Activity of TSC2 is inhibited by AKT-mediated phosphorylation and membrane partitioning.
ABSTRACT Loss of tuberin, the product of TSC2 gene, increases mammalian target of rapamycin (mTOR) signaling, promoting cell growth and tumor development. However, in cells expressing tuberin, it is not known how repression of mTOR signaling is relieved to activate this pathway in response to growth factors and how hamartin participates in this process. We show that hamartin colocalizes with hypophosphorylated tuberin at the membrane, where tuberin exerts its GTPase-activating protein (GAP) activity to repress Rheb signaling. In response to growth signals, tuberin is phosphorylated by AKT and translocates to the cytosol, relieving Rheb repression. Phosphorylation of tuberin at serines 939 and 981 does not alter its intrinsic GAP activity toward Rheb but partitions tuberin to the cytosol, where it is bound by 14-3-3 proteins. Thus, tuberin bound by 14-3-3 in response to AKT phosphorylation is sequestered away from its membrane-bound activation partner (hamartin) and its target GTPase (Rheb) to relieve the growth inhibitory effects of this tumor suppressor.
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ABSTRACT: Mammalian target of rapamycin (mTOR), which is now referred to as mechanistic target of rapamycin, integrates many signals, including those from growth factors, energy status, stress, and amino acids, to regulate cell growth and proliferation, protein synthesis, protein degradation, and other physiological and biochemical processes. The mTOR-Rheb-TSC-TBC complex co-localizes to the lysosome and the phosphorylation of TSC-TBC effects the dissociation of the complex from the lysosome and activates Rheb. GTP-bound Rheb potentiates the catalytic activity of mTORC1. Under conditions with growth factors and amino acids, v-ATPase, Ragulator, Rag GTPase, Rheb, hVps34, PLD1, and PA have important but disparate effects on mTORC1 activation. In this review, we introduce five models of mTORC1 activation by growth factors and amino acids to provide a comprehensive theoretical foundation for future research.International Journal of Molecular Sciences 11/2014; 15(11):20753-20769. · 2.46 Impact Factor
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ABSTRACT: Meningiomas are by far the most common tumors arising from the meninges. A myriad of aberrant signaling pathways involved with meningioma tumorigenesis, have been discovered. Understanding these disrupted pathways will aid in deciphering the relationship between various genetic changes and their downstream effects on meningioma pathogenesis. An understanding of the genetic and molecular profile of meningioma would provide a valuable first step towards developing more effective treatments for this intracranial tumor. Chromosomes 1, 10, 14, 22, their associated genes, and other potential targets have been linked to meningioma proliferation and progression. It is presumed that through an understanding of these genetic factors, more educated meningioma treatment techniques can be implemented. Future therapies will include combinations of targeted molecular agents including gene therapy, si-RNA mediation, proton therapy, and other approaches as a result of continued progress in the understanding of genetic and biological changes associated with meningiomas. This review provides an overview of the current knowledge of the genetic, signaling and molecular profile of meningioma and possible treatments strategies associated with such profiles.Journal of neurology and neurosurgery. 01/2014; 1(1).
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ABSTRACT: TSC1 and TSC2 mutations cause neoplasms in rare disease pulmonary LAM and neuronal pathfinding in hamartoma syndrome TSC. The specific roles of TSC1 and TSC2 in actin remodeling and the modulation of cell motility, however, are not well understood. Previously, we demonstrated that TSC1 and TSC2 regulate the activity of small GTPases RhoA and Rac1, stress fiber formation and cell adhesion in a reciprocal manner. Here, we show that Tsc1-/- MEFs have decreased migration compared to littermate-derived Tsc1+/+ MEFs. Migration of Tsc1-/- MEFs with re-expressed TSC1 was comparable to Tsc1+/+ MEF migration. In contrast, Tsc2-/- MEFs showed an increased migration compared to Tsc2+/+ MEFs that were abrogated by TSC2 re-expression. Depletion of TSC1 and TSC2 using specific siRNAs in wild type MEFs and NIH 3T3 fibroblasts also showed that TSC1 loss attenuates cell migration while TSC2 loss promotes cell migration. Morphological and immunochemical analysis demonstrated that Tsc1-/- MEFs have a thin protracted shape with a few stress fibers; in contrast, Tsc2-/- MEFs showed a rounded morphology and abundant stress fibers. Expression of TSC1 in either Tsc1-/- or Tsc2-/- MEFs promoted stress fiber formation, while TSC2 re-expression induced stress fiber disassembly and the formation of cortical actin. To assess the mechanism(s) by which TSC2 loss promotes actin re-arrangement and cell migration, we explored the role of known downstream effectors of TSC2, mTORC1 and mTORC2. Increased migration of Tsc2-/- MEFs is inhibited by siRNA mTOR and siRNA Rictor, but not siRNA Raptor. siRNA mTOR or siRNA Rictor promoted stress fiber disassembly in TSC2-null cells, while siRNA Raptor had little effect. Overexpression of kinase-dead mTOR induced actin stress fiber disassembly and suppressed TSC2-deficient cell migration. Our data demonstrate that TSC1 and TSC2 differentially regulate actin stress fiber formation and cell migration, and that only TSC2 loss promotes mTOR- and mTORC2-dependent pro-migratory cell phenotype.PLoS ONE 10/2014; 9(10):e111476. · 3.53 Impact Factor
T H E J O U R N A L O F C E L L B I O L O G Y
© The Rockefeller University Press $8.00
The Journal of Cell Biology, Vol. 173, No. 2, April 24, 2006 279–289
Tuberous sclerosis complex (TSC), caused by loss of function
of either the TSC1 or -2 tumor suppressor genes, is an auto-
somal dominant disorder that leads to mental retardation, sei-
zures, and the formation of tumors in various organs, including
the brain, kidney, heart, and skin (Young and Povey, 1998;
Gomez et al., 1999; Cheadle et al., 2000). The TSC1 gene en-
codes the 130-kD protein hamartin (van Slegtenhorst et al.,
1997), and the TSC2 gene encodes the 198-kD protein tuberin
(The European Chromosome 16 Tuberous Sclerosis Consor-
tium, 1993). Hamartin contains two coiled-coil domains,
which have been shown to mediate binding to tuberin (Hodges
et al., 2001), forming a stable, functional tumor suppressor
heterodimer within cells (Plank et al., 1998; van Slegtenhorst
et al., 1998). Lesions that develop in TSC patients are histo-
logically diverse; however, the tumors that arise as a result of
loss of function of either TSC1 or -2 share common features,
suggesting that hamartin and tuberin function within the same
pathways to regulate cell cycle, cell growth, adhesion, and
vesicular traffi cking (van Slegtenhorst et al., 1998; Hengstschlager
et al., 2001). Recent studies have indicated that the hamartin–
tuberin heterodimer regulates cell growth and proliferation as
a downstream component of the phosphoinositide 3-kinase
(PI3K)–protein kinase B (PKB/AKT) signaling pathway,
which modulates signal transduction through target of
rapamycin (TOR) in both Drosophila melanogaster and
mammalian cells (Manning and Cantley, 2003; Inoki et al.,
2005). Several distinct yet complementary genetic and bio-
chemical studies collectively show that tuberin is a GTPase-
activating protein (GAP) for the small GTPase Ras homologue
enriched in brain (Rheb), which activates TOR and its down-
stream targets, such as the ribosomal S6 kinase (RSK; Li
et al., 2004a).
