38 K2HPO4,
2.13 K2HPO4,
1 MgSO4, and 0.1 CaCl2, pH adjusted to 7.2 with
KOH) containing 0-10 mM MgATP. The metabolic inhibitors
2,4-dinitrophenol (50 mM) and 2-deoxyglucose (10 mM) were also included
in the 0 mM MgATP cytosolic solution ("low ATP"). For solutions
with ATP concentrations higher than ambient ATP concentration ("high
ATP"), 10 mM MgATP was added to the cytosolic solution. Companion
experiments with Trypan blue (molecular weight 1,000) added to the
cytosolic solution confirmed that >90% of the HeLa cells were
permeabilized under the conditions used in this study.
Immunofluorescence microscopy.
For indirect immunofluorescence, cells were fixed with 2% formaldehyde
(Polyscience) in PBS for 30 min at room temperature followed by a 3-min
incubation in cold acetone at
20°C. Fixed cells were
incubated for 1 h at room temperature with either anti-Myc MAb (9E10
culture supernatant, an IgG1 class
antibody; American Type Culture Collection) or anti-sarcomeric MHC MAb
(MF20 culture supernatant, an
IgG2b class antibody; Hybridoma
Bank of University of Iowa). Cells were washed three times
with PBS and incubated for 1 h at room temperature with either 1:500
dilution of FITC-conjugated anti-mouse
IgG1 antibody in PBS containing
3% BSA for Myc staining or Texas red (TXRD)-conjugated anti-mouse
IgG2b antibody in PBS with 3% BSA
for MHC staining. After coverslips were washed three times with PBS,
they were inverted and mounted on glass microscope slides. For double
staining, both anti-Myc MAb and anti-sarcomeric MHC MAb were applied
simultaneously to cells for 1 h. After cotransfectants were washed
three times with PBS, they were incubated with a 1:12.5 dilution of
FITC-conjugated anti-mouse IgG1
antibody (Southern Medical Association) and a 1:25 solution of
TXRD-conjugated anti-mouse IgG2b
antibody (Southern Medical Association) in PBS with 3% BSA.
Immunoblots of transfected cells.
A confluent 30-mm petri dish of HeLa cells was transfected with 10 µg
of plasmid DNA as described above. Cells were harvested 48 h after
transfection by scraping into SDS sample buffer. Peptides were
displayed on either 7.5% (MHC transfectants) or 12.5% (AMPD1 transfectants) SDS-polyacrylamide gels and electrotransferred to
nitrocellulose membranes using the Phast-Gel system (Pharmacia). Membranes were blocked with 5% dried milk powder in TBST overnight at
4°C. The supernatant of either the anti-Myc MAb or the anti-MHC MAb
was incubated with the membrane for 2 h at room temperature. After
membranes were washed three times with TBST, they were incubated with
1:5,000 dilution of peroxidase-conjugated goat anti-mouse IgG in TBST
containing 1% dried milk powder. After blots were washed three times
in TBST, they were developed with an enhanced chemiluminescence Western
blotting detection system (Amersham International).
Colocalization of AMPD1 and MHC.
In cotransfected cells that expressed both the AMPD1 and MHC peptides,
two patterns of immunofluorescence were observed. In some cells, the
AMPD1 stain was concentrated over, or superimposed on, the MHC
filaments, whereas in other cells with filamentous MHC the AMPD1 stain
was distributed throughout the cytoplasm. The percentage of dually
transfected cells exhibiting colocalization of AMPD1 and MHC was
determined for each independent transfection experiment. Two hundred
randomly encountered cells expressing both peptides were scored as
either diffuse AMPD1 stained or AMPD1 colocalized with MHC by examining
each cell with a red filter first to detect MHC filaments, followed by
examination with a green filter to determine the AMPD1 staining
pattern. Marginal images that were difficult to score as colocalized
vs. nonlocalized were classified as nonlocalized or diffuse. Three
individuals evaluated the slides independently, and their scores or
percent colocalized was concordant in all experiments.
Confocal microscopy and image reconstruction.
Cytosolic distributions of AMPD1 and MHC peptides were examined using a
confocal laser microscope illuminated with a krypton/argon laser
(Bio-Rad, MRC 600). With the use of a ×60 oil immersion lens
(Olympus, Splan Apo, 1.4 numerical aperture), optical sections were
collected at intervals of 0.8-µm increments through the entire z-axis of each cell. A single optical
section was selected from the midregion of the transfected cell to
provide a cross-sectional view of the intracellular distribution of the
AMPD1 and MHC peptides. The FITC-labeled distribution of AMPD1 and the
TXRD-labeled distribution of MHC were then combined using Adobe
Photoshop. The surface plot module of National Institutes of Health
Image 1.01 freeware was used for the integrated density volume mapping
to quantitatively assess the intracellular abundance of AMPD1 and MHC
in transfected cells (4, 14). Surface density maps displaying these
results are shown in Fig. 3
(bottom). The distributions of the
AMPD1 and MHC peptides in the cytosol are illustrated by the peaks and
troughs of high- and low-intensity staining.
