From the Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, Texas 75235
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ABSTRACT |
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Thioacylation is one of a handful of reversible
covalent protein modifications, but the enzymes responsible for
addition and removal of long chain fatty acids from protein cysteine
residues in vivo have not yet been identified. The subunits of some heterotrimeric G proteins cycle between thioacylated
and deacylated states in a receptor-regulated fashion. We have
identified, purified, and characterized an enzyme acyl-protein
thioesterase that deacylates G
proteins and at least some other
thioacyl protein substrates, including Ha-RAS. The action of this
enzyme on thioacylated heterotrimeric Gs is regulated by
activation of the G protein. Although native and recombinant
acyl-protein thioesterases act as both acyl-protein thioesterases and
lysophospholipases in vitro, we demonstrate by transfection
that the enzyme can accelerate the turnover of thioacyl groups on
Gs
in vivo.
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INTRODUCTION |
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Protein function is regulated by dynamic covalent modifications of polypeptide backbones. Although phosphorylation of serine, threonine, or tyrosine residues is the most studied of these modifications, the importance of other alterations is increasingly appreciated; included are carboxymethylation of prenylcysteine residues, acetylation of lysine residues, and thioacylation of cysteine residues (1-3).
Thioacylation (palmitoylation or S-acylation) is the
addition of long chain fatty acids to specific cysteine residues within a protein by formation of a thioester bond. Almost all heterotrimeric G
protein (G
)1 subunits,
as well as a broad range of other membrane-associated proteins of
diverse biological function, are so modified (4, 5). Thioacylation and
other lipid modifications introduce significant hydrophobicity to a
protein, which can influence both protein-lipid and protein-protein
interactions. Unlike myristoylation, isoprenylation, or glypiation,
thioacylation is a dynamic process, and thioacylated proteins cycle
between acylated and deacylated states many times during their
existence within a cell (6). Regulation of thioacylation of proteins
involved in signal transduction is apparently widespread and has been
formally demonstrated in at least four cases as follows: endothelial
nitric oxide synthase, the
-adrenergic receptor, the m2 muscarinic
receptor, and the
subunit of the heterotrimeric G protein
Gs (7-10).
Reversible thioacylation of Gs has been well documented
in vivo. In an unstimulated cell, Gs
and
presumably other G
proteins exist primarily as thioacylated
(predominantly palmitoylated) GDP-bound heterotrimers. In this state
Gs
-bound thioacyl groups turn over with a
t1/2 of 20-90 min. Activation of Gs
by
-adrenergic agonists, cholera toxin, or mutation of residues
critical for GTP hydrolysis causes a dramatic (>10-fold) increase in
the rate of palmitate turnover (10-12). However, the deacylated state
is transient, because receptor stimulation apparently does not cause a
significant alteration in the stoichiometry of thioacylation of
Gs
(13).
Thioacylation is probably involved in several aspects of G
protein-mediated transmembrane signaling, although the interpretation of some functional studies performed in vivo is confounded
by the fact that mutation of certain relevant cysteine residues
per se can cause substantial functional deficits not due to
loss of thioacylation (14). In transfected cells, mutation of
thioacylated cysteine residues in Gs, Gq
,
and Go
causes incomplete localization to the plasma
membrane (15, 16). Such mutations in Go
, GPA1p (the G
protein in Saccharomyces cerevisiae), endothelial nitric oxide synthase, and Fyn kinase result in accumulation of the protein in
internal membranes2 (17-19).
Experiments performed in vitro have been hampered by difficulties in determining the stoichiometry of thioacylation. This
obstacle has recently been surmounted in two studies. Iiri and
colleagues (20) developed a chromatographic method to separate unmodified Gs
from its thioacylated counterpart. The
affinity of thioacylated Gs
for G
was higher than
that of the nonacylated protein. Taking advantage of the ability of
some G
proteins to autoacylate in vitro (21), Tu et
al. (22) demonstrated that thioacylated
subunits of some
Gi subfamily members were resistant to the GTPase
accelerating activity of regulators of G protein signaling
proteins.