Although loss of tuberin promotes cell growth and
tumorigenesis, cells expressing tuberin must also be able to
relieve tuberin repression of mammalian TOR (mTOR) sig-
naling during conditions of mitogenic suffi ciency. In this re-
gard, tuberin contains multiple sites for AKT, MAPK, RSK,
and extracellular signal–regulated kinase phosphorylation
(Dan et al., 2002; Liu et al., 2002; Manning et al., 2002; Li
et al., 2003; Tee et al., 2003; Roux et al., 2004; Ballif et al.,
2005). Although it is clear that activation of AKT blocks tu-
berin inhibition of TOR signaling (Inoki et al., 2002; Manning
et al., 2002; Potter et al., 2002), the mechanism by which
Activity of TSC2 is inhibited by AKT-mediated
phosphorylation and membrane partitioning
Sheng-Li Cai,1 Andrew R. Tee,2 John D. Short,1 Judith M. Bergeron,1 Jinhee Kim,1 Jianjun Shen,1 Ruifeng Guo,1
Charles L. Johnson,1 Kaoru Kiguchi,1 and Cheryl Lyn Walker1
1Department of Carcinogenesis, University of Texas MD Anderson Cancer Center, Smithville, TX 78957
2Division of Molecular Physiology, University of Dundee, Medical Sciences Institute/Wellcome Building Complex, Dundee DD1 5EH, Scotland, UK
oss of tuberin, the product of TSC2 gene, increases
mammalian target of rapamycin (mTOR) signaling,
promoting cell growth and tumor development.
However, in cells expressing tuberin, it is not known how
repression of mTOR signaling is relieved to activate this
pathway in response to growth factors and how hamartin
participates in this process. We show that hamartin co-
localizes with hypophosphorylated tuberin at the mem-
brane, where tuberin exerts its GTPase-activating protein
(GAP) activity to repress Rheb signaling. In response to
growth signals, tuberin is phosphorylated by AKT and
translocates to the cytosol, relieving Rheb repression.
Phosphorylation of tuberin at serines 939 and 981 does
not alter its intrinsic GAP activity toward Rheb but parti-
tions tuberin to the cytosol, where it is bound by 14-3-3
proteins. Thus, tuberin bound by 14-3-3 in response
to AKT phosphorylation is sequestered away from its
membrane-bound activation partner (hamartin) and its
target GTPase (Rheb) to relieve the growth inhibitory ef-
fects of this tumor suppressor.
Correspondence to Cheryl Lyn Walker: firstname.lastname@example.org
Abbreviations used in this paper: CIAP, calf intestinal alkaline phosphatase;
GAP, GTPase-activating protein; IGF-1, insulin-like growth factor-1; mTOR, mam-
malian TOR; PI3K, phosphoinositide 3-kinase; Rheb, Ras homologue enriched
in brain; RSK, ribosomal S6K; S6K, S6 kinase; TOR, target of rapamycin; TSC,
tuberous sclerosis complex.
The online version of this paper contains supplemental material.
JCB • VOLUME 173 • NUMBER 2 • 2006 280
AKT inactivates this tumor suppressor is unknown (Bjornsti
and Houghton, 2004). In addition, there are confl icting data re-
garding the subcellular localization of tuberin. For instance, in-
dependent studies report that tuberin can localize to the cytosol
(Nellist et al., 1999), the membrane/particulate (100,000 g)
fraction (Wienecke et al., 1995), and even the nucleus (Lou
et al., 2001) of cells.
In this study, we sought to determine the mechanisms by
which tuberin is regulated during cell growth. We found that
tuberin is localized in membrane and cytosol fractions but not
in nuclear fractions, and the translocation of tuberin from the
membrane to cytosol is regulated by AKT signaling in response
to growth factors. Phosphorylation of tuberin by AKT causes
tuberin to become sequestered by 14-3-3 proteins in the cytosol.
Mutation of two specifi c phosphorylation sites (S939 and S981)
prevents the cytosolic translocation of tuberin from cellular
membranes and results in a constitutively active protein that
inhibits mTOR signaling. Importantly, tuberin phosphorylation
by AKT does not affect its GAP activity toward Rheb in vitro
but promotes Rheb-induced S6 kinase (S6K) 1 activation
through increased Rheb-GTP loading in vivo. Therefore, it is
likely that AKT phosphorylation inhibits tuberin as a result of
14-3-3 binding and cytosolic translocation rather than by im-
pairing its catalytic GAP activity.
Tuberin localization is regulated by Ser/Thr
phosphorylation in response to growth
To investigate the subcellular localization of tuberin, mouse
fi broblast (NIH3T3, Swiss3T3), rat kidney epithelial (TRKE2),
human embryonic kidney (HEK293), and human breast cancer
(MCF7) cell lines were fractionated, revealing that tuberin was
detected within both the cytosolic and membrane fractions but
not within the nuclear fraction (Fig. 1 A). Interestingly, tuberin
in the membrane fraction migrated faster than tuberin within the
cytosolic fraction (Fig. 1 A).
To determine whether this mobility shift was due to
changes in phosphorylation, we treated these subcellular frac-
tions with either a Ser/Thr or Tyr phosphatase (calf intestinal
alkaline phosphatase [CIAP] or YOP protein tyrosine phospha-
tase, respectively). After CIAP treatment, the mobility of tuberin
in the cytosol increased, resolving as a faster migrating band
similar to tuberin purifi ed from membrane fractions (Fig. 1 B).
However, YOP did not affect tuberin mobility, indicating that the
mobility shift was primarily due to phosphorylation of Ser/Thr
residues. Treatment with CIAP resulted in both membrane and
cytosolic tuberin migrating faster, indicating that tuberin within
Figure 1. Tuberin localization is determined
by Ser/Thr phosphorylation. (A) Western an-
alyses of subcellular fractions from the indi-
cated cell lines using an anti-tuberin antibody.
β1-integrin and lamin A/C proteins were
analyzed as fractionation controls. Similar re-
sults were also observed with canine kidney
cell lysates (not depicted). (B) Fractionated
NIH3T3 lysates were treated with either CIAP
serine/threonine phosphatase or YOP tyro-
sine phosphatase at 30°C for 1 h, and the
electrophoretic mobility of tuberin was deter-
mined by Western analysis. (C) Swiss3T3 fi -
broblasts were serum starved (SV) for 24 h
followed by serum stimulation (SM) with 20%
FBS for the indicated times, and fractionated
lysates were immunoblotted with the indicated
antibodies. Phospho-AKT (S473) and phospho-
AKT (T308) were blotted as controls for
the effectiveness of the indicated treatments.
Total lysates (top, M + C) were analyzed as
controls for tuberin levels. Tuberin protein lev-
els within the membrane and cytosolic frac-
tions were determined as the percentage of
total tuberin detected within both fractions.
