Published in Volume
119, Issue 5 (May 1, 2009)
J Clin Invest. 2009;119(5):1359–1372.
doi:10.1172/JCI37948.
Copyright © 2009, American Society for Clinical
Investigation
Research Article
Cannabinoid action induces autophagy-mediated cell death through
stimulation of ER stress in human glioma cells
María Salazar1,2, Arkaitz Carracedo1, Íñigo J. Salanueva1, Sonia Hernández-Tiedra1, Mar Lorente1,2, Ainara Egia1, Patricia Vázquez3, Cristina Blázquez1,2, Sofía Torres1, Stephane García4, Jonathan Nowak4, Gian María Fimia5, Mauro Piacentini5, Francesco Cecconi6, Pier Paolo Pandolfi7, Luis González-Feria8, Juan L. Iovanna4, Manuel Guzmán1,2, Patricia Boya3 and Guillermo Velasco1,2
1Department of Biochemistry and Molecular Biology I, School of Biology,
Complutense University, Madrid, Spain.
2Centro de
Investigación Biomédica en Red sobre Enfermedades
Neurodegenerativas (CIBERNED), Madrid, Spain.
33D Lab (Development,
Differentiation, and Degeneration), Department of Cellular and Molecular
Physiopathology, Centro de Investigaciones Biológicas, Consejo Superior de
Investigaciones Científicas (CSIC), Madrid, Spain.
4INSERM
U624, Campus de Luminy, Marseille, France.
5National Institute for
Infectious Diseases, IRCCS “L. Spallanzani,” Rome, Italy.
6Laboratory of Molecular Neuroembryology, IRCCS Fondazione Santa Lucia
and Department of Biology, University of Rome “Tor Vergata,”
Rome, Italy.
7Cancer Genetics Program, Beth Israel Deaconess Cancer
Center and Department of Medicine, Beth Israel Deaconess Medical Center, Harvard Medical
School, Boston, Massachusetts, USA.
8Department of Neurosurgery,
University Hospital, Tenerife, Spain.
Address correspondence to: Guillermo Velasco, Department of Biochemistry and
Molecular Biology I, School of Biology, Complutense University, c/ José
Antonio Novais s/n, 28040 Madrid, Spain. Phone: 34-913944668; Fax: 34-913944672;
E-mail: gvd@bbm1.ucm.es.
First published April 1, 2009
Received for publication November 3,
2008, and accepted in revised form February 11,
2009.
Autophagy can promote cell survival or cell death, but the molecular basis underlying
its dual role in cancer remains obscure. Here we demonstrate that
Δ9-tetrahydrocannabinol (THC), the main active component of
marijuana, induces human glioma cell death through stimulation of autophagy. Our data
indicate that THC induced ceramide accumulation and eukaryotic translation initiation
factor 2α (eIF2α) phosphorylation and thereby activated an ER
stress response that promoted autophagy via tribbles homolog 3–dependent
(TRB3-dependent) inhibition of the Akt/mammalian target of rapamycin complex 1
(mTORC1) axis. We also showed that autophagy is upstream of apoptosis in
cannabinoid-induced human and mouse cancer cell death and that activation of this
pathway was necessary for the antitumor action of cannabinoids in vivo. These
findings describe a mechanism by which THC can promote the autophagic death of human
and mouse cancer cells and provide evidence that cannabinoid administration may be an
effective therapeutic strategy for targeting human cancers.
Introduction
Macro-autophagy, hereafter referred to as “autophagy,” is a
highly conserved cellular process in which cytoplasmic materials — including
organelles — are sequestered into double-membrane vesicles called
autophagosomes and delivered to lysosomes for degradation or recycling (1). In many cellular settings, triggering of
autophagy relies on the inhibition of mammalian target of rapamycin complex 1 (mTORC1),
an event that promotes the activation (de-inhibition) of several autophagy proteins
(Atgs) involved in the initial phase of membrane isolation (1). Enlargement of this complex to form the autophagosome requires
the participation of 2 ubiquitin-like conjugation systems. One involves the conjugation
of ATG12 to ATG5 and the other of phosphatidylethanolamine to LC3/ATG8 (1). The final outcome of the activation of the
autophagy program is highly dependent on the cellular context and the strength and
duration of the stress-inducing signals (2–5). Thus, besides its
role in cellular homeostasis, autophagy can be a form of programmed cell death,
designated “type II programmed cell death,” or play a
cytoprotective role, for example in situations of nutrient starvation (6). Accordingly, autophagy has been proposed to play
an important role in both tumor progression and promotion of cancer cell death (2–4), although the molecular mechanisms responsible for this dual action of
autophagy in cancer have not been elucidated.
