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Proc Natl Acad Sci U S A. 2001 December 4; 98(25): 14440–14445. Published online 2001 November 20. doi: 10.1073/pnas.251541198. | PMCID: PMC64700 |
Copyright © 2001, The National Academy of Sciences Cell Biology Atrogin-1, a muscle-specific F-box protein highly expressed
during muscle atrophy Marcelo D. Gomes,*† Stewart H. Lecker,*†‡ R. Thomas Jagoe,* Ami Navon,* and Alfred L. Goldberg*§ *Department of Cell Biology, Harvard Medical School, 240 Longwood Avenue, Boston, MA 02115; and ‡Beth Israel Deaconess Medical Center, Renal Unit DA517, 330 Brookline Avenue, Boston, MA 02215 Received July 8, 2001; Accepted October 12, 2001. Muscle wasting is a debilitating consequence of fasting,
inactivity, cancer, and other systemic diseases that results primarily
from accelerated protein degradation by the ubiquitin-proteasome
pathway. To identify key factors in this process, we have used cDNA
microarrays to compare normal and atrophying muscles and found a
unique gene fragment that is induced more than ninefold in
muscles of fasted mice. We cloned this gene, which is expressed
specifically in striated muscles. Because this mRNA also markedly
increases in muscles atrophying because of diabetes, cancer, and renal
failure, we named it atrogin-1. It contains a functional F-box domain
that binds to Skp1 and thereby to Roc1 and Cul1, the other components
of SCF-type Ub-protein ligases (E3s), as well as a nuclear localization
sequence and PDZ-binding domain. On fasting, atrogin-1 mRNA levels
increase specifically in skeletal muscle and before atrophy occurs.
Atrogin-1 is one of the few examples of an F-box protein or Ub-protein
ligase (E3) expressed in a tissue-specific manner and appears to be a
critical component in the enhanced proteolysis leading to muscle
atrophy in diverse diseases. In mammals, muscle protein
serves as a primary reserve of amino acids that can be mobilized during
fasting and disease to provide a source of amino acids for hepatic
gluconeogenesis and energy production ( 1). An important physiological
adaptation to fasting is an increase in the overall rate of breakdown
of muscle proteins leading to a rapid loss of muscle mass and protein
content. A similar rapid atrophy of muscle is a common debilitating
feature of many systemic diseases including diabetes, cancer, sepsis,
hyperthyroidism, and uremia ( 2, 3) and occurs in specific muscles upon
disuse or nerve injury ( 4, 5). These different forms of muscle atrophy
are characterized by a common set of biochemical changes ( 2, 3). Loss
of muscle protein occurs primarily through enhanced protein breakdown
because of activation of the ubiquitin (Ub)-proteasome pathway,
as shown by inhibitor studies in intact muscles ( 6), increased content
of Ub-protein conjugates, and by cell-free measurements of Ub-dependent
proteolysis ( 7, 8). In addition, these various atrophying muscles show
increases in mRNA for components of this pathway including
polyubiquitin, E2 14k, and multiple proteasome
subunits ( 2, 9, 10). However, the key ubiquitination enzymes active in
degrading the bulk of muscle protein in normal or catabolic states
remain unclear ( 11). Proteins destined for degradation by the Ub-proteasome pathway are
first covalently linked to a chain of Ub molecules, which marks them
for rapid breakdown to short peptides by the 26S proteasome ( 12). The
key enzyme responsible for attaching Ub to protein substrates is a
Ub-protein ligase (E3) that catalyzes the transfer of an activated form
of Ub from a specific Ub-carrier protein (E2) to a lysine residue on
the substrate. Individual E3s ubiquitinate specific classes of
proteins; hence, the identity of the proteins degraded by the
proteasome is largely determined by the complement of E3s active in
individual cells. The present studies were undertaken to identify key factors that may be
important in the acceleration of muscle proteolysis in catabolic
states. To establish a comprehensive picture of the transcriptional
adaptations that occur during various types of muscle atrophy, and that
may be responsible for the activation of protein breakdown, we have
used Incyte cDNA microarrays to compare mRNA levels in normal mouse
muscles to those from atrophying muscles. Because much is known about
the enhancement of proteolysis and other metabolic adaptations to
fasting ( 13– 15), we initially performed microarray experiments
comparing poly(A) + RNA from muscles of normal and