A representative blot from two independent
biological replications is shown. Quantitation
obtained from quintuplicate runs (n = 5) from
each biological replicate is shown below.
(asterisks) Both the reduction in the membrane
fraction and increase in cytosolic fraction of
tuberin that occurred in response to serum
stimulation for 1 h were signifi cantly different
than starvation controls (t test, P < 0.02) and
the 6 h time point (t test, P < 0.01). (D, top) Western analyses of tuberin from membrane and cytosolic fractions of serum-starved MCF7 cells (24 h) that
were stimulated for 1 h with 30 ng/ml IGF-1 in the absence or presence of 100 nM of the PI3K inhibitor wortmannin. The vertical line indicates non-
adjacent lanes in a single blot. (bottom) Tuberin protein levels within the membrane and cytosolic fractions were determined as the percentage of total
tuberin detected within both fractions. A representative blot from three independent biological replications is shown. Quantitation is obtained from a to-
tal of seven independent Western blots from the three experiments. (asterisks) Both the reduction in the membrane fraction and increase in cytosolic frac-
tion of tuberin that occurred in response to IGF-1 stimulation were signifi cantly different from starvation controls (t test, P < 0.01) and the + wortmannin
(t test, P < 0.01).
REGULATION OF TUBERIN BY INTRACELLULAR SHUTTLING • CAI ET AL.281
the cytosolic fraction is hyperphosphorylated. In addition, the
serum-induced decrease in mobility of tuberin in both membrane
and cytosolic fractions is likely driven by multiple phosphoryla-
tion events, as CIAP treatment (removing all Ser/Thr phosphor-
ylation) increased the mobility of tuberin in both fractions.
To determine whether growth factor stimulation could
also alter tuberin phosphorylation and subcellular localization,
Swiss3T3 or MCF7 cells were serum starved and then stimu-
lated with serum or insulin-like growth factor-1 (IGF-1) before
fractionation (Fig. 1, C and D; and Fig. S1, available at http://
www.jcb.org/cgi/content/full/jcb.200507119/DC1), as were
NIH3T3 (Fig. S1 A) and TRKE2 cells (Fig. S1 B). Like serum,
IGF-1 increased the amount of tuberin in the cytosolic fraction
relative to starvation conditions (Fig. 1, C and D). In serum-
stimulated cells, phosphorylation of membrane-localized tu-
berin was also increased after 1 h (comparable to cytosolic
tuberin in starved cells but less than cytosolic tuberin from serum-
stimulated cells; Fig. 1 C), suggesting that phosphorylation at
specifi c residues, rather than total levels of phosphorylation,
was determining localization. By 6 h, tuberin became predomi-
nantly membrane localized, which correlated with a decrease in
AKT activation (Fig. 1 C). Translocation of tuberin from the
membrane to the cytosol was blocked by the PI3K inhibitors
wortmannin and LY294002 (Fig. 1 D and Fig. S1 B), implicat-
ing PI3K signaling in tuberin localization to the cytosol.
Tuberin translocation is mediated
by AKT phosphorylation
To determine whether AKT directed tuberin’s subcellular local-
ization, tuberin from membrane and cytosolic compartments of
NIH3T3 cells treated with IGF-1 or EGF was immunoprecipi-
tated and detected with a (S/T) phosphosubstrate antibody that
recognizes the consensus phosphorylation site for AKT and RSK
containing phospho-Ser/Thr with Arg at position −5 and −3
(RXRXXpS/T) (Alessi et al., 1996; Obata et al., 2000; Yaffe
et al., 2001; Roux et al., 2004). With equal tuberin loading (Fig. 2,
A and B), the (S/T) phosphosubstrate antibody predominantly
recognized tuberin within the cytosolic fraction, and the PI3K
inhibitor wortmannin signifi cantly reduced recognition of phos-
phorylated tuberin, suggesting that cytosolic tuberin was phos-
phorylated by AKT (Fig. 2, A and B). To confi rm that activation
of AKT directed tuberin to the cytosol, MCF7 cells stably trans-
fected with a constitutively active AKT (myr-AKT) were exam-
ined and found to contain more cytosolic tuberin relative to
wild-type MCF7 cells (Fig. 2 C). Recently, RSK was shown to
phosphorylate tuberin at S1798 (Roux et al., 2004). To deter-
mine whether RSK phosphorylation of tuberin could regulate
its localization, we generated a COOH-terminal deletion mutant
of TSC2 at residue 1734 (referred to as ∆73) that lacks S1798
(Roux et al., 2004). ∆73 and wild-type tuberin had a similar
distribution within membrane and cytosolic fractions under
normal growth conditions (Fig. S1 C) and in response to serum
(Fig. S2 A, available at http://www.jcb.org/cgi/content/full/
jcb.200507119/DC1). In addition, overexpression of RSK1 did
not affect localization of wild-type tuberin (Fig. S2 B). Collec-
tively, these data indicate that tuberin residing in the cytosol is
phosphorylated by AKT, suggesting that AKT (but not RSK)
directly controls tuberin’s localization and possibly its activity.
Mutation of AKT phosphorylation sites
in tuberin alters its localization
Tuberin contains multiple S/T phosphorylation sites (Fig. 3 A).
Among these, S1254 has been shown to be phosphorylated by
MK2 (Li et al., 2003), whereas S939 (Inoki et al., 2002;
Figure 2. AKT-mediated phosphorylation of tuberin leads to subcellular translocation. (A) NIH3T3 cells were treated with 100 ng/ml EGF in the absence
or presence of 200 nM wortmannin for 1 h. The fractionated cell lysates were immunoprecipitated with an anti-tuberin antibody (N19) followed by Western
blot analysis with an anti–phospho-(S/T) AKT substrate antibody and an anti-tuberin antibody (C20). (B) NIH3T3 cells were serum starved for 24 h followed
by treatment with 30 ng/ml IGF-1 in the absence or presence of 100 nM wortmannin for 1 h. The fractionated cell lysates were immunoprecipitated with
an anti-tuberin antibody (N19) followed by Western blot analysis with an anti–phospho-(S/T) AKT substrate antibody and an anti-tuberin antibody (C20).
Phospho-AKT (S473) was blotted as a control for the effectiveness of the indicated treatments. The vertical line indicates nonadjacent lanes in a single blot.
(C) Western analyses of tuberin, AKT, and phospho-AKT (S473) were performed using membrane and cytosolic fractions from wild-type MCF7 and MCF7
cells that were stably transfected with constitutively active AKT (Myr-AKT). β1-integrin and LDH proteins were used as fractionation controls.
JCB • VOLUME 173 • NUMBER 2 • 2006 282
Manning et al., 2002), S981 (Dan et al., 2002), S1130/S1132
(Inoki et al., 2002), and T1462 (Inoki et al., 2002; Manning et al.,
2002) have been reported to be AKT phosphorylation sites in vivo
and/or in vitro, and several of these also have the potential for
binding to 14-3-3 (http://scansite.mit.edu; Yaffe et al., 2001). S981
is of particular interest, as it lies within the alternatively spliced
exon 25 of tuberin. To examine these as candidate sites for the
regulation of tuberin localization, we mutated these residues to
alanine to create TSC2 constructs in frame with an NH2-terminal
Flag epitope (Flag-TSC2): S939A, S981A, and 2A (S939A +
S981A); T1462A and SATA (S939A + T1462A; Manning et al.,
2002); and S1254A and S1130A + S1132A double mutant.