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RESULTS |
Colocalization of AMPD1 and MHC peptides in transfected cells.
Preliminary experiments were performed to demonstrate that the
respective expression vectors and cell culture conditions lead to
production of AMPD1 or MHC peptides of the appropriate size (Fig.
1). Immunoblots of protein extracted from
HeLa cells transfected with the AMPD1 expression vector confirm the
presence of an AMPD1 peptide with an estimated size of 80 kDa, the
predicted size for the nonproteolyzed, full-length AMPD1 peptide (13).
The MHC peptide produced from this expression vector has an estimated molecular mass of 200 kDa, the size predicted for the
full-length MHC peptide. Transfection of COS cells with these same
vectors gave similar results (data not shown).

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Fig. 1.
Immunoblots of HeLa cells transfected with the myosin heavy chain (MHC)
or AMP deaminase 1 (AMPD1) expression vector. Lysates of HeLa cells
transfected for 48 h with either 10 µg of AMPD1 or MHC expression
plasmid were resolved on 7.5 or 12.5% SDS-polyacrylamide gels,
respectively. After transfer to nitrocellulose membranes, AMPD1 and MHC
peptides were identified by immunoblotting with the respective
monoclonal antibodies (MAbs). AMPD1 expression vector produces a
peptide that contains an epitope recognized by a Myc MAb added to the
amino terminus of AMPD1 (see Fig. 4). MHC peptide was detected by an
MAb to anti-sarcomeric MHC. Left:
lane 1, lysate of HeLa cells
transfected with MHC expression vector; lane
2, lysate of mock-transfected cells.
Right: lane
1, lysate of HeLa cells transfected with AMPD1
expression vector; lane 2, lysate of
mock-transfected cells.
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Transfection of the expression vector encoding the epitope-tagged,
native AMPD1 cDNA alone into HeLa cells produces an immunofluorescent staining pattern in which AMPD1 is distributed diffusely throughout the
cytoplasm and excluded from the nucleus (Fig.
2, top
left). Transfection of HeLa cells with the MHC
expression vector alone gives an immunofluorescent staining pattern
similar to that reported by Vikstrom et al. (30); MHC filaments
aggregate to produce flecks scattered throughout the cytoplasm (Fig. 2,
top right). Transfection of the
expression vector alone, which contains neither the AMPD1 nor MHC cDNA,
produces no detectable immunofluorescent staining under the conditions
employed. After cotransfection with both the AMPD1 and MHC expression
vectors, many cells express both peptides (Fig. 2,
bottom panels). In some cells, the
AMPD1 staining is superimposed on the MHC filaments, and, in other
cells with easily discernible MHC filaments, AMPD1 staining is
distributed diffusely throughout the cytoplasm. Confocal laser
microscopy confirms colocalization of AMPD1 and MHC in cotransfected
HeLa cells (Fig. 3). In 17 independent
transfections, 32 ± 2.3% of HeLa cells exhibited colocalization of
AMPD1 and MHC.

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Fig. 2.
Immunofluorescent staining of transfected HeLa cells. HeLa cells grown
on coverslips were transfected with the AMPD1 expression vector alone
(top left) or the MHC expression
vector alone (top right) or
cotransfected with both expression vectors
(bottom). Cells shown on
left were stained with the Myc MAb,
which detects the Myc epitope-tagged AMPD1 peptide. Cells shown on
right were stained with the
anti-sarcomeric MHC MAb. TXRD, Texas red.
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Fig. 3.
Confocal laser microscopy of transfected HeLa cells. HeLa cells were
prepared as in Fig. 2 and photographed with confocal laser microscopy.
Top left: cells were transfected with
the AMPD1 expression vector alone and stained with the Myc MAb, which
detects the Myc epitope-tagged AMPD1 peptide. Top
right: cells were transfected with MHC expression
vector alone and stained with MHC MAb.
Middle: a single cell cotransfected
with both expression plasmids visualized with a green filter
(epitope-tagged AMPD1 peptide; left)
and visualized with a red filter (MHC peptide;
right).
Bottom: 3-dimensional display of
integrated density volume calculated for the AMPD1 peptide
(left) and for the MHC peptide (right). TXRD,
Texas red.
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Initially, it was perplexing why only a fraction of cells expressing
both peptides exhibits colocalization of AMPD1 and MHC. Variability in
experimental conditions was not considered to be a likely explanation,
since the fraction of HeLa cells exhibiting colocalization of AMPD1 and
MHC was very consistent in the different transfection experiments, as
evidenced by the small coefficient of variation among the 17 independent transfections.
Cells that demonstrate colocalization of AMPD1 and MHC do not contain
more or less AMPD1 than cells that do exhibit colocalization to the
extent that this can be determined quantitatively by confocal laser
microscopy. The relative abundance of the AMPD1 and MHC peptides in a
given transfected cell was determined from the confocal laser images
using a three-dimensional display as illustrated in Fig. 3
(bottom). The integrated density
volumes were calculated for the AMPD1 and MHC peptides, respectively
(4, 14). These analyses were repeated in triplicate, and the ratio of
the AMPD1 peptide relative to that of the MHC peptide in a given cell
was determined. For the native AMPD1 peptide, the ratio of AMPD1 to MHC
was determined to be 1.09 ± 0.02 (Table
1), and this ratio was not appreciably
different in cells that exhibited colocalization of AMPD1 and MHC vs.
those that exhibited a diffuse staining pattern for AMPD1. These
findings suggest that some factor other than variation in the
intracellular level of AMPD determines the percentage of cells
demonstrating colocalization.