Despite intensive investigation, the molecular machinery responsible
for the thioacylation cycle of intracellular proteins has not been
identified. Many thioacylated proteins, including G protein subunits, spontaneously incorporate thioacyl groups at the relevant
cysteine residues when incubated in vitro with an
appropriate acyl donor (e.g. palmitoyl-CoA) (21, 23, 24). Putative enzymatic activities capable of thioacylating G
proteins and Fyn kinase have also been characterized, although neither has been
purified to homogeneity (25, 26). A protein acyltransferase that acts
on farnesylated RAS was purified (27); however, this activity is due to
a recently characterized form of thiolase, called thiolase A, and
further studies will be needed to see if thiolase A palmitoylates RAS
in vivo.3
Determination of the respective roles of autoacylation and enzymatic thioacylation in vivo awaits identification of the relevant
enzymes.
Sorting impostors from the relevant players has also been a challenge
to those studying protein deacylation. Camp and Hofmann (28) purified
an enzyme (palmitoyl protein thioesterase; PPT1) capable of deacylating
both thioacylated RAS and G proteins. However, subsequent work
revealed that the enzyme is a lysosomal resident (29, 30), and
mutations in the PPT1 gene cause a fatal lysosomal storage disease,
infantile neuronal ceroid lipofuscinosis (31, 32). Affected individuals
accumulate unidentified cysteine-containing lipid moieties, presumably
the endogenous substrates of PPT1, within the cells of brain and other
tissues (33).
We report herein the isolation of a second acyl-protein thioesterase
(APT1). Identification of the gene encoding this enzyme revealed that
the protein had been purified previously as a lysophospholipase (34).
We demonstrate that thioacylated proteins are preferred substrates for
this enzyme in vitro and that it can act in vivo to deacylate Gs. We propose that APT1 represents the
first bona fide player in the regulated thioacylation of intracellular
proteins.
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EXPERIMENTAL PROCEDURES |
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Materials--
All chromatographic resins were purchased from
Amersham Pharmacia Biotech; isotopes were from NEN Life Science
Products; all chemicals were supplied by Sigma or Calbiochem, unless
otherwise noted. Antiserum P054, directed against whole PPT1, was a
gift from Dr. Sandra Hofmann. Polyclonal antiserum 584, directed
against Gs (35), and monoclonal ascites 12CA5, directed
against the HA epitope, were supplied by Dr. Susanne Mumby.
Methods-- Molecular biological procedures were carried out using standard methods (36). Qiagen products were used for DNA preparation. The polymerase chain reaction was performed using VENT thermostable polymerase (New England Biolabs) under conditions recommended by the manufacturer. Peptide sequencing of APT1 was carried out by Carolyn Moomaw and Steve Afendis in the Biopolymer Core Facility of The University of Texas Southwestern Medical Center.
Palmitoyl-Gi1
Preparation--
Myristoylated Gi
1,
synthesized in Escherichia coli and prepared as described by
Duncan and Gilman (21), was incubated with [3H]palmitoyl-CoA (700 cpm/pmol) in HMEC buffer (50 mM NaHEPES (pH 8.0), 2 mM MgCl2, 1 mM EDTA, 7.5 mM CHAPS) supplemented with 500 µM GDP at 30 °C for 1 h. An aliquot of this
reaction mixture was assayed for incorporation of palmitate into
protein (21); more than 90% of the Gi
1 in
the reaction was palmitoylated.
[3H]palmitoyl-Gi
1 was then
separated from [3H]palmitoyl-CoA by applying the reaction
mixture to a MonoQ HR 5/5 column equilibrated in HMEC buffer and
eluting with a gradient of NaCl (0-300 mM). The fractions
containing [3H]palmitoyl-Gi
1
(identified by liquid scintillation spectrometry and SDS-PAGE) were
concentrated in a Centricon 30 (Amicon) to 150 pmol/µl.
Palmitoyl-Gi1 Thioesterase
Assay--
This assay was adapted from one for palmitoyl-RAS
thioesterase described by Camp and Hofmann (28). Reactions were started by the addition of 10 µl of enzyme to 90 µl of
[3H]palmitoyl-Gi
1 (200-500
nM) in HMEC buffer supplemented with proteinase inhibitors
(leupeptin, 6 µg/ml; lima bean trypsin inhibitor, 6 µg/ml;
L-1-tosylamido-2-phenylethyl chloromethyl ketone, 32 µg/ml; 1-chloro-3-tosylamido-7-amino-2-heptanone, 32 µg/ml; and aprotinin, 2 µg/ml). The 100-µl reaction was incubated at 30 °C for 2-10 min and stopped by addition of 25 µl of stop solution (2%
SDS; 2 mg/ml cytochrome c). Unreacted substrate was
separated from free palmitate by precipitation with 500 µl of
isopropyl alcohol and centrifugation at 16,000 × g for
10 min. Free palmitate was measured by liquid scintillation
spectrometry of the supernatant. Data shown are averages of duplicate
reactions from representative experiments.