Wild-type and mutant TSC2 constructs were transfected
into HEK293 cells and subjected to subcellular fractionation.
As shown in Fig. 3 (B and C), S939A, S981A, and 2A mutants
predominantly localized to the membrane, as did the double
alanine SATA mutant that lacks the S939 site. However, the
T1462A single mutant partitioned in the cell similarly to wild-
type tuberin (Fig. 3 B), indicating that T1462 phosphorylation
was not directing translocation of tuberin from the membrane to
the cytosol. These data were confi rmed with a phosphospecifi c
T1462 antibody, which recognized tuberin in both the mem-
brane and cytosolic fractions equally (unpublished data).
Furthermore, phosphorylation at both S939 and S981 contributes
to cytosolic localization, as phosphorylation at S939 (determined
with a phospho-S939 specifi c antibody) of the S981A mutant
was not suffi cient to partition tuberin to the cytosol (Fig. S2 C).
Other tuberin mutants, S1254A, S1130A/S1132A (Fig. 3 C),
∆73, or S1338A (Fig. S1 C), also distributed equally between
the membrane and cytosolic fractions. Thus, S939 and S981
phosphorylation are critical determinants of membrane versus
cytosolic localization of tuberin.
14-3-3 proteins mediate translocation
of tuberin into the cytosol
14-3-3 has been previously reported to directly interact with
phosphorylated tuberin (Li et al., 2002; Nellist et al., 2003).
S939 and S981 are predicted AKT phosphorylation and 14-3-3
interaction sites. When phosphorylated and nonphosphorylated
S939 and S981 peptides were used in competition assays to
block GST–14-3-3 interaction with tuberin, as shown in Fig. 4 A
and Fig. S3 (available at http://www.jcb.org/cgi/content/full/
jcb.200507119/DC1), phosphorylated but not nonphosphory-
lated S939 and S981 peptides clearly competed for the inter-
action of tuberin with several 14-3-3 isoforms. In addition, the
amount of 2A mutant tuberin affi nity purifi ed by 14-3-3 was
dramatically reduced relative to wild-type tuberin (Fig. 4 B).
Importantly, inhibition of PI3K signaling by wortmannin ab-
lated this 14-3-3 tuberin interaction (Fig. 4 B). These data indi-
cate that S939 and S981 are critical sites of interaction between
tuberin and 14-3-3 proteins and that this interaction is mediated
by PI3K/AKT phosphorylation.
To demonstrate that tuberin binding to 14-3-3 was directly
responsible for its translocation to the cytosol, we transfected
HEK293 cells with the EGFP-R18 construct that expresses a
peptide that disrupts 14-3-3 binding (Jin et al., 2004). As shown
in Fig. 4 C, the R18 14-3-3 decoy clearly repressed cytosolic
localization of tuberin, establishing a direct link between 14-3-3
and translocation of tuberin to the cytosol.
Hamartin enhances tuberin retention
at the membrane
Hamartin possesses a predicted transmembrane domain and
two coiled-coil domains that mediate its association with tu-
berin (van Slegtenhorst et al., 1997, 1998). We found that in
both human and mouse cells, hamartin was only detected in the
membrane fraction (Fig. 5, A and B; Fig. 3 B; unpublished
data). To confi rm that tuberin mutants that did or did not consti-
tutively localize to the membrane retained their ability to bind
hamartin, immunoprecipitation experiments were performed.
Immunoprecipitation showed that, similar to wild-type tuberin,
tuberin S939A, S981A, S1338A, ∆73, and 2A mutants retained
their ability to interact with hamartin (Fig. S4, A, B, and C, avail-
able at http://www.jcb.org/cgi/content/full/jcb.200507119/DC1).
To determine whether hamartin played a role in the subcel-
lular localization of tuberin, we transfected wild-type, S939A,
Figure 3. Identifi cation of S939 and S981 as phosphorylation sites that
determine tuberin localization. (A) Schematic of tuberin phosphorylation
sites with Flag-TSC2 constructs and their corresponding mutations listed
below. (B) Western analysis of HEK293 membrane and cytosolic fractions
using an anti-Flag antibody after transfection with wild-type and mutant
Flag-TSC2 constructs. (C) Wild-type and mutant Flag-TSC2 constructs were
overexpressed in HEK293 cells, and the membrane and cytosol lysates
were immunoblotted with an anti-Flag antibody. The vertical line indicates
nonadjacent lanes in a single blot.
REGULATION OF TUBERIN BY INTRACELLULAR SHUTTLING • CAI ET AL.283
and S981A Flag-TSC2 constructs into HEK293 cells with
or without Myc- or Flag-TSC1. Although S939A and S981A
mutants were primarily membrane localized, coexpression of
hamartin increased membrane retention of both mutant and
wild-type tuberin in a dose-dependent manner (Fig. 5, A and B).
These data suggest that the interaction of hamartin with
tuberin facilitates its localization to the membrane, implying
that the tuberin–hamartin heterodimer functions in this sub-
Importantly, only tuberin in the membrane fraction re-
mained associated with hamartin: no hamartin could be co-
immunoprecipitated with tuberin in the cytosolic fraction,
indicating that translocation to the cytosol dissociated tuberin
from hamartin (Fig. 5 C). Similarly, when Flag-TSC2 and
Myc-TSC1 were cotransfected into MCF7 cells, tuberin and
hamartin were observed by confocal microscopy to colocalize
in a discrete, punctate pattern (Fig. S4 D). Cells that expressed
wild-type Flag-TSC2 in the absence of Myc-TSC1 exhibited a
more diffuse staining pattern than cells that overexpressed both
Myc-TSC1 and Flag-TSC2 (Fig. S4 D, comparing γ and γ′ with
β and β′). However, the 2A mutant, which was constitutively
membrane localized as shown by cell fractionation, retained
this punctate localization pattern even in the absence of exoge-
nously expressed hamartin (Fig. S4 E).
Colocalization of tuberin and Rheb
is disrupted in response to growth
Tuberin’s GAP target Rheb is farnesylated and predicted to be
membrane localized (Clark et al., 1997), and recent data indicate
Figure 4. 14-3-3 proteins bind and sequester
tuberin in the cytosol. (A) GST–14-3-3 proteins
were used to affi nity purify proteins from
HEK293 cells in the presence of phosphory-
lated or nonphosphorylated S939 and S981
tuberin peptides. Affi nity-purifi ed complexes
were immunoblotted to detect the amount of tu-
berin interacting with 14-3-3. The vertical line
indicates nonadjacent lanes in a single blot.