A potential explanation for why some cells exhibit colocalization and
others do not is suggested by previous studies. AMPD binding to MHC in
myocytes is reversible and varies depending on the intracellular
concentration of ATP or the adenylate energy charge (13, 22, 23).
Results that are presented later in this study demonstrate
that increasing or decreasing the intracellular content of ATP
reproducibly decreases or increases, respectively, the fraction of
cells exhibiting colocalization of AMPD1 and MHC. Point mutation of the
ATP binding site in AMPD1 also reproducibly increases the fraction of
cells that exhibit colocalization. We conclude from these observations
that conditions in nonperturbed HeLa cells are near the equilibrium for
dissociation of AMPD1 from MHC. If more of the AMPD1 is bound than
free, staining over the MHC filaments is more intense and these cells
are scored as localized AMPD1 in this assay. Perturbations of the
intracellular environment or mutations in the AMPD1 peptide shift this
equilibrium, and the percentage of cells exhibiting colocalization
increases or decreases as a reflection of the shift between bound and
free AMPD1. Thus the fraction of cells exhibiting colocalization of AMPD1 and MHC provides a method for quantitating AMPD1 binding to MHC
in the intact cell.
To confirm the utility of this cotransfection model for assessing AMPD1
binding to MHC, another cell line was also utilized. COS cells
transfected with the AMPD1 expression vector alone give a diffuse
immunofluorescent staining pattern, COS cells transfected with the MHC
expression vector give a filamentous staining pattern, and COS cells
transfected with both vectors give one of two AMPD1 staining patterns:
diffuse or colocalization with MHC. In four independent cotransfection
experiments, 58 ± 14% of COS cells exhibited colocalization of
AMPD1 and MHC. Although fewer experiments were performed with COS
cells, mutations of the AMPD1 peptide that increase or decrease binding
to MHC in HeLa cells also increase or decrease binding in COS cells.
Thus binding of AMPD1 to MHC is not restricted to the environment of
the HeLa cell, and the model reflects qualitatively similar changes in
binding in both cell types.
Transfection experiments with the AMPD1 expression vectors were also
attempted with a soleus 8 myoblast cell line to assess binding of AMPD1
to intact myofibrils. Although the AMPD1 peptide was easily detectable
in myoblasts 48-72 h posttransfection, expression was extinguished
or the myoblasts killed after 72 h, before myotube formation.
Subsequent experiments have confirmed that expression of the AMPD1
peptide during the late myoblast stage is deleterious to the
differentiation process.
Identification of the MHC binding domain in AMPD1.
As illustrated in Fig. 4, a series of
amino- and carboxy-terminal deletions were made in the AMPD1 peptide.
The diagram of the AMPD1 peptides illustrated in Fig. 4 has
been broken into sections to indicate the location of the Myc epitope
tag; the nonconserved, noncatalytic region of AMPD1; the conserved
catalytic domain; and the positions of the catalytic and ATP binding
sites (8). The expression vectors encoding these deletion constructs produce AMPD1 peptides of the predicted size in HeLa cells as demonstrated in Fig. 5.

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Fig. 4.
MHC binding of truncated AMPD1 peptides. Progressive deletions were
made from the amino terminus (N series) and carboxy terminus (C
series) of AMPD1 peptide. All peptides were engineered to contain Myc
epitope at the amino terminus. AMPD1 peptide is divided into conserved
and nonconserved regions, and locations of the catalytic center and ATP
binding site in the carboxy terminus are illustrated. E2+, wild-type
AMPD1 peptide that retains exon 2-encoded sequences. Number of amino
acid residues deleted from either the amino or carboxy terminus of E2+
peptide is indicated at left, adjacent
to each construct. At right are the
percentages of HeLa cells expressing both the AMPD1 and MHC peptides
that exhibit superimposition of AMPD1 on MHC filaments. Numbers in
parentheses are the number of independent transfection experiments
performed with each AMPD1 construct.
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Fig. 5.
Top: progressive deletions were made
from the amino terminus (N series) and carboxy terminus (C
series) of AMPD1 peptide as described in Fig. 4.
Bottom: immunoblots of HeLa Cells
transfected with AMPD1 deletion constructs. Lysates of HeLa cells
transfected with the AMPD1 deletion constructs illustrated in Fig. 4
were displayed on SDS-gels and immunoblotted as described in Fig. 1. A
comparable amount of protein extract (1 µg) was loaded in each well.
Lane 1, E2+ construct;
lane 2, E2 construct (isoform
that excludes sequences encoded by exon 2); lane
3, N 65 construct; lane
4, N 178 construct; lane
5, N 333 construct; lane
6, C 85 construct; lane
7, C 118 construct; lane
8, C 426 construct.