Purification of APT1 from Rat Liver Cytosol--
All
manipulations were performed at 4 °C, and all buffers were
supplemented with proteinase inhibitors as described above. Forty grams
of frozen rat liver (PelFreez Biologicals) was pulverized with a hammer
and then diluted with 400 ml of ice-cold HE buffer (50 mM
NaHEPES (pH 8.0), 1 mM EDTA). The mixture was blended with six 30-s pulses of a Polytron and then further disrupted by 20 strokes
in a Dounce homogenizer. This homogenate was filtered through
cheesecloth and centrifuged for 20 min at 1500 × g to remove large particulate matter. The resulting supernatant was centrifuged at 100,000 × g for 45 min, resulting in a
soluble fraction (S100) and pellet (P100). The P100 was suspended in
200 ml of HE buffer for assay. The S100 (500 ml) was supplemented with
200 ml of 3 M
(NH4)2SO4, and, after 10 min, the
mixture was again centrifuged at 100,000 × g for 45 min. The supernatant was applied to 100 ml of butyl-Sepharose FF
equilibrated in HE buffer containing 1 M
(NH4)2SO4. The column was eluted
with sequential 350-ml aliquots of HE buffer containing 500 or 250 mM or no (NH4)2SO4. Fractions containing thioesterase activity were diluted 10-fold in TE
buffer (50 mM Tris-HCl (pH 8.5), 1 mM EDTA) and
applied to 50 ml of Q-Sepharose FF equilibrated in TE buffer. The
flow-through was collected, and the column was subsequently eluted with
200 ml of TE buffer containing 1 M NaCl. The flow-through
fractions containing thioesterase were brought to 1 M
(NH4)2SO4 by slow addition of 3 M (NH4)2SO4. After
stirring for 15 min, the sample was centrifuged at 100,000 × g for 30 min, and the supernatant was applied to 40 ml of
phenyl-Sepharose HP packed in a 26/10 FPLC column. This column was
washed with 150 ml of TE buffer and then eluted with a 300-ml gradient
of ethylene glycol from 0 to 75% in TE buffer. Thioesterase-containing
fractions were dialyzed for 16 h against ME buffer (75 mM NaMES (pH 6.0), 1 mM EDTA) prior to
application to 12 ml of SP-Sepharose FF equilibrated in ME buffer; the
flow-through was collected. The column was eluted with a single 100-ml
application of ME buffer containing 1 M NaCl. Thioesterase
activity was found in both the flow-through and eluate. However,
immunoblots indicated that PPT1 was in the eluate (consistent with its
original purification); the flow-through was devoid of PPT1. The
flow-through was brought to 1 M
(NH4)2SO4 and then applied to 10 ml
of phenyl-Sepharose HP packed in a 16/10 column. The column was washed
with 40 ml of TE buffer and then eluted with a 75-ml gradient of
ethylene glycol (0-75%). Fractions containing thioesterase were
pooled and run on a Superdex-200 16/60 preparative gel filtration
column equilibrated in HMEC buffer (with no proteinase inhibitors).
Fractions containing thioesterase were concentrated to 100 ng/ml in a
Centricon 10 (Amicon), frozen in liquid N2, and stored at
80 °C.
Palmitoyl-CoA Hydrolase Assay-- [3H]Palmitoyl-CoA (15 µl; 200-1000 cpm/pmol) in HMEC buffer was added to 10 µl of enzyme, also in HMEC buffer; reactions were incubated at 30 °C for 2-15 min and stopped by addition of 200 µl of 6% phosphoric acid/CH3CN (1:9). Free palmitate was separated from palmitoyl-CoA by addition of 100 µl of toluene. After centrifugation for 1 min in a microcentrifuge, 100 µl of the organic (upper) phase was processed for liquid scintillation counting. Data shown are averages of duplicate reactions from representative experiments.
Lysophospholipase Assay-- [14C]Lysophosphocholine (lyso-PC) substrate was dried in a speed vac and suspended in HMEC buffer. This substrate was added to enzyme in the same buffer. After incubation at 30 °C for 10-30 min, free palmitate was separated from lyso-PC substrate as described by Lee et al. (37).