(B) HEK293 cells were transfected with Flag-
TSC2-WT or Flag-TSC2-2A in the absence or
presence of 200 nM wortmannin. Western
analysis of exogenous tuberin was performed
after using GST–14-3-3 proteins to affi nity purify
Flag-tuberin. (C) HEK293 cells were transfected
with Flag-TSC2 in the presence or absence of
GFP-R18 14-3-3 decoy expression construct,
and fractionated lysates were used for Western
analyses with the indicated antibodies. LDH
was used as a fractionation control.
Figure 5. Hamartin increases the amount of
tuberin retained in the membrane. (A) HEK293
cells were transfected with wild-type and mu-
tant Flag-TSC2 constructs in the presence or
absence of a Myc-TSC1 construct. Cells were
fractionated and immunoblotted with indicated
antibodies. (B) HEK293 cells were transfected
with wild-type TSC2 constructs along with in-
creasing amounts of Flag-TSC1. Cells were
then fractionated and immunoblotted with indi-
cated antibodies. (C) HEK293 cells were
transfected with Flag-TSC2 and Myc-TSC1
constructs. The fractionated cell lysates were
immunoprecipitated with an anti-Flag antibody
followed by Western analysis with anti-Flag
(mouse) and anti-Myc (mouse). The same blot
was then reprobed with anti-tuberin (rabbit)
and anti-hamartin (rabbit) as indicated.
JCB • VOLUME 173 • NUMBER 2 • 2006 284
that Rheb is localized in endomembranes (Takahashi et al.,
2005). To determine whether phosphorylation at S939/S981
affected tuberin’s intrinsic GAP activity for Rheb, we fi rst de-
termined Rheb’s subcellular localization. Rheb was detected
only in the membrane fractions from HEK293 (Fig. 3 B) and
MCF7 (Fig. S5 A, available at http://www.jcb.org/cgi/content/
full/jcb.200507119/DC1) as well as NIH3T3, Swiss3T3, and
TRKE2 cells (not depicted). Treatment with EGF or wortmannin
did not affect this localization over a period of 10 min to 3 h
(Fig. S5 A and not depicted). Recognition of the 21-kD band in
the membrane fraction using this anti-Rheb antibody was spe-
cifi c for Rheb, as antibody binding was ablated when Rheb
RNAi was used to knock down Rheb (Fig. S5 B). When un-
tagged TSC1/Flag-TSC2 and Myc-Rheb were cotransfected into
HeLa cells, tuberin and Rheb were observed with confocal
microscopy to colocalize in a discrete, punctate pattern in the
absence of serum (Fig. 6). In contrast, stimulation with IGF-1
resulted in partitioning of wild-type tuberin away from Rheb
(Fig. 6), indicating that in response to growth factor stimulation,
tuberin is no longer retained in physical proximity to its down-
stream target, Rheb. Collectively, these data indicate that the
localization pattern of tuberin is modulated by both hamartin
and growth factor signaling and that Rheb is in physical prox-
imity to membrane-localized tuberin and hamartin within the
cell. In addition, these data, along with confocal microscopy
using 2A mutant tuberin and biochemical fractionation, sug-
gest that S939 and S981 are crucial sites for determining the
sub cellular localization and, thus, function of tuberin in re-
sponse to growth factor signaling.
Tuberin localization regulates its function
To elucidate the relationship between tuberin localization and
function, we examined the effect of wild-type, S939A, S981A,
and 2A tuberin mutants on mTOR-S6K activity. T389 is a
known rapamycin-sensitive phosphorylation site of S6K that
correlates with activation by mTOR (Pearson et al., 1995).
In cells transfected with HA-tagged S6K, cotransfection of
wild-type TSC1 and -2 expression constructs diminished phos-
phorylation of exogenously expressed S6K (Fig. 7 A). However,
cotransfection of the 2A mutant with TSC1 resulted in an even
more dramatic reduction in phospho-S6K levels (Fig. 7 A).
Similar experiments were performed to determine the effect of
S939A and S981A single and double mutants on phosphoryla-
tion of endogenous S6K. In the presence of serum, wild-type
Flag-tuberin was located in the membrane and cytosol, whereas
S939A, S981A, and 2A Flag-tuberin mutants remained primar-
ily at the membrane (Fig. 7 B, top). AKT was equally activated
by serum in cells expressing wild-type or mutant tuberin con-
structs, and endogenous hamartin remained membrane local-
ized (Fig. 7 B). In cells transfected with wild-type tuberin,
phosphorylation of endogenous S6K at T389 increased in re-
sponse to serum, coincident with translocation of tuberin to the
cytosol. However, in cells transfected with S939A, S981A, or
2A tuberin mutants, phosphorylation of S6K was inhibited,
with the 2A mutant most effectively blocking S6K activation
(Fig. 7 B). Thus, tuberin mutants that were retained at the mem-
brane were constitutively active and exhibited enhanced ability
to repress S6K activation. These data suggest that AKT phos-
phorylation of tuberin regulates its activity by decreasing the
amount of tuberin located at the membrane, thereby reducing its
inhibitory effect on the Rheb–mTOR–S6K signaling pathway.
Phosphorylation sites that determine
localization and function of tuberin
do not change its intrinsic RhebGAP
activity in vitro
Physical sequestration of tuberin from its target Rheb suggested
a mechanism whereby AKT modulated tuberin by partitioning
it away from the membrane rather than altering its intrinsic
GAP activity for Rheb. We compared the relative RhebGAP
activity of wild-type TSC2 and 2A mutant (S939A and S981A)
in cells during activation of the PI3K–AKT pathway (Fig. 8).
Figure 6. Tuberin colocalization with Rheb is
disrupted in response to growth factor stimula-
tion. HeLa cells were transfected in serum-free
media with Flag-TSC2 WT and Myc-Rheb
along with an untagged TSC1 construct. After
12 h without serum, cells were treated with
30 ng/ml IGF-1 for 1 h. Localization of exoge-
nous Flag-tuberin (green) and Myc-Rheb (red)
was detected by double immunofl uorescence
labeling and confocal microscopy. Colocaliza-
tion is detected in the merged image with the
yellow signal (right).
REGULATION OF TUBERIN BY INTRACELLULAR SHUTTLING • CAI ET AL.285
To do this, we quantifi ed the ratio of GTP and GDP Myc-Rheb
(percentage of GTP bound Myc-Rheb) when these TSC2 con-
structs were coexpressed during a time course of insulin
stimulation. The 2A mutant enhanced the GTPase function of
Rheb more effectively than wild-type tuberin, as indicated by
impaired accumulation of the active GTP form of Rheb after
15 min of insulin stimulation (32% GTP bound) when compared
with wild-type TSC2 (42% GTP bound; Fig. 8 A). As an in-
crease in GTP bound Rheb would be predicted to enhance
mTOR-mediated cell signaling, we measured the activity of
HA-S6K1 in these cells (Fig. 8 B). As expected, insulin-induced
activation of S6K1 was markedly impaired in cells expressing
the 2A mutant with the 2A mutant blocking insulin-induced ac-
tivation of S6K1 by 50% (after 15 min) when compared with
wild-type TSC2 (Fig. 8 B), refl ecting the reduced levels of ac-
tive GTP bound Rheb (Fig. 8 A).