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The wild-type peptide used for comparison in this set of experiments is
the full-length, exon 2 plus (E2+) isoform of AMPD1. Deletion of the
amino-terminal 65 residues of AMPD1 (N
65) virtually abolishes
binding of this peptide to MHC; the percentage of HeLa cells exhibiting
colocalization falls from 31.7 ± 2.3% to 2.2 ± 0.9%
(P < 0.01) (Fig. 4). This truncated
peptide accumulates in amounts relative to MHC that are comparable to
that for the wild-type peptide (Table 1), excluding peptide abundance
as the explanation for its reduced binding to MHC. Prior studies have shown that deletion of these amino-terminal residues has only a minor
effect on the catalytic activity of AMPD1 (8), providing evidence that
this truncated form of AMPD1 retains some structural features similar
to that of the native enzyme.
Deletion of another 113 residues (N
178) from the amino terminus not
only restores binding but cells expressing this truncated peptide
exhibit an even higher percentage of colocalization than observed in
cells expressing the wild-type peptide (60 ± 2.7% vs. 31.7 ± 2.3%, P < 0.05) (Fig. 4). Deletion
of an additional 155 residues (N
333) from the amino terminus reduces
binding of AMPD1 to MHC to very low levels; only 8.6 ± 0.9% of
cells expressing this truncated peptide exhibits colocalization
(P < 0.05 compared with the
wild-type peptide). These truncated peptides accumulate in cells in
amounts relative to MHC which are comparable to that observed for the
wild-type peptide (Table 1), excluding abundance of the mutant AMPD1
peptides relative to that of MHC as the explanation for observed
differences in binding.
Deletions from the carboxy-terminal region can also affect AMPD1
binding to MHC. Removal of 85 residues (C
85) from the carboxy terminus has no discernible effect on binding; 31.4 ± 4.4% of cells expressing this truncated peptide exhibits colocalization, a
value comparable to that observed in cells expressing the wild-type peptide (P > 0.5). Deletion of an
additional 118 residues (C
118) removes both the catalytic site and
the ATP binding site (8). Cells expressing this truncated peptide
exhibit a significantly higher percentage of colocalization of AMPD1
and MHC compared with cells expressing the wild-type peptide (61.8 ± 4.2% vs. 31.7 ± 2.3%, P < 0.05). Deletion of another 426 residues (C
426) from the carboxy
terminus does not appreciably change binding from that observed with
the C
118 construct; 77.6 ± 1.8% of cells that express the
C
426 peptide exhibits colocalization. As with the other mutant
constructs, abundance of the AMPD1 peptides relative to that of MHC in
the transfected cells does not explain the observed differences in
binding observed with these mutants (Table 1).
These deletion constructs define a minimal region within the AMPD1
peptide that is essential for binding of this peptide to MHC (Fig. 4).
Up to 178 residues can be deleted from the amino terminus, and up to
426 residues can be deleted from the carboxy terminus with preservation
of binding of the truncated AMPD1 peptides to MHC. These analyses
suggest that a myosin binding domain(s) may reside within the region of
AMPD1 encompassed by residues 178-333. To confirm that the region
of AMPD1 encompassed by residues 178-333 includes a myosin binding
site, these sequences were ligated into the expression vector to make a
construct called N178-333. In three experiments with the
N178-333 construct, colocalization of this AMPD peptide with MHC
was observed in >90% of dually transfected cells (Fig. 4).
ATP modulates AMPD1 binding to MHC.
Prior studies with resting and stimulated skeletal muscle (1, 23) and
isolated MHC and AMPD1 peptides (13) indicate that ATP, or some
derivative of the adenylate energy charge, modulates binding of AMPD1
to MHC. Two approaches were taken in the present study to define the
role of ATP in the control of AMPD1 binding to MHC and to identify
regions of the AMPD1 peptide that control binding of AMPD1 to MHC in
response to changes in the intracellular concentration of ATP. One
series of experiments employed the AMPD1 mutants described above. These
mutants were transfected into HeLa cells, and the cells were
subsequently permeabilized to permit external control of the ATP
concentration in the cell.
As illustrated in Fig. 6, binding of
wild-type AMPD1 peptide to MHC in permeabilized HeLa cells responds to
changes in the intracellular concentration of ATP. At lower than
ambient ATP concentrations, the percentage of cells exhibiting
colocalization of AMPD1 and MHC increases from 31.7 ± 2.3% to 70.6 ± 3.4% (P < 0.05); at higher
than ambient ATP concentrations, the percentage of cells exhibiting
colocalization decreases from 31.7 ± 2.3% to 20.3 ± 1.8%
(P < 0.1). Deletions in the
carboxy-terminal region of AMPD1 (C
426 and C
118), which
eliminates the segment of this peptide containing the ATP binding site
and the catalytic site (Fig. 4), render the truncated AMPD1 peptides
resistant to changes in ATP concentrations with respect to their
binding to MHC (Fig. 6). A more restricted deletion of carboxy-terminal
residues (C
85) that preserves the region of AMPD1 that contains the
ATP binding site and the catalytic site produces a peptide that
exhibits binding characteristics in response to changes in the
intracellular concentration of ATP similar to those observed with the
wild-type peptide.

