Download A caspaselike activity is triggered by LPS and is

IMMUNOBIOLOGY
A caspaselike activity is triggered by LPS and is required for survival
of human dendritic cells
Luigi Franchi, Ivano Condò, Barbara Tomassini, Chiara Nicolò, and Roberto Testi
Bacterial endotoxin (lipopolysaccharide
[LPS]) is a potent inducer of human dendritic cell (DC) maturation and survival.
Here we show that immature DCs exposed to LPS trigger an early and sustained caspaselike activity, which can be
blocked by zVAD (z-Val-Ala-Asp), in the
absence of detectable caspase 8 and
caspase 10 activation, or poly(ADP-ribose)
polymerase (PARP)–cleaving activity. Preventing LPS-induced caspaselike activation in DC results in massive cell death.
Importantly, triggering of the caspaselike
activity is required for LPS-induced activation of extracellular signal-regulated kinases (ERKs) and for LPS-induced upregulation of cFLIP (Fas-associating
protein with death domain–like interleu-
kin-1␤–converting enzyme [FLICE]–like
inhibitory protein). Therefore, a caspasedependent pathway initiated by LPS controls survival of human DCs. (Blood. 2003;
102:2910-2915)
© 2003 by The American Society of Hematology
Introduction
Dendritic cells (DCs) are specialized leukocytes evolved to provide
a link between innate and adaptive immunity.1 Due to their
extremely high capability in capturing and processing macromolecules, they effectively patrol the interstitial environment of many
tissues and eventually present antigens to naive T lymphocytes to
initiate a specific immune response.2 In addition to acting as very
efficient antigen-presenting cells, they also convey information to
T cells about the presence and the nature of infection and help
direct T-cell responses.3 DCs sense potential pathogens via surface
pattern-recognition receptors (PRRs)4 and, upon engagement of
PRRs, undergo “maturation” (ie, morphologic and functional changes
that include the modulation of adhesion/costimulatory receptors, major
histocompatibility complex [MHC]/peptide complexes, chemokine
receptors, and cytokine production). The complexity of these
responses provides the information required to activate or inhibit
specific T-cell subsets, depending on the type of infection.5,6
PRRs are engaged by pathogen-associated molecular patterns
(PAMPs), structures constitutively and invariantly expressed in
microbes. Among PRRs, Toll-like receptors (TLRs) are characterized by extracellular leucine-rich repeats (LRRs) and by an
intracellular Toll/interleukin 1 (IL1) receptor domain (TIR).7,8
TLRs form a large receptor family counting at least 10 members in
humans, 9 in Drosophila melanogaster and 1 in Caenorhabditis
elegans, generally implicated in microbial immunity. LRRs are
involved in microbe ligand recognition, whereas the cytoplasmic
TIR domain is responsible for receptor/protein interactions and
downstream signal transduction. In fact, human TLR engagement
and clustering results in the recruitment to the receptor of the
adaptor myeloid differentiation primary response 88 (MyD88)
through homophilic interaction with its TIR domain. MyD88 then
interacts with the serine/threonine kinases, IRAKs (interleukin-1
receptor-associated kinase), via homophilic interaction between
their respective death domains (DDs). IRAKs can also be recruited
to the TLR, independently from MyD88, by Toll-interacting
protein (TOLLIP).9 This is followed by IRAK’s autophosphorylation, dissociation from the receptor, and association with tumor
necrosis factor receptor–associated factor 6 (TRAF6). TRAF6
becomes phosphorylated, oligomerizes, and is responsible for
downstream signaling leading to the activation of nuclear factor–␬
B (NF␬B)–dependent genes. TLRs share with the IL1 receptor
(IL1R) important structural features, including the TIR domain,
and most of the intracellular signal transduction machinery.