food-deprived mice, and we have identified a group of genes whose
transcripts increase markedly in the atrophying muscles. One expressed
sequence tag was of particular interest because its level increased
most dramatically on fasting. Therefore, we have cloned this protein
and defined its properties. We demonstrate here that this protein has
the properties of an E3 of the SCF class and is unusual in being
expressed selectively in striated muscle. We have also studied further
the expression of this gene on food deprivation and in several other
models of human diseases in which there is a marked acceleration of
muscle proteolysis. These studies demonstrate the existence of a unique
ubiquitination enzyme that appears to increase when muscles undergo
atrophy. cDNA Library Production and Screening. Total RNA was isolated from the gastrocnemius muscle of 2 d
food-deprived mice by using TRIzol (Life Technologies, Grand Island,
NY) reagent, and poly(A) + RNA was purified by
using the Oligotex mRNA isolation kit (Qiagen). A cDNA library was
constructed from mouse RNA by using the Superscript Plasmid cDNA
Synthesis system (Life Technologies). Analysis of the library revealed
an average insert size of 2–3 kb. Hybridization screens were performed
according to the procedures of Sambrook et al. ( 16) and
Church and Gilbert ( 17). From the 30,000 recombinants screened, six
positive clones were identified. Creation of Atrogin-1 Mutants. Atrogin-1 in BlueScriptII KS + (Stratagene) served
as the phagemid for the generation of single-strand DNA for
site-directed mutagenesis of the F-box sequence. The plasmid was
transformed into f+ Escherichia coli strain (RZ1032). A
single colony was grown to mid-log phase, superinfected with M13KO7
helper phage, and grown overnight at 37°C. The bacteria were removed
by centrifugation, and the phage was precipitated from the supernatant
with NaCl-polyethylene glycol solution. Single-strand DNA was purified
by phenol-chloroform extraction followed by ethanol precipitation and
resuspension in water. An aliquot of the preparation was used as a
template for oligonucleotide site-directed mutagenesis ( 16) to generate
both the deletion of the F-box motif (ΔFb-atrogin-1; amino acids
228–284; primer 5′-GAAGTGGTACTGGCAGAGTCGATCGGTGATCGTGAGGCCTTTGAAG-3′)
as well as for mutation of the first two residues of the F-box sequence
(atrogin-1 3A; D-L-P to A-A-A; primer
5′-GTTGTAAGCACACAGCCGCGGCGGTGATCGTGAGG-3′). Oligonucleotide primers for
the amplification of DNA fragments containing wild-type or mutant
versions of the atrogin-1 gene were designed with flanking attB1 or
attB2 sites for insertion into the GATEWAY donor vector pDONR201 (Life
Technologies). Primers with the following sequences were synthesized by
MWG Biotec (Ebersberg, Germany): AT1 (forward),
5′-attB1-GGGGACAAGTTTGTACAAAAAAGCAGGCTTCCTTGGGCAGGACTGGCGG-3′; and
AT1 (reverse),
5′-attB2-GGGGACCACTTTGTACAAGAAAGCTGGGTTATTCAGAACTTGAACAAATTG-3′.
The PCR products were cloned directly into pDONR201, and the resulting
plasmids were used to transfer the atrogin-1 gene sequences into
pDEST27 [glutathione S-transferase (GST) fusion
vector] via homologous recombination. Coprecipitation Experiments. For expression of GST-, Myc5-, FLAG-, and
hemagglutinin (HA)-tagged proteins, 293T cells were grown in
DMEM (Life Technologies) supplemented with 10% FBS. GST-atrogin-1,
GST-atrogin-13A, or GST-ΔFb-atrogin-1 in
pDEST27 (5 μg) was cotransfected with 5 μg of Myc5-Skp1 (kindly
provided by N. Ayad, Harvard Medical School) or HA-Skp1, FLAG-Cul1, and
FLAG-Roc1 (kindly provided by K. Tanaka, Tokyo Metropolitan Institute
of Medical Science) into 293T cells at 60–80% confluence by using
Superfect reagent (Qiagen) according to the manufacturer's
instructions. The cells were maintained in medium with 10% of FBS for
24 h before beginning the experiment. After harvest, the cells
were resuspended in cell lysis buffer [20 mM Na-Hepes, pH 7.7/225 mM
KCl/1% Triton X-100, supplemented with protease inhibitor mixture
(Roche Biomedical)] and lysed by vortexing. The lysates were cleared
by centrifugation at 10,000 × g for 20 min in an
Eppendorf microcentrifuge. Coprecipitation was carried out by
incubating 900 μl of cell lysate (containing 1.5–3.0 mg of protein)
with 20 μl of glutathione-Sepharose beads (Amersham Pharmacia) for
≈16 h at 4°C with gentle rocking. The beads were then washed three
times with 1 ml of cell lysis buffer containing 0.5% Triton X-100 and
225 mM KCl, resuspended in SDS loading dye, and resolved by SDS/PAGE.