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Fig. 6.
Effect of truncations at the carboxy terminus on ATP-induced changes in
AMPD1 binding to MHC. Wild-type (E2+) and carboxy-terminal truncation
mutants of AMPD1 were cotransfected with MHC into HeLa cells, and
transfectants were subsequently permeabilized to permit perfusion with
solutions containing different ATP concentrations. Individual cells
were scored for colocalization of AMPD1 and MHC as described in Fig. 4.
For each AMPD1 construct (E2+, C 85, C 118, and C 426),
percentage of cells expressing both peptides that exhibit
superimposition of AMPD1 on MHC filaments is illustrated for 3 different conditions [0 mM ATP and metabolic inhibitors in
perfusion buffer (solid bars); ambient ATP, i.e., nonpermeabilized
cells (open bars); and 10 mM ATP in perfusion buffer (hatched
bars)]. Number of independent experiments performed for each
condition is indicated above bars, and SE observed among the different
transfection experiments are illustrated by vertical lines on each
bar.
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The above studies with the carboxy-terminal deletion mutants are
consistent with the hypothesis that the ATP binding site located in
this region of the peptide is responsible for the observed effects of
ATP in modulating binding of AMPD1 to MHC, but these experiments have
several limitations. As demonstrated in Fig. 5, the C
85 and
especially the C
118 mutants are subject to proteolysis. We cannot
exclude that this proteolysis occurred in the cell and that the
resultant mixture of native and degraded peptides contributed to some
of the observed differences in ATP responsiveness of these carboxy-terminal deletion mutants. Furthermore, these deletion constructs removed residues in addition to the ATP binding site. Therefore, another set of experiments was performed that utilized AMPD1
peptides engineered to contain point mutations in the ATP binding site
or the catalytic center of AMPD1. Chimeric rat/human AMPD1 cDNAs were
used for these studies to take advantage of point mutants in human
AMPD1 characterized previously (8). As illustrated in Fig.
7, substitution of a glycine for aspartate
at position 650, a mutation known to destroy the catalytic center of
this enzyme (8), has no effect on binding of this chimeric AMPD1 peptide to MHC at ambient ATP concentrations in HeLa cells (32 ± 13.8% vs. 31.7 ± 2.3% for the mutant and wild-type peptides, respectively, P > 0.5). In contrast,
mutation of residue 663 (glutamine to lysine) within the ATP binding
site of the rat/human chimeric AMPD1 peptide (8) results in a peptide
that exhibits increased binding to MHC at ambient ATP concentrations in
HeLa cells (82 ± 8.3% vs. 31.7 ± 2.3% colocalization of the
mutant chimera vs. the wild-type rat peptide,
P < 0.05). When HeLa cells
containing the 663 glutamine-to-lysine mutation are permeabilized and
exposed to media containing different ATP concentrations,
colocalization of the ATP binding site mutant is not appreciably
altered by either higher or lower than ambient ATP concentrations (data
not shown). These two chimeric peptides accumulate in amounts relative
to MHC that are comparable to that observed for the wild-type rat AMPD1
peptide, and their sizes are comparable to that of the native rat AMPD1
peptide (data not shown).

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Fig. 7.
Effect of point mutations in the ATP binding site and catalytic site on
AMPD1 binding to MHC. Chimeric cDNAs were constructed that fuse the
amino terminus of the wild-type (E2+) rat AMPD1 to the carboxy terminus
of a mutant human AMPD1, as illustrated in the diagram at
top. Rat/human chimeras were selected
for these experiments because prior studies have characterized the
physiological consequences of the point mutations in the human peptide
(26). Transfections and scoring for colocalization of AMPD1 and MHC
were performed as described in Fig. 2 and MATERIALS
AND METHODS. All experiments depicted were performed in
nonpermeabilized cells. Number of independent transfections performed
with each construct is indicated above bars
(bottom), and SE observed among the
different experiments are illustrated by vertical line on each bar.
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Effect of the amino terminus on AMPD1 binding to MHC.
Results presented in Fig. 4 demonstrate that residues in the
amino-terminal region of AMPD1 have a profound effect on the binding of
this peptide to MHC. Deletions within the amino terminus appear to have
paradoxical effects on binding of AMPD1 to MHC; the N
65 mutant
reduces binding at ambient ATP concentrations to essentially background
levels, whereas deletion of an additional 113 residues (N
178)
results in binding levels at ambient ATP concentrations that are
greater than that observed with the wild-type peptide. This paradox is
explained by results obtained from experiments that examine binding of
the various amino-terminal deletion mutants at different ATP
concentrations (Fig. 8).

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Fig. 8.