To determine whether phosphorylation at S939 and S981
had an impact on tuberin’s intrinsic GAP activity, we analyzed the
ability of wild-type or mutant tuberin proteins to activate Rheb
GTPase in vitro. RhebGAP assays were conducted on immuno-
precipitated tuberin–hamartin heterodimers containing either
the wild-type or AKT-phosphorylation mutants of TSC2 (Fig. 9).
Figure 8. AKT phosphorylation of tuberin promotes Rheb-induced S6K1
activation through increased Rheb-GTP loading. (A) HEK293 cells co-
expressing HA-S6K1, Myc-Rheb, Flag-TSC1, and either Flag-TSC2-WT or
Flag-TSC2-2A (S939A and S981A) were serum starved and, where indi-
cated, the PI3K–AKT signaling pathway was stimulated with 100 nM insu-
lin for 7.5 and 15 min. These cells were subjected to in vivo radiolabeling,
and the level of guanine nucleotide bound to immunoprecipitated Myc-
Rheb was quantifi ed. A representative blot from three independent biologi-
cal replications is shown. The mean of the percentage of total Myc-Rheb
bound to GTP (active state) is shown in the bar fi gure. (B) In parallel,
HEK293 cells treated as in A were subjected to S6K1 activity assays.
HA-S6K1 used in the activity assay was analyzed with an anti-HA antibody.
32P- incorporation into GST-S6 substrate was quantifi ed using a phospho-
rimager, and the fold activation of S6K1 is graphed. Protein levels of Flag-
TSC1, Flag-TSC2, and AKT, and level of AKT phosphorylation at S473,
were determined using Western analyses.
Figure 7. AKT-mediated phosphorylation of tuberin relieves repression of
mTOR-S6K signaling. (A) HEK293 cells were transfected with HA-S6K,
Myc-TSC1, and Flag-TSC2 constructs as indicated. Cells were serum
starved for 24 h and treated with or without 30 ng/ml IGF-1 for 1 h. Cell
lysates were immunoblotted with the indicated antibodies, and phopho-
S6K levels were compared with the amount of HA-S6K protein levels.
Phospho-AKT (S473) was blotted as a control for the effectiveness of IGF-1
treatment. (B) Western analyses of membrane and cytosolic fractions from
HEK293 cells transfected with wild-type or mutant Flag-TSC2 constructs.
After 9 h, serum was either removed or allowed to remain for 24 h. The
protein levels of phosphorylated endogenous S6K were compared with
total S6K protein levels. Phospho-AKT (S473) and phospho-AKT (T308)
were blotted as controls for the effectiveness of the indicated treatments.
JCB • VOLUME 173 • NUMBER 2 • 2006 286
Tuberin–hamartin complexes containing wild-type tuberin from
both unstimulated and insulin-stimulated cells enhanced the in-
trinsic GTPase activity of Rheb at comparable rates. Induction
of tuberin phosphorylation by insulin was confi rmed in these
lysates by detection of phospho-T1462 (Fig. 9, bottom).
Furthermore, tuberin mutants lacking these sites (S939A,
S981A, and 2A) also possessed similar RhebGAP activity to
wild-type tuberin in vitro, even after insulin treatment (Fig. 9).
These fi ndings imply that AKT-mediated phosphorylation of
tuberin does not directly alter the rates at which tuberin en-
hances the GTPase activity of Rheb, at least in vitro. However,
these data support a mechanism whereby AKT phosphorylation
suppresses tuberin function by translocating tuberin to the cyto-
sol away from its membrane-associated binding partner, hamar-
tin, and its downstream target, Rheb. Indeed, as the effi ciency
with which tuberin functions as a RhebGAP in vitro is signifi -
cantly reduced in the absence of hamartin, physical separation
from both hamartin and Rheb may be contributing to reduced
tuberin activity (Tee et al., 2002; Li et al., 2004b).
A mechanism by which tuberin function
is regulated through subcellular localization
Based on our data, we propose a model where in the absence of
growth stimulatory signals, hamartin facilitates localization of
hypophosphorylated tuberin to membranes in physical proxim-
ity to Rheb. The membrane-associated tuberin–hamartin com-
plex binds Rheb and acts as a GAP to inactivate membrane
Rheb signaling by stimulating GTP hydrolysis (Fig. 10 A).
However, during mitogenic suffi ciency (i.e., after growth factor
stimulation), activation of PI3K signaling leads to activation of
AKT, which then directly phosphorylates membrane-associated
tuberin. In response to AKT phosphorylation, 14-3-3 proteins
bind tuberin and sequester it in the cytosol. The defi ciency of
membrane-associated tuberin results in the accumulation of
GTP bound Rheb, which leads to increased mTOR signaling,
enhanced cell growth, and proliferation (Fig. 10 B).
In this study, we provide evidence that AKT inhibits the tumor
suppressor function of tuberin by altering its subcellular
localization. Previously, AKT was shown to directly phosphorylate
and inhibit tuberin function upon stimulation with growth fac-
tors (Marygold and Leevers, 2002). However, it was not known
how AKT phosphorylation of tuberin regulated its function or
how hamartin contributed to tuberin regulation of Rheb and
mTOR (Garami et al., 2003; Tee et al., 2003; Zhang et al.,
2003). We have shown that although tuberin is found in both the
membrane and cytosolic fractions of cells, cytosolic tuberin ac-
cumulates upon growth factor stimulation and is hyperphos-
phorylated. The translocation of tuberin from the membrane
to the cytosol can be blocked by PI3K inhibitors, and AKT-
phosphorylated tuberin is predominantly found in cytosolic
fractions. The AKT phosphorylation sites S939 and S981 are
Figure 9. Tuberin’s GAP activity for Rheb in vitro is not affected by AKT
phosphorylation. HEK293 cells coexpressing Flag-TSC1 and Flag-TSC2-
WT, Flag-TSC2-S939A, Flag-TSC2-S981A, or Flag-TSC2-2A were serum
starved and then stimulated with 100 nM insulin for 30 min. TSC1 and -2
were immunoprecipitated from the lysates and subjected to RhebGAP activ-
ity assays for 15, 30, or 60 min. Immunocomplexes containing wild-type
and mutant tuberin activated Rheb similarly, as indicated by quantitation of
percentage of GDP. The vertical line indicates nonadjacent lanes resolved
on a single thin layer chromatography plate. For quality control of the
α-[32P]GTP used in the assay, the guanine nucleotide eluted from the 60-min
vector control was resolved on a separate thin layer chromatography plate
and shows that Rheb is 98% bound to GTP in the absence of hamartin and
tuberin. Each assay contained approximately equal amounts of TSC1 and -2,
and AKT activation was confi rmed with a phospho-T1462 antibody by
Western blot analysis (bottom).
Figure 10. Model for regulation of tuberin under conditions of mitogenic
suffi ciency. (A) Tuberin normally functions as a GAP for Rheb within an in-
tracellular membrane compartment, inhibiting Rheb and mTOR signaling,
which suppresses cell growth. (B) Upon stimulation by growth factors, AKT
is activated, leading to phosphorylation and cytosolic sequestration of
tuberin by 14-3-3 proteins. This cytosolic translocation relieves tuberin
repression of the Rheb-mTOR signaling, stimulating cell growth.