Proteins were transferred to PDVF membrane (Millipore) and probed with
mouse monoclonal anti-HA, -FLAG, or -GST primary antibodies (Sigma) in
5% milk/PBS, at a dilution of 0.1 μg/ml, 0.1 μg/ml and
1:8000, respectively. Membranes were washed twice with PBS/0.1%
Tween-20, once with PBS and incubated with an alkaline
phosphatase-linked anti-mouse IgG fragment (Promega). Detection was by
enhanced chemiluminescence (CDP-Star/Tropix, Bedford, MA). Animal Models of Muscle Atrophy. Six-week-old male C57BL6 mice were deprived of solid food for 1 or
2 d but given free access to water. During this period, the
gastrocnemius lost 15–20% of its initial weight (R.T.J., S.H.L.,
M.D.G., and A.L.G., unpublished data). Both gastrocnemius muscles from
10 control or food-deprived mice were harvested and pooled to prepare
total RNA for the gene microarray, cDNA library preparation, and
northern analyses. Gastrocnemius muscles from uremic and diabetic rats
and controls were kindly provided by S. R. Price, J. L.
Bailey, and W. E. Mitch (Emory University, Atlanta). The uremic
rats were prepared by 5/6 nephrectomy and demonstrate marked muscle
atrophy and accelerated protein breakdown ( 18). Acute diabetes was
induced by streptozotocin administration, and muscles were harvested
3 d later when proteolysis was accelerated and animals demonstrate
marked muscle atrophy ( 19). Gastrocnemius muscles from rats implanted
with Yoshida Ascites Hepatoma and controls were provided by V. Baracos
(University of Alberta, Edmonton, AB, Canada). These muscles were
harvested 6 d after tumor implantation and demonstrate marked
muscle atrophy ( 20). A systematic analysis of the transcriptional
changes in these different conditions will be presented elsewhere. Cloning and Structure of the Atrogin-1 Gene. The gene transcript that was increased most dramatically (seven- to
ninefold) on the Incyte microarray after food deprivation corresponded
to an expressed sequence tag (GenBank accession no. AW051824) and was
chosen for further study. [The mouse gene has subsequently also been
identified by the RIKEN Mouse Gene Encyclopedia Project ( NP_080622)
( 42)]. After conducting sequence analysis in available expressed
sequence tag databases ( http://www.labonweb.com), we identified a
sequence extended at both 3′- and 5′- ends, which allowed us to clone a
1.1-kb fragment by PCR from a human skeletal muscle cDNA library. To
identify the full-length cDNA, the PCR-amplified human gene fragment
was used as a probe to screen a mouse cDNA library derived from the
atrophying hindlimb muscles of mice deprived of food for 2 d.
These tissues were chosen because of the high level of expression of
this gene at this time point compared to levels in fed controls (see
below). A positive clone containing a 2.1-kb insert was excised by
complete SalI and NotI double digestion,
subcloned into the BlueScriptII KS +, and
sequenced in its entirety. The nucleotide and deduced amino acid
sequences of the gene, which we term atrogin-1 (for atrophy gene-1),
are shown in Fig. A.
Analysis of the mouse DNA sequence (nucleotides 1–2071) indicated the
presence of a single 1,068-bp ORF with a predicted initiation codon
(ATG) at nucleotide 328 and a termination codon at nucleotide 1,395.
This ORF codes for a polypeptide of 355-aa residues with a molecular
mass of 41,503 Da and a pI of 9.46. The initiation codon (nucleotide
328) is embedded in the sequence ACCATG, which matches perfectly with
the eukaryotic translation initiation consensus sequence ACCATG ( 21).