Effect of truncations at amino terminus on ATP-induced changes in AMPD1
binding to MHC. Wild-type (E2+) and amino-terminal truncation mutants
of rat AMPD1 were cotransfected with MHC into HeLa cells. Transfectants
were permeabilized and perfused with solutions containing different ATP
concentrations, as described in Fig. 6. For each AMPD1 construct (E2+,
N 65, N 178, and N 333), percentage of cells expressing both
peptides that exhibit superimposition of AMPD1 on MHC filaments is
illustrated for 3 different conditions [0 mM ATP and metabolic
inhibitors in perfusion buffer (solid bars); ambient ATP, i.e.,
nonpermeabilized cells (open bars); and 10 mM ATP in perfusion buffer
(hatched bars)]. SE observed among the different transfection
experiments are illustrated by vertical line on each bar, and number of
experiments is indicated in parentheses.
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The N
65 mutant exhibits colocalization with MHC in only 2.2 ± 0.9% of HeLa cells at ambient ATP concentrations. However, when the
intracellular ATP concentration is reduced in permeabilized cells, the
percentage of cells exhibiting colocalization increases to 20 ± 4.0% (P < 0.05). Thus this deletion
mutant is capable of binding to MHC, but binding is reduced at all ATP
concentrations. These results suggest that the amino-terminal 65 residues of AMPD1 influence binding of this peptide to MHC through
enhancing sensitivity to ATP-induced inhibition of binding.
In contrast to the N
65 mutant, the N
178 is less sensitive to
ATP-induced inhibition of binding to MHC. This mutant exhibits a level
of binding to MHC at ambient ATP concentrations that is comparable to
that observed with the wild-type peptide at reduced ATP concentrations.
Moreover, the percentage of cells exhibiting colocalization of the
N
178 mutant with MHC is unchanged at low, ambient, and high ATP
concentrations (Fig. 8). These results suggest that residues within the
region of the AMPD1 peptide encompassed by amino acids 66-177 are
necessary for ATP to inhibit AMPD1 binding to MHC.
The results with the amino-terminal deletion mutants indicate that the
more proximal regions of the AMPD1 peptide have the potential to
influence binding of this enzyme to MHC through modulation of the
sensitivity to ATP-induced inhibition of binding. For this region of
the peptide to exert a regulatory effect on binding, there would need
to be a physiological mechanism for altering the structure of the
amino-terminal domain of AMPD1. The second exon of AMPD1, which encodes
residues 8-12 of this peptide, is retained or excluded from the
mature messenger RNA derived from the AMPD1 primary transcript in
response to stage-specific and tissue-specific signals (16, 18). These
sequences fall within the region of the amino terminus that influences
the sensitivity of the AMPD1 peptide to ATP-induced inhibition of
binding.
To determine if the alternative isoforms of AMPD1 exhibit differences
in binding to MHC, an expression vector was constructed that encodes an
AMPD1 peptide that excludes exon 2-derived sequences (E2
) for
comparison with the parent construct that includes exon 2-derived
sequences (E2+). Binding to MHC was assessed under several culture
conditions following cotransfection with the MHC expression vector
(Fig. 9).

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Fig. 9.
Effect of exon 2-encoded sequences on AMPD1 binding to MHC. Primary
transcript of the AMPD1 gene is alternatively spliced into two mature
mRNAs, one that retains exon 2 (E2+) and one that excludes exon 2 (E2 ). cDNAs for both the E2+ and E2 isoforms of AMPD1
were cloned into the pRC/CMV expression vector and transfected into
HeLa cells. Percentage of HeLa cells expressing one of the 2 AMPD1
isoforms and MHC were scored for colocalization of AMPD1 and MHC under
a variety of culture conditions. A:
percent colocalized was determined for both AMPD1 isoforms in cells
under routine culture conditions, i.e., medium containing 10% fetal
bovine serum (FBS), under culture conditions in which serum content of
the medium was reduced to 2.5% FBS, and under culture conditions in
which medium containing 2.5% FBS was supplemented with epidermal
growth factor, endothelial growth factor, insulin, transferrin,
triiodothyronine, progesterone, 17 -estradiol, testosterone,
hydrocortisone. B: percent colocalized
was determined for both AMPD1 isoforms in cells cultured in 10% FBS
and subsequently permeabilized and perfused with different ATP
concentrations as described in Fig. 6.
C: percent colocalized was determined
for both AMPD1 isoforms in cells cultured in 2.5% FBS and subsequently
permeabilized and perfused with different ATP concentrations as
described in Fig. 6. SE observed among the different transfection
experiments are illustrated by vertical line on each bar, and number of
experiments is indicated in parentheses.
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|
Routine culture conditions for the studies described above utilized
medium containing 10% FBS. Under these culture conditions, the
percentage of HeLa cells exhibiting colocalization of AMPD1 and MHC is
comparable for the E2+ and E2
isoforms (31.7 ± 2.3% vs.
25.6 ± 3.4%, P > 0.5) (Fig.
9A). In the course of testing different culture conditions, it was noted that the serum concentration of the medium has a profound effect on the binding of the E2
isoform to MHC at ambient ATP concentrations. Binding of the E2
isoform increases from 25.6 ± 3.4% to 61.5 ± 1.7%
(P < 0.05) in 2.5% serum, whereas
binding of the E2+ isoform is essentially unchanged (21 ± 9.2% vs.