REGULATION OF TUBERIN BY INTRACELLULAR SHUTTLING • CAI ET AL.287
crucial residues that affect tuberin localization, with localiza-
tion of tuberin to the membrane accounting for the ability of tu-
berin to inhibit mTOR signaling via its GAP activity for Rheb.
We show that two tuberin phosphorylation sites, S939
and S981, are responsible for interaction with 14-3-3 in a
phosphorylation-dependent manner. 14-3-3 proteins are highly
acidic dimeric intracellular proteins that chiefl y bind to phos-
phoserine motifs (Yaffe and Elia, 2001). They play a key reg-
ulatory role in many cellular processes, including signal
transduction, apoptosis, and cell cycle checkpoint control
( Muslin and Xing, 2000; Tzivion et al., 2001; Yaffe, 2002;
Hermeking, 2003). In many instances, 14-3-3 binding leads to
altered subcellular localization of target proteins, which modu-
lates their function. As this appears to be the case for tuberin as
well, we propose a mechanism whereby growth factor stimula-
tion activates AKT, leading to tuberin phosphorylation and
mislocalization within the cell. Although some studies have
suggested that the tuberin–hamartin heterodimer is destabilized
upon AKT-mediated phosphorylation of tuberin (Inoki et al.,
2002; Potter et al., 2002), our data and those of others indicate
that AKT-mediated phosphorylation of tuberin does not change
its affi nity for hamartin (Dan et al., 2002; Manning et al., 2002;
Tee et al., 2003). An alternative hypothesis is that in response to
phosphorylation of S939 and S981 by AKT, 14-3-3 binds to
these phosphorylated motifs to localize tuberin in the cytosol,
physically sequestering tuberin away from hamartin. The pres-
ence of two adjacent 14-3-3 binding sites at S939 and S981 of
tuberin may be important, as they would be predicted to stabi-
lize binding of tuberin in the central channel of the 14-3-3 dimer
more effectively than would a single binding site.
In this regard, it is interesting that loss of either the S939
or S981 14-3-3 binding sites reduces localization of tuberin to
the cytosol. As one of these binding sites is in the alternatively
spliced exon 25 of tuberin, it suggests that alternative splicing
may allow for wider or more varied regulation of this tumor
suppressor. In splice isoforms lacking exon 25, the absence of
the S981 binding site could result in retention at the membrane,
potentially enhancing tuberin activity. This could provide a
mechanism via alternative exon splicing for cells and tissues to
regulate tuberin localization and function. It might be noted that
in previous studies that failed to identify the S939 site within
tuberin as a 14-3-3 binding site, the TSC2 construct used lacked
the alternatively spliced exon 25 and the S981 14-3-3 binding
site (Li et al., 2002).
Hamartin has been shown to signifi cantly enhance
tuberin’s GAP activity toward Rheb GTPase (Garami et al.,
2003; Tee et al., 2003), and our observations also emphasize the
importance of the interaction between tuberin and hamartin as a
functional unit. We found that hamartin directly interacts with
tuberin and promotes tuberin colocalization at the membrane in
proximity to its GAP target Rheb and enhances tuberin’s ability
to repress mTOR signaling. Both endogenously and exoge-
nously expressed hamartin were detected in the membrane frac-
tion of cells, even when cells were treated with serum or growth
factors, data that are consistent with the previous reports that
hamartin is a membrane protein (van Slegtenhorst et al., 1997;
Plank et al., 1998). The colocalization of tuberin in proximity to
its activation partner hamartin and its GAP target Rheb at the
membrane could explain why increased retention of tuberin at
the membrane enhances its ability to inhibit phosphorylation of
S6K. Given that the S939 and S981 residues lie distal from the
COOH-terminal GAP region (residues 1517–1674) of tuberin
and mutation of these sites does not affect the Rheb GAP activ-
ity of tuberin, we conclude that phosphorylation of S939 and
S981 does not directly regulate the GAP function of TSC2.
However, mutation of these sites signifi cantly inhibits Rheb-
induced S6K1 activation through decreased Rheb-GTP load-
ing, suggesting that AKT regulates tuberin function by causing
tuberin translocation to the cytosol, rather than by directly in-
hibiting its intrinsic GAP activity toward Rheb.
Materials and methods
Antibodies and reagents
The following antibodies were used: tuberin, hamartin, lamin A/C, HA,
and Myc (Santa Cruz Biotechnology, Inc.); Rheb, phosphotuberin (T1462),
S6K, phospho-S6K (T389), AKT, phospho-AKT (S473), phospho-AKT
(T308), and phospho-(S/T) AKT substrate (Cell Signaling Technology); Flag
M2 and Flag M2 immobilized agarose beads (Sigma-Aldrich); LDH
(Chemicon International); EGFP (Abcam); and β1-integrin (CLONTECH
Laboratories, Inc.). The following reagents were used: EGF, insulin, and
wortmannin (Sigma-Aldrich) and IGF-1 (R & D Systems). Tuberin peptides
were synthesized by W.M. Keck Biotechnology Resource Center.
Full-length human TSC1 and -2 cDNAs (supplied by J. DeClue, National
Cancer Institute, Bethesda, MD) were subcloned into pcDNA3.1 (Invitro-
gen) and pCMV-Tag2 (Stratagene) expression vectors with NH2-terminal
Myc or Flag epitopes, respectively. COOH-terminal truncation mutant
of TSC2 (∆73) (residues 1–1734) was created by EcoR V digestion fol-
lowed by religation. TSC2 mutations were generated by site-directed
Other constructs used in this study were generously provided as
follows: Flag-TSC1 and HA-S6K from J. Blenis (Harvard Medical School,
Boston, MA), HA-RSK1 from J. Avruch (Massachusetts General Hospital,
Boston, MA), GFP-R18 from T. Pawson (Mount Sinai Hospital, Toronto,
Canada), and the Flag-TSC2 SATA and T1462A from B. Manning
(Harvard School of Public Health, Boston, MA). GST–14-3-3 constructs
were described previously (Liu et al., 2002).
Cell culture, transfection, and immunoprecipitation
Cell lines were grown as follows: MCF7 cells were grown in improved mini-
mum essential medium (Biosource International). TRKE2 cells were grown in
DF8 complete medium (Walker and Ginsler, 1992). HeLa cells were grown
in MEM, and HEK293, Swiss3T3, and NIH3T3 cells were grown in DME
(Life Technologies, Inc.). All media contained 10% FBS (Hyclone) unless
otherwise noted. Myr-Akt MCF7 cells were cultured as previously described
(DeGraffenried et al., 2003). Transfections were performed using the Lipo-
fectamine 2000 reagent (Invitrogen) according to manufacturer’s instructions.