APT1 Expression in E. coli--
APT1 cDNA was amplified from
rat brain cDNA (CLONTECH) using the polymerase
chain reaction with the following primers:
5'-GCCCATGGGCGGCAACAACATGTCC-3' (sense) and
5'-GCAAGCTTTCATGGGGAAAGGTTTATACTCC-3' (antisense); note that
Cys2 of APT1 was mutated to Gly to facilitate cloning. The
amplified cDNA was digested with NcoI and
HindIII and ligated into the bacterial expression vector
pQE60-6H (38), which had been digested with the same restriction
enzymes. E. coli carrying pQE60-6HAPT1 or pQE60-6H-galactosidase were grown in 50 ml of LB containing 50 µg/ml ampicillin. When cultures reached an absorbance of 0.6, protein
synthesis was induced by addition of isopropylthiogalactoside to 200 µM. After 16 h at 30 °C, cells were harvested by
centrifugation, and the pellets were frozen in liquid N2.
The cells were thawed in 5 ml of TE buffer with proteinase inhibitors
and lysed with lysozyme (1 mg). Soluble lysate was generated by
treatment of the lysed cells with 100 ng of DNase, followed by
centrifugation at 100,000 × g for 30 min.
Stable Expression of APT1 in HEK293 Cells--
A vector for
expression of APT1 carrying two tandem copies of HSV-Tag at the amino
terminus was generated as follows. The polymerase chain reaction
was performed using 5'-GCGGTACCTTGCGGCAACAACATGTCCGCC-3' (sense) and
5'-GCCTCGAGTCATGGGGAAAGGTTTATACTCC-3' (antisense) primers. The product
was digested with KpnI and XhoI and ligated into
the vector pCDA3.1-HSV (provided by Elizabeth Duncan, University of
Texas Southwestern), such that the amino-terminal sequence, to
Cys2, of the HSV-tagged APT1 was as follows:
MQPELAPEDPEDQPELAPEDPEDIDGTVPC. HEK293 cells were transfected
with either pCDNA3.1-HSV-APT1 or pCDNA3.1-his3--galactosidase (Invitrogen) using the Cal-Phos Maximizer Transfection kit (CLONTECH) according to
the manufacturer's recommendations. The transfected cells were
switched to and maintained in selection media (Dulbecco's modified
Eagle's medium, 10% fetal calf serum, 5 units/ml penicillin, 5 µg/ml streptomycin, 4 µg/ml G418 (Life Technologies, Inc.)) after
30 h. Clonal cell lines were isolated using standard
techniques.
Metabolic Labeling of Gs--
Five 100-mm dishes
of HEK293
-galactosidase and HEK293-APT1 cells were
transfected with HA-
s-pCDNAI (15) using the Cal-Phos Maximizer Transfection Kit (CLONTECH). After
36 h, the cells were removed from the plates by washing with
Puck's saline G with 2 mM EDTA and pelleted by
centrifugation for 2 min at 500 × g. The pellet was
then suspended (106 cells/ml) in 10 ml of labeling media
containing Dulbecco's modified Eagle's medium, 10% fetal calf serum,
and 250 µCi/ml [3H]palmitate (50 Ci/mmol). For labeling
time courses, 1 ml of suspended cells was removed at each time point
and pelleted by centrifugation for 15 s in a microcentrifuge; the
labeling medium was aspirated, and the cell pellet was frozen
immediately in liquid N2. For pulse-chase experiments,
cells were labeled for 30 min and centrifuged for 2 min at 500 × g. The labeled cell pellet was then suspended in chase
medium (Dulbecco's modified Eagle's medium and 10% fetal calf serum,
supplemented with 50 µM unlabeled palmitate), and at
indicated times, the cells were harvested as described above.