31.7 ± 2.3%, P > 0.5). The rate
of growth of HeLa cells in 2.5% and 10% serum was comparable,
excluding this variable as an explanation for the difference in binding observed with the two AMPD1 isoforms. The abundance of the E2+ and
E2
peptides relative to that of MHC is essentially unchanged when the serum concentration is reduced from 10 to 2.5% FBS (Table 1).
In addition, supplementation of medium containing 2.5% serum with
epidermal growth factor, endothelial growth factor, insulin, transferrin, triiodothyronine, progesterone, 17
-estradiol,
testosterone, and hydrocortisone does not suppress binding of the
E2
isoform at ambient ATP concentrations (Fig. 9).
Because of the results obtained with the amino-terminal deletion
mutants, the alternatively spliced isoforms of AMPD1 were evaluated for
differences in sensitivity to ATP-induced inhibition of binding. After
permeabilization of HeLa cells cultured in either 10% serum (Fig.
9B) or 2.5% serum (Fig.
9C), the percentage of cells
exhibiting colocalization was determined at low, ambient, and high ATP.
In 10% serum, the two isoforms of AMPD1 exhibit comparable changes in
binding to MHC in response to alterations in the intracellular
concentration of ATP. However, in 2.5% serum, the E2
, but not
the E2+, isoform of AMPD1 exhibits reduced sensitivity to ATP-induced
inhibition of binding. The E2
isoform, in 2.5% serum, resembles
the N
178 deletion mutant in its binding characteristics in that it
is relatively insensitive to ATP-induced inhibition of binding.
 |
DISCUSSION |
Results of this and prior studies suggest that there are a number of
domains within the AMPD1 peptide that interact to control the activity
of this enzyme in response to the physiological needs of the myocyte
(Fig. 10). The peptide can be
subdivided into a nonconserved amino terminus that is unique to the
AMPD1 gene product and a conserved carboxy terminus that is shared
among the various members of this multigene family (24). The catalytic
site and an ATP binding site are located in the conserved,
carboxy-terminal region of the AMPD1 peptide (8, 13, 15). The present
study has identified a myosin binding domain in the amino terminus that resides within the region encoded by residues 178-333. These
studies do not exclude the presence of other myosin binding domains in the AMPD1 peptide, and the presence of one or more additional MHC
binding domains could contribute to the low level of binding observed
with the N
333 mutant. A portion of the amino-terminal region of
AMPD1 that encompasses the myosin binding domain, residues 182-209, shares sequence similarity with another myosin binding protein, titin (Table 2). This region of
titin, referred to as the PEVK domain, is thought to disrupt or
destabilize local tertiary structures because of the reduced complexity
of this repeating amino acid sequence and the cluster of negative
charges in the PEVK domain (12). These conserved sequences could
participate in the formation of the MHC binding domain in AMPD1 or play
a role in regulating accessibility of the MHC binding domain through their influence on local tertiary structure. The latter is an attractive hypothesis because of the data presented in this report, which suggest that a secondary or higher-order structure of the intact
AMPD1 peptide influences binding to MHC in response to changes in ATP
concentration, mediated by a site remote to the MHC binding domain, and
through alternative splicing in a region of the amino terminus distinct
from the MHC binding domain.

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Fig. 10.
Domains in AMPD1. Open portion of this bar diagram represents segment
of the AMPD1 peptide that is conserved among all members of this
multigene family. Stippled portion at amino terminus is the segment
that determines sensitivity to ATP-induced inhibition of binding to
MHC. Hatched portion in the middle of the peptide is the segment that
encompasses the MHC binding domain. Locations of exon 2-encoded
sequences, catalytic site, and ATP binding site are indicated by
vertical bars. Numbers at top of the
bar diagram indicate position of specific amino acid residues.
|
|
The model illustrated in Fig. 10 suggests a mechanism for regulating
the binding of AMPD1 to MHC and thereby controlling the activity of
this enzyme in the myocyte. This model may be oversimplified because
transfections were performed in nonmyocytes for the express purpose of
limiting the complexity of AMPD1 interaction with other myocyte-specific proteins. In addition, the model is based on forced
expression of AMPD1 and MHC using a transient transfection assay, and
there is no assurance that the relative abundance and/or higher-order structure of the AMPD1 and MHC peptides produced in these
nonmyocytes might not influence the interaction between these peptides.
Although these recognized limitations to the assay used in the present
study exist, none of the results obtained with the transient expression
of the mutant AMPD1 peptides in this model system is at variance with
prior studies that have examined the control of AMPD1 binding to MHC in
vivo (22, 23) or with purified protein preparations (1, 2, 10, 13). The
additional information obtained from the current study using the two
naturally occurring variants of AMPD1 suggests a heretofore unrecognized regulatory mechanism by which alterations in physiological conditions may differentially control the binding of these two AMPD1
isoforms to MHC.