Because no other ATG was found upstream of this initiation codon, this
region probably represents the complete coding sequence. Comparison of the atrogin-1 cDNA sequence with the National Center for
Biotechnology Information sequence database identified four other
similar genes, including the human homolog, orthologs in both
Caenorhabditis elegans (Dy3.6) and
Drosophila melanogaster (cg11658), and a similar human
protein, FBXO25. Human and mouse atrogin-1 are almost identical,
differing in only 11 amino acids. Atrogin-1 shares slightly greater
overall amino acid sequence identity with FBOX25 (60%) than with other
members of the family. Atrogin-1 is 27% identical to C.
elegans Dy3.6 and 26% identical to the cg11658 protein predicted
from Drosophila genome sequence (Fig. C). Atrogin-1 Is an F-Box Protein, a Component of an SCF E3. A systematic search for functional domains within the atrogin-1
sequence by using the pfscan algorithm
( http://www.isrec.isb-sib.ch) and cansite
( http://www.cansite.bidmc.harvard.edu) indicated the presence of
four conserved motifs (Fig. B). An F-box is present at
amino acid 228–267 (Fig. D). This highly degenerate motif
is found in a family of proteins, most of which function as one
component of the SCF family of Ub-protein ligases (E3s) ( 22). The F-box
protein functions as an adaptor that binds proteins to be ubiquitinated
and, through the F-box, associates with the Skp1 protein (or a homolog)
and thus with other components of the E3 complex ( 23). Atrogin-1 lacks
the common motifs found in other F-box proteins that are known to
interact with protein substrates (e.g., leucine-rich regions, WD40
repeats). However, at its extreme carboxy terminus (amino acids
346–355), atrogin-1 does contain a motif known to interact with
proteins containing class II PDZ domains. PDZ domain-containing
proteins bind to specific sequences at the extreme C-termini of target
proteins ( 24, 25) and may represent a new type of substrate-binding
region in F-box proteins. By using the convention proposed by
Cenciarelli et al. ( 26) and Winston et al. ( 27),
we categorize atrogin-1 as an FBXO class of F-box protein, X referring
to its lack of a recognized substrate-binding region. Other features
also present in the molecule include a potential bipartite nuclear
localization sequence (amino acid 267–284) and a cytochrome
c family heme-binding site signature (amino acid 317–322). To demonstrate that the F-box domain in atrogin-1 is functional, we
tested whether this protein associates with Skp1, the classical
F-box-binding partner in SCF complexes. For these studies, we also
created mutant forms of atrogin-1 in which the amino acids at positions
227–229 containing the first two highly conserved residues of the
putative F-box were replaced by alanines
(atrogin-13A) and a second mutant in which the
putative F-box was deleted completely (ΔFb-atrogin-1). These
wild-type and mutant forms of atrogin-1 were expressed as GST-fusions
in 293T cells also expressing Myc5-Skp1, and the
cell lysate was subjected to a GST pull-down assay. Atrogin-1 was
precipitated together with Skp1, as detected by immunoblot analysis
(Fig. E). However, both mutant forms that lacked the
complete F-box sequence failed to associate with Skp1 (Fig.
E, lanes 2 and 3). Thus, atrogin-1 can interact
directly with Skp1 through the F-box domain. Further evidence for a
direct interaction between atrogin-1 and Skp1 was our finding that the
abundance of GST-atrogin-1 was increased when it was coexpressed with
Myc5-Skp1 (data not shown). This in
vivo stabilization depended on the F-box, because the abundance of
mutant forms of atrogin-1 lacking the F-box was unaffected by
Myc5-Skp1 coexpression (data not shown). To learn whether atrogin-1 exists in cells as part of an SCF complex,
we tested whether the other components, Cul1 and Roc1, were also
associated with GST-atrogin-1 and Skp1. 293T cells were transiently
transfected with constructs encoding FLAG- or HA-tagged versions of
these various proteins, the cell lysed, and atrogin-1 was then
precipitated. These pellets contained also Skp1 as well as Cul1 and
Roc1 (Fig. ), which were identified by
immunoblot. In contrast, these other SCF subunits were not found
coprecipitated with the versions of atrogin-1 containing a mutation or
deletion in the F-box. These observations provide strong evidence that
atrogin-1 is a component of an SCF E3. The functional significance of
the other putative domains and its substrates are presently being
studied.