Δ9-Tetrahydrocannabinol (THC), the main active component of
marijuana (7), exerts a wide variety of biological
effects by mimicking endogenous substances — the endocannabinoids
— that bind to and activate specific cannabinoid receptors (8). One of the most exciting areas of research in the
cannabinoid field is the study of the potential application of cannabinoids as
antitumoral agents (9). Cannabinoid administration
has been found to curb the growth of several types of tumor xenografts in rats and mice
(9, 10).
Based on this preclinical evidence, a pilot clinical trial has been recently run to
investigate the antitumoral action of THC on recurrent gliomas (11). Recent findings have also shown that the pro-apoptotic and
tumor growth–inhibiting activity of cannabinoids relies on the upregulation
of the transcriptional co-activator p8 (12) and
its target the pseudo-kinase tribbles homolog 3 (TRB3) (13). However, the mechanisms that promote the activation of this signaling
route as well as the targets downstream of TRB3 that mediate its tumor
cell–killing action remain elusive. In this study we found that ER
stress–evoked upregulation of the p8/TRB3 pathway induced autophagy via
inhibition of the Akt/mTORC1 axis and that activation of autophagy promoted the
apoptotic death of tumor cells. The uncovering of this pathway, which we believe is
novel, for promoting tumor cell death may have therapeutic implications in the treatment
of cancer.
Results
Autophagy mediates THC-induced cancer cell death. As a first approach to gain insight into the morphological changes induced in cancer
cells by cannabinoid administration, we performed electron microscopy analysis of
U87MG human astrocytoma cells. Interestingly, double membrane vacuolar structures
with the morphological features of autophagosomes were observed in THC-treated cells
(Figure 1, A–C). The conversion of
the soluble form of LC3 (LC3-I) to the lipidated and autophagosome-associated form
(LC3-II) is considered one of the hallmarks of autophagy (1), and thus we observed the occurrence of LC3-positive dots as
well as the appearance of LC3-II (Figure 1D) in
cannabinoid-challenged cells. In addition, co-incubation with the lysosomal protease
inhibitors E64d and pepstatin A, which blocks the last steps of autophagic
degradation (14), enhanced THC-induced
accumulation of LC3-II (Figure 1E), confirming
that cannabinoids induce dynamic autophagy in U87MG cells. Furthermore, incubation
with the cannabinoid receptor 1 (CB1) antagonist SR141716 prevented THC-induced LC3
lipidation and formation of LC3 dots (Figure 1D), indicating that induction of autophagy by cannabinoids relies on CB1
receptor activation.
Since autophagy has been implicated in promotion and inhibition of cell survival, we
next investigated its participation in the cancer cell death–inducing
action of THC. Pharmacological inhibition of autophagy at different levels
(Supplemental Figure 1, A–C; supplemental material available online with
this article; doi:10.1172/JCI37948DS1) or selective knockdown of ATG1 (an essential
protein in the initiation of autophagy; ref. 1)
(Figure 1, F and G), ATG5 (an essential protein
in the formation of the autophagosome; ref. 1)
(Supplemental Figure 1, D–F), or AMBRA1 (a recently identified
beclin-1–interacting protein that regulates autophagy; ref. 15) (Supplemental Figure 1, D–F)
strongly reduced cannabinoid-induced autophagy and cell death. Moreover, transformed
Atg5-deficient mouse embryonic fibroblasts (MEFs), which are
defective in autophagy (16), were more
resistant than their wild-type counterparts to THC-induced cell death (Figure 1H) and did not undergo autophagy upon cannabinoid
treatment (Figure 1I). Taken together, these
findings demonstrate that autophagy plays a prominent role in THC-induced cancer cell
death.
THC induces autophagy via ER stress–dependent upregulation of p8 and
TRB3. In addition to the presence of autophagosomes, electron microscopy analysis of
cannabinoid-treated cells revealed the presence of numerous cells with dilated ER
(Figure 2A). In line with this observation,
immunostaining of the ER luminal marker protein disulphide isomerase (PDI) showed a
striking dilation in the ER of THC-treated U87MG cells (Figure 2B), an event that was associated with an increased
phosphorylation of the α subunit of eukaryotic translation initiation
factor 2 (eIF2α), a hallmark of the ER stress response (17) (Figure 2C). In addition, THC-induced ER dilation and eIF2α
phosphorylation were prevented by pharmacological blockade of the CB1 receptor
(Figure 2, B and C).