Immunoprecipitation of HA-Gs--
Pellets from 1 ml of cell culture were suspended in 1 ml of RIPA buffer (50 mM NaHEPES (pH 7.4), 150 mM NaCl, 1% Nonidet
P-40, 0.5% sodium deoxycholate, 0.1% SDS) with proteinase inhibitors and solubilized by five passages through a 251/2 gauge needle. Debris
was removed by centrifugation in a microcentrifuge for 30 min at
4 °C. Monoclonal antibody 12CA5 ascites (1 µl) was added to each
solubilized extract. The extracts were incubated with rocking at
4 °C for 1 h. HA-Gs
-12CA5 complexes were
precipitated by addition of 25 µl of protein G-Sepharose and washed
three times with 500 µl of RIPA buffer. HA-Gs
was
eluted by addition of SDS-PAGE sample buffer (with 5 mM
dithiothreitol) and incubation for 3 min at 95 °C. The
immunoprecipitates were subjected to SDS-PAGE and transferred to
nitrocellulose. These filters were exposed to phosphorimaging analysis
(TR2040S imaging plates and BAS1500 scanner; Fuji Medical Systems)
after a 1-month exposure. The amount of Gs
present in
each sample was determined by immunoblotting with antiserum 584 (35)
and quantitation using NIH Image (version 1.61) software on a scanned
image of the film.
Metabolic Labeling and Separation of Cellular Lipids--
Cells
were incubated with [3H]palmitate as described for
metabolic labeling of Gs. At each time point, 100 µl
of culture was added to 1 ml of CHCl3/MeOH (1:1). The
lipids were extracted and separated by thin layer chromatography as
described by Patterson and Skene (39).
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RESULTS |
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Purified myristoylated Gi1 was
palmitoylated in vitro with [3H]palmitoyl-CoA
to provide a source of radiolabeled, thioacylated Gi
1 (21), a suitable substrate for palmitoyl
protein thioesterase assays similar to those previously described for
palmitoyl-RAS (28). To identify the relevant thioesterase activity, we
sought to separate it from the lysosomal protein exhibiting the same biochemical activity, PPT1. We chose rat liver as a source of thioesterase activity because of the relatively low abundance of PPT1
mRNA in this tissue (29). The majority of the thioesterase activity
was in the soluble fraction (S100) of rat liver homogenate (Fig.
1A). However, immunoblot
analysis of aliquots containing equal amounts of activity from the
particulate (P100) and S100 fractions with an antiserum directed
against PPT1 revealed that the membrane fraction contained
approximately four times more PPT1 than did the soluble fraction (Fig.
1B). The S100 fraction was further fractionated by
concanavalin A-Sepharose chromatography. Approximately 70% of the
soluble thioesterase activity flowed through the column, and this
fraction was devoid of immunologically detectable PPT1 (Fig. 1,
A and B). The enzyme(s) present in this flow-through fraction removed thioester-bound palmitate from
Cys3 of Gi
1 but failed to remove
amide-linked [3H]myristate from Gly2 of the
protein (data not shown). Separation of reaction products by thin layer
chromatography demonstrated that palmitate was the isopropyl
alcohol-soluble radiolabeled product measured in the thioesterase assay
(data not shown).
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The acyl-protein thioesterase activity not attributable to PPT1
(i.e. APT1) was purified from rat liver S100 by ammonium
sulfate precipitation and six chromatographic steps (Table
I). During the final step, Superdex-200
gel filtration chromatography, a single peak of thioesterase activity
comigrated with a single protein peak, measured by absorbance at 280 nm, with an apparent molecular mass of 29 kDa (Fig.
2A). A single silver-staining
band, with a mobility corresponding to a molecular mass of 27 kDa, was detected by SDS-PAGE in fractions containing this activity (Fig. 2B). All thioesterase activity (PPT1 and APT1) behaved
homogeneously at every step of this procedure with two exceptions.
First, PPT1 bound to SP-Sepharose, permitting separation of PPT1 and
APT1 (Fig. 2C). Second, a minor peak of thioesterase
activity (approximately 10% of that recovered) eluted between 30 and
35% ethylene glycol on the first phenyl-Sepharose column; the bulk of
the activity (including PPT1 and APT1) eluted between 40 and 55%
ethylene glycol (data not shown). The activity in the minor peak was
purified by subsequent chromatographic steps identical to those
described for the major peak. Although this protein exhibited a lower
specific thioesterase activity, it was indistinguishable by SDS-PAGE
and silver staining from the purified APT1 contained in the major peak
(data not shown). Sequencing of tryptic peptides from this protein
revealed it to be the same polypeptide as APT1 (data not shown). When
rat liver cytosol was depleted of PPT1 using concanavalin A-Sepharose,
detectable thioesterase activity migrated homogeneously on seven
separate chromatographic resins (data not shown). Furthermore, immunoblot analysis of particulate (P100) thioesterase activity and the
soluble activity that bound to concanavalin A-Sepharose indicates that
PPT1 accounts for virtually all membrane-associated palmitoyl-Gi1 thioesterase activity (Fig.