Lipopolysaccharide (LPS) is a constitutive component of the
bacterial cell wall in Gram-negative bacteria and its invariant
molecular pattern, lipid A, is specifically recognized by Toll-like
receptor 4 (TLR4). TLR4 is expressed and functional in human
DCs.10-12 To be fully competent, TLR4 also requires CD14,
lipopolysaccharide-binding protein (LBP), and MD2. The recruitment to the TLR4 of the private adaptor MyD88 adapter-like
protein/Toll-interleukin 1 precursor domain–containing adapter
protein (MAL/TIRAP) confers to the receptor complex unique
signaling specificities compared with other TLRs and with the
IL1R.13,14 TLR4 indeed can signal to TRAF6 independently from
MyD88 and IRAKs via TIRAP and PKR, an RNA-dependent
protein kinase. LPS/TLR4 interactions in murine DCs eventually
give rise to 2 distinct intracellular signaling pathways, one
involving NF␬B, which controls DC maturation and may proceed
independently from MyD88,15 and the other involving extracellular
signal-regulated kinases (ERKs), which promote DC survival.16,17
Similarly, human DC differentiation appears to involve NF␬B18
and to be negatively regulated by ERK activation.19 No information
From the Laboratory of Immunology and Signal Transduction, Department of
Experimental Medicine and Biochemical Sciences, University of Rome “Tor
Vergata,” Rome, Italy.
Reprints: Roberto Testi, Laboratory of Immunology and Signal Transduction,
Department of Experimental Medicine and Biochemical Sciences, University of
Rome “Tor Vergata,” via Montpellier 1, 00133 Rome, Italy; e-mail:
[email protected].
Submitted March 28, 2003; accepted June 16, 2003. Prepublished online as
Blood First Edition Paper, June 26, 2003; DOI 10.1182/blood-2003-03-0967.
Supported by grants from Associazione Italiana Ricerca sul Cancro, Ministero
Istruzione Universita’ e Ricerca, Agenzia Spaziale Italiana, and European
Commission V Framework Program.
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The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 U.S.C. section 1734.
© 2003 by The American Society of Hematology
BLOOD, 15 OCTOBER 2003 䡠 VOLUME 102, NUMBER 8
BLOOD, 15 OCTOBER 2003 䡠 VOLUME 102, NUMBER 8
is available concerning LPS-initiated intracellular signaling controlling survival in human DCs.
Caspases are a class of cystein proteases that cleave protein
substrates after selected aspartate residues.20 They participate in the
proteolytic modifications of several proteins involved in intracellular signaling pathways governing cellular adaptation, differentiation, proliferation, and apoptosis. They are also primarily responsible for the ordered proteolysis associated with cellular apoptosis.21
At least 11 different caspases have been described in humans and
they can be grouped into different subfamilies according to
structural features, substrate specificity, and function. Caspases
have been shown to participate in ultraviolet B (UVB)–induced
apoptotic cell death of human DCs,22 but little is known about their
involvement in other aspects of DC biology. Here we show that an
early and sustained caspaselike activation participates in the
LPS-generated signaling in DCs and that, importantly, it is required
for LPS-induced survival of human DCs.
Materials and methods
Reagents
Escherichia coli serotype 055:B5 LPS was from Sigma (St Louis, MO).
Human recombinant IL1 was from Pharmingen (San Diego, CA). zVADfmk (z-Val-Ala-Asp(OMe)-fluoromethylketone), z-WEHD-fmk (z-Trp-GluHis-Asp-fluoromethylketone), Ac-YVAD-cmk (Ac-Tyr-Val-Ala-Asp-chloromethylketone), cell-permeable YVAD-CHO (N-acetyl-Ala-Ala-ValAla-Leu-Leu-Pro-Ala-Val-Leu-Leu-Ala-Pro-Tyr-Val-Ala-Asp-CHO), and
z-IETD-fmk (z-Ile-Glu(OMe)-Thr-Asp(OMe)-fluoromethylketone) were
from Biomol (Plymouth Meeting, PA). The cell-permeable cathepsin B
inhibitor zFA-fmk (z-Phe-Ala-fluoromethylketone) and calpain inhibitor
zVF-CHO (z-Val-Phe-CHO) were from Calbiochem (San Diego, CA).