Time-course analysis of PDI and LC3 immunostaining, eIF2α
phosphorylation, and LC3 lipidation of cannabinoid-treated cells revealed that ER
stress occurred earlier than autophagy (Figure 2, D and E). Of interest, cannabinoid administration produced similar
activation of ER stress and autophagy, as well as cell death, in other human
astrocytoma cell lines (Supplemental Figure 2, A–F), a primary culture of
human glioma cells (Supplemental Figure 2, G–I), and several human cancer
cell lines of different origin, including pancreatic cancer (Supplemental Figure 2,
J–L), breast cancer, and hepatoma (data not shown). However, neither ER
dilation nor eIF2α phosphorylation or autophagy was evident in normal,
nontransformed primary astrocytes (Supplemental Figure 3), which are resistant to
cannabinoid-induced cell death (13).
We next investigated whether activation of ER stress is involved in the induction of
autophagy in response to cannabinoid treatment of cancer cells. We have previously
shown that THC-induced accumulation of de novo–synthesized ceramide, an
event that occurs in the ER (18), leads to
upregulation of the stress-regulated protein p8 and its ER stress–related
downstream targets, ATF4, CHOP, and TRB3, to induce cancer cell death (13). Of importance, incubation with ISP-1 (a
selective inhibitor of serine palmitoyltransferase, the enzyme that catalyzes the
first step of sphingolipid biosynthesis; ref. 18) prevented ceramide accumulation (Supplemental Figure 4A); THC-induced ER
dilation (Supplemental Figure 4B); eIF2α phosphorylation (Figure 3A); p8, ATF4, CHOP, and TRB3 upregulation
(Supplemental Figure 4C); and autophagy (Figure 3B), supporting that ceramide accumulation is involved in
cannabinoid-triggered ER stress and autophagy. We also verified by means of RNA
interference that CaCMKKβ — which had been previously
implicated in activating autophagy in response to ER stress–associated
calcium release (19) — was not
involved in THC-induced autophagy and cell death (data not shown). As phosphorylation
of eIF2α on Ser51 attenuates general protein synthesis while enhancing
the expression of several ER stress response genes (17), we used cells derived from eIF2α S51A knockin mice to
test whether eIF2α phosphorylation regulates the expression of p8 and its
downstream targets. In agreement with this hypothesis, THC treatment (which promoted
ceramide accumulation in both wild-type and eIF2α S51A immortalized MEFs;
Supplemental Figure 5A) triggered p8, ATF4, CHOP, and TRB3 upregulation (Figure 3C) as well as autophagy (Supplemental Figure 5B)
in wild-type cells but not in their eIF2α S51A counterparts.
We subsequently asked whether p8 and its downstream targets regulate autophagy.
Knockdown of p8 or TRB3 prevented THC-induced autophagy (Figure 3, D and E) but not ER dilation (Supplemental Figure 4D) in U87MG
cells. Furthermore, THC induced autophagy in p8+/+ but
not p8-deficient transformed MEFs (Figure 3F and Supplemental Figure 5C). Altogether, these findings reveal
that THC induces autophagy of cancer cells via activation of an ER
stress–triggered signaling route that involves stimulation of ceramide
synthesis de novo, eIF2α phosphorylation, and p8 and TRB3 upregulation.
THC inhibits Akt and mTORC1 via TRB3. Inhibition of mTORC1 is considered a key step in the early triggering of autophagy
(6). We therefore tested whether
cannabinoid-induced upregulation of the p8 pathway leads to autophagy via inhibition
of this complex. THC treatment of U87MG cells reduced the phosphorylation of p70S6
kinase (a well-established mTORC1 substrate) and the ribosomal protein S6 (a
well-established p70S6 kinase substrate) (Figure 4, A and C), indicating that mTORC1 is inhibited in cannabinoid-challenged
cells. In addition, the cannabinoid-induced decrease in p70S6 kinase and S6
phosphorylation, autophagy, and cell death were not evident in
Tsc2–/– cells, in which
mTORC1 is constitutively active (20) (Figure
4B and Supplemental Figure 6, A and B),
further supporting a major role for mTORC1 inhibition in the induction of autophagic
cell death by cannabinoids.
The protein kinase Akt positively regulates the activity of the mTORC1 complex by
phosphorylating and inhibiting TSC2 and PRAS40 (a well-established Akt substrate
within the mTORC1 complex). Thus, Akt inhibition decreases mTORC1 activity and
promotes autophagy (20). In line with this
idea, THC decreased the phosphorylation of Akt, TSC2, and PRAS40 as well as p70S6
kinase and S6 (Figure 4C). This inhibition of
the Akt/mTORC1 pathway was abrogated by incubation with a CB1 receptor antagonist
(Supplemental Figure 6C) or a ceramide synthesis inhibitor (Supplemental Figure 6D).
Likewise, cells overexpressing a myristoylated (constitutively active) form of Akt
were resistant to THC-induced mTORC1 inhibition, autophagy, and cell death (Figure
4D and Supplemental Figure 6, E and F),
further supporting that THC induces autophagy via Akt inhibition.