1B). We thus believe that PPT1 and APT1 account for
essentially all of the palmitoyl-Gi
1 thioesterase activity detected in rat liver homogenate.
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We tested the thioacyl protein substrate specificity of APT1 using
autoacylated Gi1 and
[3H]palmitoyl-RAS (isolated from Sf9 cells as
described by Camp and Hofmann (28)). Purified APT1 removed palmitate
from both proteins (Fig. 3A).
Because the stoichiometry of palmitoylation of Ha-RAS isolated in this
manner is unknown, detailed comparison of kinetic parameters for the
two substrates is not possible. However, the assays were performed at
substrate concentrations well below the KM of the
enzyme, demonstrated by linear increases in initial reaction rates with
increasing substrate concentration (data not shown). Under these
conditions, the catalytic efficiencies
(Vmax/KM) of the enzyme for
each substrate should be inversely proportional to the
t1/2 of substrate in the reaction. The catalytic
efficiency of APT1 for palmitoyl-Gi
1 was
roughly 3-fold higher than the value for palmitoyl-RAS.
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Activation of a heterotrimeric G protein in vivo by an
appropriately coupled receptor causes increased turnover of palmitate on the G protein subunit. Wedegaertner and Bourne (12) found that
addition of AlF4
, which activates
heterotrimeric G proteins, increased the rate of palmitate loss from
Gs
in broken cell preparations. We sought to confirm the
activation-dependent regulation of deacylation using
purified proteins. Gs heterotrimer was autoacylated as
described previously (21). When this palmitoylated protein was used as a substrate for APT1 in the presence or absence of
AlF4
, the activated, dissociated
subunit was deacylated more rapidly than the inactive heterotrimer
(Fig. 3B). Similar results were obtained with
Gi
1 (data not shown).
Palmitoyl-Gs
was never fully depalmitoylated in these
reactions; during substrate preparation some
palmitoyl-Gs
is denatured, and this protein is no longer a substrate for APT1. Since the rate of depalmitoylation of G
proteins by APT1 is not affected by the identity of the bound nucleotide (GDP or GTP
S; data not shown), the enhanced rate of depalmitoylation in vitro following activation is explained
by the fact that the free
subunit is a better substrate for APT1 than is the heterotrimer. Thus, the cycle of G protein subunit dissociation and association can apparently account for regulation of
G
depalmitoylation in vivo.
To identify the gene encoding APT1, the purified protein was subjected
to tryptic digestion and peptide sequencing. Three peptide sequences
were found in the deduced amino acid sequence of a recently purified
25-kDa rat lysophospholipase (Fig. 4)
(34). The mouse ortholog of this protein has also been cloned, again based on purification of a lysophospholipase activity (40). Data from
various genome sequencing projects allowed us to identify putative
orthologs of this protein in many species, including Schistosoma
mansoni, Saccharomyces cerevisiae, Caenorhabditis elegans, and Homo sapiens (Fig. 4). These orthologs
contain several highly conserved regions, including one between
residues 113 and 131 of the rat amino acid sequence. This region
contains a Gly-X-Ser-X-Gly motif found in a wide
variety of proteins that exhibit esterase activity. APT1 lacks the
sequence Gly-Asp-His near its carboxyl terminus; this sequence is found
in PPT1 and other enzymes with thioesterase activity (29). The
predicted molecular weight of 25,000 is consistent with the observed
migration of purified APT1 by SDS-PAGE and gel filtration
chromatography. Although there is similarity between lysophospholipid
and palmitoyl-Gi1 substrates (an
oxyester-bound fatty acid versus a thioester-bound fatty
acid), we wished to confirm the notion that APT1 has both
lysophospholipase and palmitoyl-Gi
1
thioesterase activity. The rat lysophospholipase gene was expressed in
E. coli as a hexahistidine-tagged fusion protein. Lysates
from bacteria expressing rat lysophospholipase contained substantially
higher activities of both lysophospholipase and
palmitoyl-Gi
1 thioesterase than did their
-galactosidase-expressing counterparts (Fig.