Expression of Atrogin-1 in Atrophying Muscles. Northern blot analysis of total RNA from various tissues of control and
food-deprived mice revealed three major transcripts (≈2.5, 5.0, and
7.3 kb) (Figs. A and
). In fed controls, gastrocnemius
muscles and to a much lesser extent heart were the only tissues
expressing significant amount of these transcripts. Liver, kidney,
brain, spleen, and testis (data not shown) contained only trace
amounts. In addition to being restricted to striated muscle normally,
Northern blot analysis showed that after fasting, the levels of all
three transcripts increased at least sevenfold in skeletal muscles but
not in heart or other tissues (Fig. A). The increase in
atrogin-1 mRNA is among the largest we have identified for any gene in
different types of atrophying muscle (see below). For example,
polyubiquitin mRNA, whose expression has served as the benchmark for
activation of Ub-mediated proteolysis in atrophying muscles, is
increased typically 2- to 5-fold in muscles during fasting ( 28, 29).
To test whether the induction of atrogin-1 occurs before and may thus
trigger the loss of muscle tissue mass, we examined the time course of
its gene expression in the gastrocnemius muscle after food deprivation.
The level of atrogin-1 mRNA was clearly increased as early as 16 h
after food removal and reached a maximum level by 24 h, which was
sustained at 30, 48, and 72 h (Fig. B and data not
shown). In the same experiment, muscle weights (expressed as a
proportion of initial body weight as an indirect measure of muscle
weight loss) were compared between fasted and fed animals. In contrast
to the rapid rise in the level of atrogin-1 mRNA, no significant
decrease in the mass of the gastrocnemius muscle was detected in the
fasted animals until 30 h after food removal. At 30 h,
gastrocnemius weight (mg)/initial body weight (g) decreased by 8%
(from 5.52 in the fed to 5.09 in fasted animals) corrected for body
weight (P = 0.02, Mann–Whitney u test).
Similarly, in previous experiments with larger groups of mice of
similar age, significant muscle weight loss was always found 48 h
after food removal but not at 24 h, whereas atrogin-1 mRNA was
markedly increased at both these times (data not shown). Thus, the
increase in expression of atrogin-1 precedes muscle weight loss and
remains at a high level during the course of muscle atrophy. These
findings suggest that activation of this gene is important in the
development and progression of muscle protein loss in fasting. To test whether this gene is induced generally in atrophying muscles,
we analyzed the mRNA levels of atrogin-1 in muscles from a number of
rat models of human diseases in which there is also enhanced
proteolysis and marked muscle wasting. In atrophying muscles from rats
with streptozotocin-induced diabetes, rats bearing a peritoneal Yoshida
hepatoma, or rats with experimentally induced uremia, at times when
proteolysis by the ATP-dependent cytosolic pathway and Ub conjugation
are markedly accelerated ( 18– 20), the mRNA levels of atrogin-1 were
also increased at least 10-fold (Fig. ). Thus, atrogin-1 is a
muscle-specific gene, whose expression rises under diverse conditions
when muscle wasting occurs. By using cDNA microarrays to define the transcriptional changes in
atrophying muscle from food-deprived mice, we have discovered a new
muscle-specific F-box protein termed atrogin-1. Because atrogin-1 is
very strongly induced in many catabolic states, it is likely to play a
key role in the generation of muscle atrophy. The strong induction of
atrogin-1 expression at an early stage of muscle wasting, in fact
before muscle weight loss was detectable, and the maintenance of its
high expression during the period when overall proteolysis is
accelerated, strongly suggest a role in both initiation and maintenance
of the accelerated proteolysis. The signals that trigger the loss of muscle mass in the varied
pathological states modeled in this study differ. For instance,
glucocorticoids and low insulin levels are essential for the enhanced
proteolysis and muscle wasting in starvation and diabetes ( 28, 30, 31),
whereas TNF-α, prostaglandins, and glucocorticoids play important
roles in muscle wasting in experimental models of sepsis and
cancer-induced cachexia, including that induced by the Yoshida hepatoma
( 32– 35). Acidosis and glucocorticoids have been shown to be important
in the enhancement of muscle proteolysis in uremic animals ( 36, 37). By
themselves, glucocorticoids can stimulate protein breakdown in muscle
( 35), and in related studies, we have also found a marked up-regulation
of atrogin-1 mRNA on treatment of animals or cultured muscle cells with
glucocorticoids and also increased atrogin-1 mRNA in rat muscles
undergoing atrophy because of unloading by hind-limb suspension
(unpublished observations). Despite the different physiological
signals, the induction of atrogin-1 in all these catabolic states is
strong evidence for our suggestion that the muscle atrophy in these
diseases ( 2, 3, 10, 35) proceeds through a common set of adaptations
leading to enhanced proteolysis. The changes seen here in mRNA for atrogin-1 are much larger than those
reported by others and us for mRNAs encoding other components of this
proteolytic pathway. For example, the expression of polyubiquitin
increases in all forms of muscle atrophy; however this induction (two
to five times) is much smaller than the increase in atrogin-1.