Since TRB3 has been shown to directly interact with and inhibit Akt (21, 22),
we investigated whether upregulation of TRB3 was responsible for THC-induced
Akt/mTORC1 inhibition. Several observations support that this is indeed the case: (a)
THC increased the amount of Akt coimmunoprecipitated with TRB3 from U87MG extracts
(Figure 4E), (b) knockdown of TRB3 prevented the
effect of THC on Akt, TSC2, PRAS-40, p70S6 kinase, and S6 phosphorylation (Figure
4F), and (c) TRB3 overexpression decreased
Akt, TSC2, PRAS40, p70S6 kinase, and S6 phosphorylation, enhanced the inhibitory
effect of THC on the phosphorylation of these proteins, and promoted autophagy
(Figure 4G). In line with these observations,
THC failed to inhibit Akt, p70S6 kinase, and S6 phosphorylation of eIF2α
S51A knockin or p8-deficient MEFs, in which TRB3 did not become upregulated upon
cannabinoid treatment (Figure 4H and
Supplemental Figure 6, G and H). Altogether, these data demonstrate that upregulation
of p8 and TRB3 induce autophagy of tumor cells via inhibition of the Akt/mTORC1
pathway.
THC-induced autophagy promotes the apoptotic death of cancer cells. While analyzing the mechanism of cannabinoid cell-killing action, we observed that
incubation with the pan-caspase inhibitor ZVAD-fmk prevented cell death to the same
extent as genetic (Figure 5A) or pharmacological
(Supplemental Figure 7) inhibition of autophagy. Furthermore,
Bax/Bak double knockout (DKO) immortalized MEFs, which are protected
against mitochondrial apoptosis (23), were
resistant to THC-induced cell death and apoptosis (Figure 5B) but underwent eIF2α phosphorylation and autophagy
(Figure 5C) upon THC treatment. We therefore
investigated whether cannabinoid-induced autophagy promoted the apoptotic death of
cancer cells. Time-course analysis of LC3 and active caspase-3 immunostaining in
U87MG cells revealed that autophagy preceded the appearance of apoptotic features in
THC-treated cells (Figure 5D). In addition,
selective knockdown of ATG1 (Figure 5D) as well
as of AMBRA1 or ATG5 (Supplemental Figure 8) prevented THC-induced caspase-3
activation. Moreover, unlike their wild-type counterparts,
Atg5-deficient immortalized MEFs did not undergo phosphatidylserine
translocation to the outer leaflet of the plasma membrane (Figure 5E), loss of mitochondrial membrane potential
(Figure 5F), or increased production of reactive
oxygen species (Supplemental Figure 9) in response to cannabinoid treatment. These
findings indicate that activation of the autophagy-mediated cell death pathway occurs
upstream of apoptosis in cannabinoid antitumoral action.
Activation of autophagy is necessary for cannabinoid antitumoral action in vivo. To determine the in vivo relevance of our findings, we first investigated whether THC
promotes the activation of the above-described autophagy-mediated cell death pathway
in U87MG cell–derived tumor xenografts, in which we have recently shown
that cannabinoid treatment reduces tumor growth (specifically, THC administration for
14 days decreased tumor growth by 50%; ref. 13). Analysis of these tumors revealed that cannabinoid administration
increases TRB3 expression and decreases S6 phosphorylation (Figure 6A). Likewise, formation of LC3 dots as well as
increase in LC3-II and active caspase-3 immunostaining were observed in THC-treated,
but not vehicle-treated, tumors (Figure 6B).
To further investigate whether activation of the p8 pathway mediates cannabinoid
antitumoral action, we also analyzed tumors derived from
p8+/+ and
p8–/–
RasV12/E1A-transformed MEFs (in this case, THC administration for 8 days
decreased by 45% the growth of p8+/+ tumors but had no
significant effect on p8–/–
tumors; ref. 13). THC treatment increased TRB3
expression, decreased S6 phosphorylation, and increased autophagy as well as TUNEL
and active caspase-3 immunostaining in p8+/+ but not
p8–/– tumors (Figure 6C and Supplemental Figure 10). Moreover, THC treatment enhanced
the number of cells with LC3 dots and TUNEL-positive nuclei in p8+/+ but
not in p8–/– tumors (Figure 6C).