5). Purification of recombinant APT1
using Ni-NTA agarose and gel filtration chromatography indicated that this protein was indeed responsible for the observed activities (data
not shown).
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Because of the broad substrate specificity of APT1 in vitro,
we were unsure of its physiological substrate. To address this issue
biochemically, we assayed the activity of the enzyme while titrating
three different substrates presented in mixed detergent micelles (10 mM CHAPS) under otherwise identical conditions (Fig. 6). The apparent KM of
APT1 for the acyl protein substrate was 25- and 250-fold lower than the
values observed for palmitoyl-CoA or lysophosphocholine (lyso-PC),
respectively. In addition, the catalytic efficiency
(Vmax/KM) for the acyl
thioester substrates (palmitoyl-CoA and
palmitoyl-Gi1) was at least 200-fold higher
than that for the acyl ester containing
lyso-PC.4 Although the acyl
moiety of all three substrates presumably projects into the hydrophobic
core of the detergent micelles, the relative accessibility of each
substrate under these or physiological conditions (with substrates
present on the surface of a phospholipid bilayer) is not known. Thus,
under these assay conditions, acyl proteins are the preferred substrate
for APT1 in vitro.
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To demonstrate that palmitoyl-G proteins could be substrates for
this enzyme in vivo, we stably transfected 293 cells with plasmids carrying the rat APT1 cDNA fused to the HSV Tag, an
epitope derived from the herpes simplex glycoprotein D, or the E. coli
-galactosidase cDNA. Whole cell homogenates from these
cells were assayed for their ability to hydrolyze lyso-PC,
palmitoyl-Gi
1, and palmitoyl-CoA. The cells
transfected with APT1 showed roughly 2-fold elevations in APT and
lysophospholipase activity (Fig. 7A). However, there was no
detectable increase in palmitoyl-CoA thioesterase activity, presumably
because of the large number of palmitoyl-CoA hydrolase activities
within normal cells (41). The expression of APT1 was confirmed by
immunoblot analysis using a monoclonal antibody specific for the HSV
Tag epitope (data not shown). We used these cell lines to determine if
APT1 could act on thioacylated proteins in vivo. The
thioacylation of Gs
is well characterized; deacylation
of the protein is the rate-limiting step for incorporation of
exogenously added palmitate (10). These cell lines were transfected
with a cDNA encoding Gs
with an HA tag (15). After
incubation with [3H]palmitate, cells expressing APT1
exhibited a moderate increase in the rate of incorporation of label
into Gs
compared with cells expressing
-galactosidase
(Fig. 7B), consistent with the level of expression of APT1.
A similar increase in rate was observed when loss of radioactivity from
Gs
was monitored in four separate pulse-chase
experiments (Fig. 7C). We were concerned that the palmitoyl-CoA hydrolase activity of APT1 might increase the rate of
turnover of palmitoyl-CoA and cause an apparent increase in the rate of
Gs
deacylation. To be incorporated into cellular lipids,
exogenously added palmitate must first be incorporated into the
palmitoyl-CoA pool. We observed no change in the rate of incorporation
of [3H]palmitate into cellular lipids in cells
overexpressing APT1 (Fig. 7D). This set of experiments
demonstrates that APT1 is able to remove thioacyl groups from
Gs
in vivo.
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DISCUSSION |
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Isolation of enzymes based on biochemical activities can lead to the identification of molecules whose function is tangential to the activity in hand. The search for an enzyme responsible for deacylation of intracellular thioacylated proteins highlights two such tales. The first palmitoyl protein thioesterase activity to be purified and cloned was PPT1. A variety of data indicate that the role of this lysosomal enzyme is to hydrolyze cysteine-containing lipid products rather than to deacylate intracellular proteins (33). We have now purified and characterized a second palmitoyl protein thioesterase activity; APT1 had already been purified as a lysophospholipase (34).
Several lines of evidence suggest that intracellular deacylation of at
least some thioacylated proteins is the actual function of this enzyme.
During the purification of APT1, the
palmitoyl-Gi1 thioesterase activity was
purified at least 18,000-fold, and only one other species of activity
was observed (PPT1). During the purification of lysophospholipase (the
same polypeptide as APT1), lysophospholipase activity was enriched only
500-fold over the starting material because only one of many
characterized lysophospholipase activities was purified (34). We have
shown that thioacylated proteins are the preferred substrate of APT1
in vitro. APT1 has both a substantially lower
KM and a higher catalytic efficiency
(Vmax/KM) for
palmitoyl-Gi
1 substrate when compared with
lyso-PC.