mRNAs for Ub carrier protein (E2) genes, such as
E2 14k, are increased two- to threefold in
atrophying muscles ( 7), and similar increases may also occur with many
other E2s ( 38) (unpublished observations). mRNA for E3α, the Ub
protein ligase of the N-end rule pathway, is increased twofold in
muscles from diabetic and septic animals ( 7, 39), and this
ubiquitination pathway appears to contribute to the enhanced
proteolysis in these catabolic states ( 7, 8). Because atrogin-1 shares
no sequence homology with E3α, it is likely to act on different cell
proteins (see below). In SCF complexes, the F-box protein binds the substrate and links it to
the other subunits (RING-H 2, Cullin, and Skp1
proteins) and the E2 involved in ubiquitination. Accordingly, our
coprecipitation experiments showed that the F-box in atrogin-1 is
necessary for this protein to interact with the Skp1 protein because
coprecipitation from cell lysates did not occur when critical residues
in the F-box were mutated. Like certain other F-box proteins ( 40),
atrogin-1 is stabilized in vivo by its interaction with
Skp1. In fact, unless Skp1 was coexpressed with atrogin-1, only trace
amounts of this protein could be found in the cultured cells. Moreover,
atrogin-1, but not mutants in the F-box domain, assembled with Cul1,
Skp1, and Roc1 to form the minimal SCF module
(SCF Atrogin-1) characteristic of this family of
E3s. Most F-box proteins bind to substrates through regions C-terminal
to the F-box. Atrogin-1 may bind to substrates through the predicted
PDZ-binding motif at its extreme C terminus or through an as yet
undetermined region in its N-terminal portion. Other F-box proteins
with similar C-terminal F-box domains are known ( 26, 27), although
their substrate-binding domains have not been identified. In fact, in
related studies, we have shown that these complexes can ligate
125I-ubiquitin to form high molecular weight
conjugates (unpublished observation). Also of appreciable interest is the presence in atrogin-1 of a putative
nuclear localization signal. Many other F-box proteins also contain
nuclear localization sequences ( 41). The presence of such a sequence in
atrogin-1, along with its muscle-specific expression, suggests that it
may function in ubiquitinating muscle-specific transcription
factors or other nuclear proteins involved in muscle growth. However,
atrogin-1 may also function to ubiquitinate cytosolic proteins
such as components of the myofibril, which are rapidly degraded during
muscle atrophy. Previous studies have suggested that although diverse conditions can
lead to rapid muscle atrophy, the loss of muscle tissue in each case
results primarily from activation of a common biochemical program that
stimulates muscle proteolysis ( 2, 3, 10, 35). The identification of a
new E3 dramatically up-regulated in various types of atrophy further
strengthens the argument for such a common “atrophy program” and
for the existence of a group of atrophy-related genes. We will present
further evidence to support this concept in subsequent papers based on
transcriptional profile analysis of a variety of types of atrophying
muscles. Identification of the transcription factors that regulate
atrogin-1 expression will be an important step to understand the
atrophy response. Also, defining the substrates that atrogin-1
ubiquinates will be essential to clarify its role in the atrophy
process. Genetic manipulations or pharmacological agents that reduce
atrogin-1 activity might even prove useful in counteracting the
debilitating effects of muscle wasting. We thank Sandy Ryeom for her help and advice; Vickie Baracos for
providing muscles from tumor-bearing rats; James Bailey, Russ Price,
and William E. Mitch for providing muscles from diabetic and uremic
rats; and Dr. Edouard Vannier for providing the human muscle cDNA
library. We are also grateful to Dr. Keiji Tanaka and Tomiki Chiba for
providing cDNA plasmids and for advice on carrying out coprecipitation
experiments. This study was supported by grants from the National Space
Biomedical Research Institute and the Muscular Dystrophy Association
(to A.L.G.) and from the National Institutes of Health (DK02707 to
S.H.L.) During these studies, M.D.G. was a fellow of the
Fundaçâo de Amparo à Pesquisa do Estado de Sâo
Paulo. R.T.J. held a fellowship from US–UK Fulbright Commission and
the Warren-Whitman-Richardson Fellowship. | | Ub | ubiquitin | | | HA | hemagglutinin | | | GST | glutathione S-transferase |
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