In order to verify the importance of autophagy for cannabinoid antitumoral action, we
next generated tumors with Atg5+/+ and
Atg5–/–
RasV12/T-large antigen transformed MEFs. THC administration reduced by
more than 80% the growth of tumors derived from wild-type cells but had no
significant effect on those tumors generated by autophagy-deficient cells (Figure
7A). Furthermore, cannabinoid administration
increased autophagy, TUNEL (Figure 7B), and
active caspase-3 immunostaining (Supplemental Figure 11) in
Atg5+/+ but not
Atg5–/– tumors. Likewise,
cannabinoid administration increased the number of cells with LC3 dots and
TUNEL-positive nuclei in Atg5+/+ but not
Atg5–/– tumors (Figure
7B). Taken together, these findings
demonstrate that activation of the autophagy-mediated cell death pathway is
indispensable for cannabinoid antitumoral action.
Finally, we analyzed the tumors of 2 patients enrolled in a clinical trial aimed at
investigating the effect of THC on recurrent glioblastoma multiforme. The patients
were subjected to intracranial THC administration, and biopsies were taken before and
after the treatment (11). In the 2 patients,
cannabinoid inoculation increased TRB3 immunostaining and decreased S6
phosphorylation (Figure 8A). Interestingly, the
number of cells with autophagic phenotype (Figure 8B) as well as with active caspase-3 immunostaining (Figure 8C) was increased in the tumor samples obtained
after THC treatment. Although these studies were only conducted in specimens from 2
patients, they are in line with the preclinical evidence shown above and suggest that
cannabinoid administration might also trigger autophagy-mediated cell death in human
tumors.
Discussion
In this study we show that cannabinoids, a new family of potential antitumoral agents,
induce autophagy of cancer cells and that this process mediates the cell
death–promoting activity of these compounds. Several observations strongly
support this idea: (a) THC induced autophagy and cell death in different types of cancer
cells but not in nontransformed astrocytes, which are resistant to cannabinoid killing
action, (b) pharmacological or genetic inhibition of autophagy prevented THC-induced
cell death, (c) autophagy-deficient tumors were resistant to THC growth-inhibiting
action, and (d) THC administration activated the autophagic cell death pathway in 3
different models of tumor xenografts as well as in 2 human tumor samples.
Depending on the cellular context and the strength and duration of the triggering
stimulus, autophagy is involved in the promotion or inhibition of cancer cell survival
(4, 5,
24, 25). However, the molecular bases of this dual role of autophagy in cancer
remain unknown. Data presented here demonstrate that induction of autophagy by
cannabinoids leads to cancer cell death and identify the signaling route responsible for
the activation of this cellular process. Thus, our findings suggest that THC
— via activation of the CB1 receptor and stimulation of ceramide synthesis
de novo — activates an early ER stress response that leads to increased
phosphorylation of eIF2α on Ser51. Experiments performed with
eIF2α S51A mutant cells have shown that phosphorylation of this residue,
which is known to attenuate general protein translation while enhancing the expression
of several genes related with the ER stress response (17), is required for the upregulation of the stress protein p8 and its ER
stress–related downstream targets ATF4, CHOP, and TRB3 as well as for the
induction of autophagy by cannabinoids. Furthermore, we demonstrate that the
upregulation of p8 and TRB3, which has been previously implicated in cannabinoid-evoked
cell death (13), is a crucial event in the
triggering of autophagy. Ceramide accumulation has been proposed to induce ER stress
(26, 27) and autophagy (28), and
eIF2α phosphorylation has been implicated in the induction of autophagy in
response to different situations (29–31). However, the
molecular mechanisms responsible for these actions have not been clarified. Findings
presented here now suggest that upregulation of the p8-TRB3 pathway constitutes a
mechanism by which de novo–synthesized ceramide and eIF2α
phosphorylation promote autophagy, thus identifying what we believe is a novel
connection between ER stress and autophagy.
Our data also demonstrate that the autophagy-promoting activity of the p8-regulated
pathway is based on its ability to inhibit the Akt/mTORC1 axis. Regulation of mTORC1
largely relies on the activity of the prosurvival kinase Akt, whose inhibition leads to
mTORC1 inactivation and, in turn, to autophagy (20). Our findings reveal that THC upregulates TRB3, promoting its interaction
with Akt and leading to decreased phosphorylation of this kinase as well as of its
direct substrates TSC2 and PRAS40, which triggers mTORC1 inhibition and induction of
autophagy. TRB3 has been previously shown to inhibit Akt (21, 22), although the precise
contribution of this pseudo-kinase to the regulation of Akt activity in different
cellular contexts is unclear (32). Here we
demonstrate that TRB3 inhibition of the Akt/mTORC1 axis is essential for
cannabinoid-induced autophagy of cancer cells. Moreover, we show that this pathway is
essential for cannabinoid antitumoral action. Thus, THC administration leads to TRB3
upregulation, mTORC1 inhibition, induction of autophagy, and reduction of tumor growth
in different models of tumor xenografts, but not in p8-deficient tumors that are
defective in the upregulation of the p8/TRB3 pathway. Furthermore, activation of this
pathway was also evident in 2 glioma patients that had been treated with THC. These
results thus uncover a role for TRB3 that may be of great importance in the regulation
of cancer cell death.