Binding of AMPD1 to MHC is essential for activation of this enzyme
under conditions prevailing in the contracting myocyte (22, 23). The
E2+ isoform of AMPD1, the predominant isoform in adult, fast-twitch
glycolytic fibers (16, 18), is prevented from associating with MHC in
resting muscle by ATP binding to the nucleotide regulatory site in this
enzyme. During strenuous exercise when ATP levels in the myocyte fall
as a consequence of utilization of this high-energy substrate, the
inhibitory effect of ATP on binding of AMPD1 to MHC is diminished.
Under these circumstances, the E2+ isoform of AMPD1 translocates to the
myofibril where it is activated. Deamination of AMP by AMPD shifts the
adenylate kinase equilibrium toward ATP formation, and this increases
the adenylate energy charge (24). Thus activation of the E2+ isoform of
AMPD1 through binding to MHC could help to preserve the adenylate energy charge at a time when energy production is limited. This function of AMPD could protect the myocyte from irreversible injury, which would ensue if the ATP or adenylate energy charge fell below a
critical level following intense muscle contraction.
Another function that has been ascribed to AMPD in myocytes is the role
this enzyme plays in controlling flux through the purine nucleotide
cycle (24). When IMP is formed as a consequence of AMP deamination, IMP
can be condensed with aspartate, subsequently leading to the formation
of fumarate and AMP. This series of reactions is referred to as the
purine nucleotide cycle (in which SAMP is succinyl
AMP)
In myocytes, fumarate production via the purine nucleotide cycle is the
only known pathway for expanding the pool of citric acid cycle
intermediates during muscle contraction (24), and flux through the
purine nucleotide cycle is inhibited when GTP levels are reduced (24).
The simultaneous requirement for a high level of GTP and activation of
AMPD presents a metabolic dilemma if the E2+ isoform of AMPD1 is needed
for flux through the purine nucleotide cycle. As discussed above, the
E2+ isoform is activated only after the adenylate energy falls, and
such a fall in the adenylate energy charge will also lead to a drop in GTP levels in the cell. Under these conditions, the two arms of the
purine nucleotide cycle are not coordinated.
The different myosin binding properties of the E2
isoform provide a potential mechanism for bypassing this metabolic
dilemma. The E2
isoform of AMPD1 can bind to MHC in the presence
of high-ATP concentrations under specified conditions, and activation
of AMPD under these conditions would lead to flux through the purine
nucleotide cycle and fumarate production. In vitro, the E2
isoform can overcome ATP-induced inhibition of binding to MHC in
response to a signal produced by some component of the tissue culture
medium. In vivo, a hormonal or neural stimulus may provide a signal to
the E2
isoform of AMPD1, which releases it from ATP-induced
inhibition of binding to MHC. The relative abundance of the E2
isoform of AMPD1 in different skeletal muscle fiber types is consistent
with a potential role of the E2
isoform in controlling flux
through the purine nucleotide cycle and generation of citric acid cycle intermediates. Fast-twitch glycolytic fibers have the lowest amount of
the E2
relative to the E2+ isoform, and slow-twitch oxidative fibers have the highest amount of the E2
relative to
the E2+ isoform (16, 18). Through regulation of the alternative
splicing pathway, the myocyte can produce varying ratios of the
E2
and E2+ isoforms of AMPD1. Binding of these two AMPD1
isoforms to MHC with subsequent activation of this enzyme could be
differentially regulated to meet the particular needs of different
fiber types under various physiological conditions.
 |
ACKNOWLEDGEMENTS |
We thank Dr. Judy Swain of the Division of Cardiology, Dr. Lee Sweeney
of the Muscle Institute, and Dr. Haruhiko Siomi of the Howard Hughes
Medical Institute for numerous discussions regarding this work and
their critical review of this manuscript. We also thank Dr. Gideon
Dryfus of the Howard Hughes Medical Institute for the generous use of
the inmmunofluorescence and confocal laser microscopes in his
laboratory.
 |
FOOTNOTES |
The MAb MF20 developed by Evans was obtained from the Departmental
Studies Hybridoma Bank maintained by the Department of Pharmacology and
Molecular Sciences, Johns Hopkins University School of Medicine,
Baltimore, MD, and the Department of Biological Sciences, University of
Iowa, Iowa City, IA, under contract N01-HD-2-3144 from the
National Institute of Child Health and Human Development. This work was
supported by National Institute of Diabetes and Digestive and Kidney
Diseases Grant DK-12413 (E. W. Holmes).
Present addresses: I. Hisatome, 1st Department of Medicine, Tottori,
University Faculty of Medicine, Nishimachi 36-1, Yonago 683, Japan; H. Kamma, Department of Pathology, Institute of Basic Medical Sciences,
University of Tsukuba, Ibaraki, 305-8575, Japan; H. Morisaki,
Department of Bioscience, National Cardiovascular Center Research
Institute, Osaka, Japan.
Address for reprint requests: E. W. Holmes, Stanford Univ. School of
Medicine, Office of the Dean, M-121, 300 Pasteur Drive, Stanford, CA
94305-5119.
Received 15 August 1997; accepted in final form 19 May 1998.
 |
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