Cells were lysed using PBS containing 0.1% SDS, 1% NP-40, 0.5% deoxy-
cholic acid, 1 μM PMSF, 20 μg/ml aprotinin, 10 μM leupeptin, and 1 μM
Na3VO4. For immunoprecipitation, cell lysates were immunoprecipitated
with the indicated antibodies and protein A– or protein G–Sepharose
beads (GE Healthcare) and washed with buffer (10 mM Tris-HCl, pH 7.5,
1% NP-40, 1% Triton X-100, 100 mM NaCl, 50 mM NaF, 2 mM EDTA,
1 mM PMSF, and Complete protease inhibitor cocktail [Roche]). Immuno-
complexes were subjected to SDS-PAGE and Western blotting.
Subcellular fractionation and protein phosphatase treatments
Cells (70–80% confl uent in 15-cm plates) were washed and collected by
scraping into ice-cold PBS, pelleted by centrifugation at 4°C, resuspended
in hypotonic buffer (10 mM Hepes, pH 7.2, 10 mM KCl, 1.5 mM MgCl2,
0.1 mM EGTA, 20 mM NaF, and 100 μM Na3VO4), and disrupted using
a Dounce homogenizer. Crude nuclei and unbroken cells were then pel-
leted by centrifugation at 3,000 rpm at 4°C for 5 min. The postnuclear
supernatant was separated by ultracentrifugation at 100,000 g for 1 h at
4°C. The supernatant, or cytosolic fraction, was removed and the pellet
JCB • VOLUME 173 • NUMBER 2 • 2006 288
was lysed in 1× lysis buffer (20 mM Tris-HCl, pH 7.5, 150 mM NaCl,
1 mM EDTA, 1 mM EGTA, 1% Triton X-100, 2.5 mM NA4P207, 1 mM
β-glycerophosphate, 1 mM Na3VO4, and 1 μg/ml leupetin). The insoluble
fractions were removed by centrifugation at 14,000 rpm for 10 min, and
the supernatant was collected as the membrane fraction. Crude nuclei
were resuspended with hypotonic buffer and homogenized using a Dounce
homogenizer. After centrifugation at 4°C for 5 min, the pellet was further
washed with wash buffer (10 mM Tris-HCl, pH 7.4, 0.1% NP-40, 0.05%
sodium deoxycholate, 10 mM NaCl, and 3 mM MgCl2) and lysed in high-
salt lysis buffer (20 mM Hepes, pH 7.4, 0.5 M NaCl, 0.5% NP-40, and
1.5 mM MgCl2). All lysis and wash buffers contained 1× Complete prote-
ase inhibitor cocktail. The nuclear, membrane, and cytosolic lysates were
normalized using the BCA Protein Assay kit (Pierce Chemical Co.) and
subjected to SDS-PAGE and immunoblot analysis. For phosphatase treat-
ment, equal amounts of membrane and cytosolic lysates were treated with
CIAP serine/threonine phosphatase or YOP tyrosine phosphatase at 30°C
for 1 h.
GST–14-3-3 pull-down assays
GST pull-down assays were performed as previously described (Liu et al.,
2002). For peptide competition assays, incubations were performed in the
presence or absence of 100 μM phospho-Ser939 or Ser939 tuberin peptide
(S G S G F R A R S T S  LNERPK) or phospho-Ser981 or Ser981 tuberin peptide
(S G S G F R C R S I S  VSEHVV).
Rheb GAP and S6K1 assay
Anti-Flag M2 antibody was used to immunoprecipitate Flag-tuberin from
HEK293E cells. Immunocomplexes of TSC1/TSC2 were used for in vitro
RhebGAP assays as previously described (Tee et al., 2003). α-[32P]GTP
and α-[32P]GDP were eluted from Rheb and resolved by thin layer chroma-
tography on PEI cellulose (Sigma-Aldrich) with KH2PO4. The relative levels
of radiolabeled GTP and GDP were quantifi ed with a phosphorimager.
S6K1 activity assays on immunoprecipitated HA-S6K1 were performed
as previously described (Martin et al., 2001) using recombinant GST-S6
(32 COOH-terminal amino acids of ribosomal protein S6) as substrate.
In vivo Rheb nucleotide binding
Analysis of Rheb guanine nucleotide binding in cells was determined as
previously described (Tee et al., 2005).
Immunofl uorescent staining
Intracellular localization of wild-type tuberin and 2A mutant and colocal-
ization studies was determined by immunofl uorescence analysis of HeLa
cells. Transfected cells were plated onto glass chamber slides in the ab-
sence of serum. After 12 h of serum starvation, transfected cells were
treated with either 20% FBS or 30 ng/ml IGF-1 for 1 h. Cells were then
fi xed in 50% ethanol plus 10% acetic acid for 1 h at 4°C, and nonspecifi c
antigens were blocked for 1 h in PBS containing 7.5% BSA at 37°C.
Rabbit anti-Flag (Sigma-Aldrich) and/or mouse anti-Myc (Santa Cruz
Biotechnology, Inc.) primary antibodies were incubated in PBS/7.5% BSA
at 37°C for 2 h. Primary antibodies were detected with FITC-conjugated
goat anti–rabbit (Abcam) and Cy3-conjugated donkey anti–mouse ( Jackson
ImmunoResearch Laboratories) antibodies, respectively.
Fluorescence images were analyzed either on a confocal microscope
(Fluoview Scanning Laser Biological Microscope IX 70 system; Olympus)
equipped with two lasers (Ar 488 and Kr-Ar 488–564) using UplanFl
100× oil-immersion (NA 1.30) objective or a conventional microscope
with fl uorescence attachment (BX40 attached with BX-FLA; Olympus) using
UplanFl 40× (NA 0.75) objective. Fluoview version 1.26 software
( Olympus) and MagnaFire version 2.1C (Olympus) were used for image
acquisition from confocal microscopy and conventional microscopy,
respectively. Photoshop 8.0 software (Adobe) was used for minor adjust-
ments and processing of images.
Online supplemental material
Fig. S1 shows that AKT-mediated phosphorylation of tuberin leads to sub-
cellular translocation. Fig. S2 shows that S939 and S981 are required for
altered subcellular localization of tuberin. Fig. S3 demonstrates that 14-3-3
proteins bind phosphorylated S939 and S981 residues. Fig. S4 demon-
strates that tuberin mutants retain their ability to interact with hamartin. Fig.
S5 shows that Rheb is retained in the membrane fraction in the presence or
absence of growth factor treatment. Online supplemental material is avail-
able at http://www.jcb.org/cgi/content/full/jcb.200507119/DC1.
We are grateful to Drs. Jeffrey DeClue, John Blenis, Joseph Avruch, Tony
Pawson, and Brendan Manning for providing DNA constructs and to Dr. Linda
deGraffenried for the Myr-Akt MCF7 cell line. We are especially indebted to
Dr. Brendan Manning for intellectual contributions to the project and critical
reading of the manuscript. We also thank Hannah Gattis and Jennifer Widener
for technical assistance.
This work was supported in part by National Institutes of Health grants
CA63613 and HD46282 (to C.L. Walker), ES08263-06S1 (to J.M. Bergeron),
and ES07784 and CA16672 (to the University of Texas MD Anderson
Cancer Center). A British Hearth Foundation intermediate research fellowship
(FS/04/002) was awarded to A.R. Tee.
The authors declare that they have no competing fi nancial interests.
Submitted: 27 July 2005
Accepted: 21 March 2006
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