Autophagy has been proposed to protect from apoptosis, act as an apoptosis-alternative
pathway to induce cell death, or act together with apoptosis as a combined mechanism for
cell death (6, 33). However, very little is known about the role of the interplay between these
2 cellular processes in the control of tumor growth in response to anticancer agents.
Our results now clearly demonstrate that induction of autophagy is involved in the
mechanism by which cannabinoids promote the activation of the mitochondrial
pro-apoptotic pathway. Thus, neither tumors in which the p8-regulated pathway has been
ablated (and in which, therefore, THC treatment does not induce autophagy) nor tumors
intrinsically deficient in autophagy undergo apoptosis in response to THC, and so they
are resistant to THC antitumoral action. These findings reveal that autophagy is
required for the activation of apoptosis in response to cannabinoid treatment in vivo.
It is worth noting that the concentrations of THC used in this study are in the same
range as those administered intracranially to the patients in which we observed
activation of the autophagy-mediated cell death pathway (11) and could be thus considered clinically relevant. Of interest,
intraperitoneal administration of THC to U87MG tumor xenografts produces a similar
decrease in tumor growth (that occurs in concert with increased autophagy and apoptosis)
to that observed when the cannabinoid is administered peritumorally (our unpublished
observations). Considering that no signs of toxicity were observed in the clinical trial
patients (11) or in tumor-bearing animals treated
intracranially, peritumorally, or intraperitoneally with THC (refs. 34 and 35 and
data not shown), and that no overt toxic effects have been reported in other clinical
trials of cannabinoid use in cancer patients for various applications (e.g., inhibition
of nausea, vomiting, and pain) and using different routes of administration (e.g., oral,
oro-mucosal) (9, 36), our findings support that safe, therapeutically efficacious doses of THC
may be reached in cancer patients.
In summary, in this study we identify what we believe is a new route that links the ER
stress response to the activation of autophagy and promotes the apoptotic death of tumor
cells (Figure 7C). The identification of this
pathway will help to understand the molecular events that lead to activation of
autophagy-mediated cell death by anticancer drugs and may contribute to the design of
new therapeutic strategies for inhibiting tumor growth.
Methods
Cell culture and viability. Cortical astrocytes were prepared from 24-hour-old mice as previously described
(13). Primary cultures of brain tumor cells
were prepared and cultured as described in the Supplemental Methods. U87MG, T98G,
U373MG, and MiaPaCa2 cells, p8+/+ and
p8–/– RasV12/E1A
MEFs, Atg5+/+ and
Atg5–/– T-large antigen
MEFs (provided by Noboru Mizushima, Tokyo Medical and Dental University, Tokyo,
Japan), Bax/Bak wild-type and Bax/Bak DKO T-large
antigen MEFs (provided by Luca Scorrano, Dulbecco Telethon Institute, Milan, Italy,
and Patrizia Agostinis, Catholic University of Leuven, Leuven, Belgium),
eIF2α S51S WT and eIF2α S51A T-large antigen MEFs (provided
by Richard Kaufman, University of Michigan, Ann Arbor, Michigan, USA, and Cesar de
Haro and Juan J. Berlanga, Centro de Biología Molecular Severo Ochoa,
Autonoma University, Madrid, Spain), Tsc2+/+ and
Tsc2–/–
p53–/– MEFs, empty vector (pBABE)
and pBABE-myr-Akt MEFs, and Atg5+/+ and
Atg5–/–
RasV12/T-large antigen MEFs were cultured in DMEM containing 10% FBS and
transferred to medium containing 0.5% FBS (except RasV12/E1A-transformed
MEFs, which were transferred to medium containing 2% FBS) 18 h before performing the
different treatments. p8+/+ and
p8–/– RasV12/E1A
MEFs as well as Atg5+/+ and
Atg5–/–
RasV12/T-large antigen MEFs correspond to a polyclonal mix of at least 20
different selected clones. Unless otherwise indicated, THC was used at a final
concentration of 5 μM. Cell viability was determined by the MTT
[3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide] test (Sigma-Aldrich).
Flow cytometry. Briefly, cells (approximately 5 × 105 cells per assay) were
trypsinized, divided in 2 tubes, washed, and collected by centrifugation at 1,500
g for 5 min. One aliquot was incubated for 10 min at
37°C with Annexin V–FITC (BD Biosciences). Propidium iodide
(1 μg/ml) was added just before cytofluorometric analysis. The other
aliquot was simultaneously labeled with 3,3ι-dihexyloxacarbocyanine
iodide (DiOC6[3], 40 nM; Invitrogen) and hydroethidium (5 μM;
Invitrogen) for 10 minutes at 37°C, followed by cytofluorometric
analysis. Cells (10,000) were recorded in each analysis. Fluorescence intensity was
analyzed in an EPICS XL flow cytometer (Beckman Coulter).