In addition to these suggestive arguments, two other pieces of evidence
support our hypothesis that APT1 is an acyl-protein thioesterase
in vivo. First, the product of the phospholipase B gene
(PLB1) of S. cerevisiae has both phospholipase B activity and lysophospholipase activity. Disruption of PLB1 eliminates more than
99% of lysophospholipase activity, revealed by both biochemical assay
and metabolic labeling studies (37). These data suggest that the
putative lysophospholipase activity of S. cerevisiae APT1
contributes insignificantly to the total lysophospholipase activity
within this species. Finally, we have also demonstrated that
overexpression of APT1 leads to an increase in the basal rate of
turnover of palmitate in Gs. Studies on the effects of such overexpression on the turnover of thioacyl moieties bound to other
proteins are in progress.
When appropriate G protein-coupled receptors are activated, they
trigger at least a 10-fold increase in the rate of deacylation of
Gs; however, such stimulation has been shown to not
alter the stoichiometry of thioacylation of Gs
(10, 13).
It is thus unlikely that the 2-fold overexpression of APT1 that we
achieved by stable transfection causes a decrease in the steady-state
stoichiometry of thioacylation of G protein
subunits. Dramatic
overexpression of APT1 might lower the stoichiometry of G
thioacylation in vivo. However, it is important to note that
interpretation of results from studies of G protein signaling under
these conditions would be complicated by thioacylation of at least five
components of the G protein signaling machinery, including G
proteins themselves, seven transmembrane receptors, G protein-coupled
receptor kinases, adenylyl cyclases, and certain regulator of G protein
signaling proteins (42-45). Despite repeated attempts to isolate 293 cell clones with higher levels of expression of APT1, enzyme activity was never increased more than 3-fold and HSV-tagged protein, assayed by
immunoblotting, never varied by more than 2-fold among clones (data not
shown). These observations are consistent with toxicity resulting from
high levels of APT1 or regulation of APT1 protein levels
posttranscriptionally (a phenomenon noted by Sugimoto et al.
(34) while studying APT1 as a lysophospholipase). Dramatic overexpression of APT1 (e.g. with recombinant adenoviral
vectors) or targeting of overexpressed APT1 to specific protein
complexes (e.g. with APT1 fusion proteins) may provide a
tool for specifically probing the role of thioacyl cysteine residues in
cellular processes.
Genetic or pharmacological inactivation of APT1 will ultimately
demonstrate whether it is the only enzyme within the cell responsible
for deacylating G subunits or other thioacylated proteins. To this
end, putative orthologs of APT1 have been identified in mouse and
S. cerevisiae, systems commonly used for genetic manipulations. We have used targeted disruption to create S. cerevisiae strains lacking APT1. These yeast lack detectable
acyl-protein thioesterase activity, and we are currently characterizing
their phenotype.5 We believe
the identification of APT1 will allow many studies of the function of
thioacylation cycles that were previously not feasible.
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ACKNOWLEDGEMENTS |
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We thank Drs. Sandra Hofmann and Susanne Mumby for helpful discussion.
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FOOTNOTES |
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* This work was supported by National Institutes of Health Grants GM34497 and GM07062 and The Raymond and Ellen Willie Distinguished Chair in Molecular Neuropharmacology.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.
1
The abbreviations used are: G, heterotrimeric
G protein
subunit; G
, heterotrimeric G protein
and
subunit complex; PPT1, palmitoyl protein thioesterase 1; APT1,
acyl-protein thioesterase 1; GTP
S, guanosine
5'-3-O-(thio)triphosphate; lyso-PC,
L-
-lysophosphatidylcholine; HA, influenza hemagglutinin;
HSV, herpes simplex virus; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid, MES, 4-morpholineethanesulfonic acid.
2 Dr. Susanne Mumby, personal communication.
3 Dr. Michael Gelb, personal communication.
4
Partially purified hexahistidine-tagged APT1,
expressed in E. coli, exhibited substrate specificity
(Vmax/KM for palmitoyl-Gi1 > palmitoyl-CoA
lyso-PC) and specific activities in vitro similar to the
purified, nonrecombinant enzyme (data not shown).
5 J. A. Duncan and A. G. Gilman, unpublished observations.
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REFERENCES |
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