Western blot. Western blot analysis was performed following standard procedures. A list of the
antibodies used can be found in Supplemental Methods. Densitometric analysis was
performed with Quantity One software (Bio-Rad).
Transfections. U87MG cells (75% confluent) were transfected with siRNA duplexes using the DharmaFECT
1 Transfection reagent (Dharmacon). Cells were trypsinized and seeded 24 h after
transfection, at a density of 5,000 cells/cm2. Transfection efficiency was
greater than 70% as monitored with a control fluorescent (red) siRNA (siGLO RISC-Free
siRNA; Dharmacon). In immunofluorescence experiments, control and selective siRNAs
were used in a 1:5 ratio, and cells with red spots were scored as transfected.
Infections with adenoviral vectors. U87MG cells (75% confluent) were transduced for 1 h with supernatants obtained from
HEK293 cells infected with adenoviral vectors carrying EGFP (provided by Javier G.
Castro, Hospital Infantil Universitario Niño Jesús, Madrid,
Spain), rat HA-tagged TRB3 (donated by Patrick Iynedjian, University of Geneva,
Geneva, Switzerland) (32), or human EGFP-LC3
(provided by Aviva Tolkovsky and Christoph Goemans, University of Cambridge,
Cambridge, United Kingdom). Infection efficiency was greater than 80% as determined
by EGFP fluorescence.
RNA interference. Double-stranded RNA duplexes were purchased from Dharmacon. A list of sequences can
be found in the Supplemental Methods.
RT-PCR analysis. RNA was isolated using Trizol Reagent (Invitrogen). cDNA was obtained with
Transcriptor Reverse transcriptase (Roche Applied Science). Primers and amplification
conditions can be found in the Supplemental Methods.
Real-time quantitative PCR. cDNA was obtained using Transcriptor (Roche Applied Science). Real-time quantitative
PCR assays were performed using the FastStart Universal Probe Master mix with Rox
(Roche Applied Science), and probes were obtained from the Universal ProbeLibrary Set
(Roche Applied Science). Primer sequences can be found in the Supplemental Methods.
Amplifications were run in a 7900 HT-Fast Real-Time PCR System (Applied Biosystems).
Each value was adjusted by using 18S RNA levels as a reference.
Immunoprecipitation. U87MG cells were lysed in HEPES lysis buffer (see Supplemental Methods for buffer
composition). Lysate (1–4 mg) was precleared by incubating with
5–20 μl of protein G–Sepharose conjugated to
pre-immune IgG. The lysate extracts were then incubated with 5–20
μl of protein G–Sepharose conjugated to 5–20
μg of the anti-TRB3 antibody or pre-immune IgG. TRB3 antibody
(aminoterminal end, ab50516; Abcam) was covalently conjugated to protein
G–Sepharose using dimethyl pimelimidate. Immunoprecipitations were
carried out for 1 h at 4°C on a rotatory wheel. The immunoprecipitates
were washed 4 times with HEPES lysis buffer, followed by 2 washes with HEPES kinase
buffer. The immunoprecipitates were resuspended in 30 μl of sample buffer
(not containing 2-mercaptoethanol) and filtered through a 0.22-μm Spin-X
filter, and 2-mercaptoethanol was added to a concentration of 1% (vol/vol). Samples
were subjected to electrophoresis and immunoblot analysis.
Ceramide levels. Ceramide levels were determined as previously described (37).
Confocal laser scanning microscopy. Standard protocols for immunofluorescence microscopy were used (see
Supplemental Methods for the antibodies used). To quantify the percentage of cells
with LC3 or PDI dots, at least 200 cells per condition were counted in randomly
selected fields. In all cases, only those cells with 4 or more prominent dots of
either LC3 or PDI were scored positively.
In vivo treatments. Tumors derived from U87MG cells and p8+/+ and
p8–/– MEFs were induced and treated as
previously described (13). Tumors derived from
Atg5+/+ or
Atg5–/–
RasV12/T-large antigen MEFs (see Supplemental Methods for the procedure
used to generate these cells) were induced in nude mice by subcutaneous injection of
107 cells in PBS supplemented with 0.1% glucose. Tumors were allowed to
grow until an average volume of 200–250 mm3, and animals were
assigned randomly to the different groups. At this point, vehicle or THC (15 mg/kg/d)
in 100 μl of PBS supplemented with 5 mg/ml BSA was administered daily in
a single peritumoral injection. Tumors were measured with an external caliper, and
volume was calculated as (4π/3) × (width/2)2
× (length/2). All procedures involving animals were performed with the
approval of the Complutense University Animal Experimentation Committee according to
Spanish official regulations.
Human tumor samples. Tumor biopsies were obtained from 2 recurrent glioblastoma multiforme patients who
had been treated with THC. The characteristics of the patients and the clinical study
have been described in detail elsewhere (11).
Briefly, THC dissolved in 30 ml of physiological saline solution plus 0.5% (wt/vol)
human serum albumin was administered intratumorally to the patients. Patient 1
received a total of 1.46 mg of THC for 30 days, while patient 2 received a total of
1.29 mg of THC for 26 days (it was estimated that doses of 6–10
μM THC were reached at the site of administration; ref. 11). Samples were fixed in formalin, embedded in
paraffin, and used for immunomicroscopy.
Immunomicroscopy of tumor samples. Samples from tumor xenografts were dissected, Tissue-Tek (Sakura) embedded, frozen,
and, before the staining procedures were performed, fixed in acetone for 10 min at
room temperature. Samples from human tumors were subjected to deparaffinization,
rehydration, and antigen retrieval before the staining procedures were performed.
Standard protocols for immunofluorescence or immunohistochemistry microscopy were
used (see Supplemental Methods). Nuclei were counterstained with TOTO-3 iodide (U87MG
and human tumor samples; Invitrogen) or Hoechst 33342 (MEF tumors; Invitrogen).
Fluorescence images were acquired using Metamorph-Offline 6.2 software (Universal
Imaging) and Zeiss Axioplan 2 Microscope.
TUNEL. Tumor samples were fixed, blocked, and permeabilized, and TUNEL was performed as
previously described (13).
Electron microscopy. Ultrastructural analysis of vehicle- and THC-treated cells was assessed by
conventional embedding in the epoxy-resin EML-812 (Taab Laboratories). Ultrathin (20-
to 30-nm-thick) sections of the samples were obtained using a Leica-Reichert-Jung
ultramicrotome and then stained with saturated uranyl acetate–lead
citrate by standard procedures. Ultrathin sections were analyzed in a JEOL 1200-EX II
transmission electron microscope operating at 100 kV.
Statistics. Statistical analysis was performed by ANOVA with a post-hoc analysis using the
Student-Neuman-Keuls test. Differences were considered significant when the
P value was less than 0.05.
Supplemental data
View Supplemental data
Acknowledgments
This work was supported by grants from the Spanish Ministry of Education and Science
(MEC) (HF2005/0021, to G. Velasco; SAF2006/00918, to M. Guzmán; and
BFU2006-00508, to P. Boya), Santander-Complutense PR34/07-15856, to G. Velasco),
Comunidad de Madrid (S-SAL/0261/2006, to M. Guzmán), and La Ligue contre le
Cancer and Canceropole PACA (to J.L. Iovanna). M. Salazar was the recipient of a
fellowship from the MEC. A. Carracedo was the recipient of fellowships from Gobierno
Vasco, the Federation of European Biochemical Societies, and the European Molecular
Biology Organization. M. Lorente and P. Boya have a Juan de la Cierva and a
Ramón y Cajal contract from the MEC, respectively. S.
Hernández-Tiedra has a technician contract from the Spanish Ministry of
Education and the Fondo Social Europeo. The authors thank Dario Alessi (University of
Dundee, Dundee, United Kingdom) for donating anti-PRAS40 antibodies and for technical
support for immunoprecipitation experiments; Gemma Fabriàs, Josefina Casas,
and Eva Dalmau (Instituto de Investigaciones Químicas y Ambientales,
Barcelona, Spain) for analyzing ceramide samples; José Lizcano,
José Bayascas, María M. Caffarel, and Patrizia Agostinis for
their experimental suggestions; and other members of our laboratory for their continual
support.
Footnotes
Conflict of interest: The authors have declared that no conflict of
interest exists.
Nonstandard abbreviations used: Atg, autophagy protein;
eIF2α, eukaryotic translation initiation factor 2α; MEF,
mouse embryonic fibroblast; THC, Δ9-tetrahydrocannabinol;
mTORC1, mammalian target of rapamycin complex 1; PDI, protein disulphide isomerase;
TRB3, tribbles homolog 3.
Citation for this article:
J. Clin. Invest.
119:1359–1372 (2009). doi:10.1172/JCI37948
Arkaitz Carracedo and Ainara Egia’s present address is: Cancer Genetics
Program, Beth Israel Deaconess Cancer Center and Department of Medicine, Beth Israel
Deaconess Medical Center, Harvard Medical School, Boston, Massachusetts, USA.
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