DC generation
Peripheral blood mononuclear cells (PBMCs) from different healthy donors
were isolated on lymphoprep cushions and monocytes were purified by
positive sorting using anti-CD14–conjugated magnetic microbeads according to manufacturer’s instructions (Mylteni Biotech, Auburn, CA). The
recovered cells were 99% CD14⫹ as determined by flow cytometry with
anti-CD14 phycoerythrin (PE)–conjugated antibody (Pharmingen). Cells
were then cultured for 4 to 6 days in 6-well plates (Costar, High Wycombe,
United Kingdom) at the initial concentration of 5 ⫻ 105/mL in RPMI, 10%
fetal calf serum (FCS), penicillin, streptomycin, glutamine, HEPES (N-2hydroxyethylpiperazine-N⬘-2-ethanesulfonic acid) 10 mM, 1% non–
essential amino acids, 1% sodium pyruvate, supplemented with IL4 (2.5
ng/mL; R&D Systems, Minneapolis, MN), and granulocyte-macrophage
colony-stimulating factor (GM-CSF; 50 ng/mL; a kind gift from Prof
Paolo Rossi, University of Rome, Tor Vergata, Italy). Cells were routinely controlled for CD1, CD14, and CD86 expression using the respective PE-conjugated antibodies (Pharmingen) and were consistently
CD14⫺CD1highCD86low.
Antibodies and Western blotting
Cells were washed 2 times with phosphate-buffered saline (PBS) and the
pellet was resuspended in lysis buffer (150 mM NaCl, 10 mM Tris
[tris(hydroxymethyl)aminomethane; pH 7.4], 1 mM EDTA [ethylenediaminetetraacetic acid], 1 mM EGTA [ethylene glycol tetraacetic acid], 1%
Triton X-100, 1% Nonidet P40 [NP40]) and protease inhibitors (1 mM
PMSF [phenylmethylsulfonyl fluoride], 1 ␮g/mL leupeptin, 1 ␮g/mL
aprotinin, 1 ␮g/mL pepstatin, 0.5 ␮g/mL antipain). When necessary, cells
were detached with scrapers in cold PBS. Samples were separated by
sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE)
and transferred to nitrocellulose membrane. Membranes were incubated
with the following antibodies: rabbit polyclonal anti-ERK1/2 (Santa Cruz
Biotechnology, Santa Cruz, CA), mouse polyclonal anti–phospho-ERK1/2
CASPASE ACTIVATION IN DENDRITIC CELLS
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(anti–P-ERK1/2; Santa Cruz Biotechnology), mouse monoclonal anti–
caspase 10 clone 4C1 (MBL, Nagoya, Japan), mouse monoclonal anti–
caspase 8, clone 5F7 (Upstate Biotechnology, Lake Placid, NY), rat
monoclonal anti-FLIP (Fas-associating protein with death domain–like
interleukin-1␤–converting enzyme [FLICE]–inhibitory protein) clone Dave
2. For detection, the appropriate secondary antibodies antimouse, antirabbit, and antirat conjugated to horseradish peroxidase were used (Santa Cruz
Biotechnology) followed by enhanced chemioluminescence. Mouse monoclonal immunoglobulin G1 (IgG1) anti–human tumor necrosis factor ␣
(TNF-␣) was from BD Pharmingen.
Fluorescence-activated cell sorter (FACS) analysis
To detect caspase activity we used CaspACE (Promega, Madison, WI) and
modified the original protocol. Cells were pretreated with 2 ␮M fluorescein
isothiocyanate (FITC)–VAD-fmk for 1 hour, with or without 100 ␮M
zVAD-fmk, and then stimulated with LPS. In some experiments, DCs were
exposed to UV 5400 J/m2 to induce apoptotic cell death, as previously
described.22 After the indicated times, cells were recovered, washed
extensively with cold PBS, and analyzed by a FACScan cytofluorimeter
(Becton Dickinson, San Jose, CA) using CellQuest software (Becton
Dickinson). To detect necrosis, cells were incubated for 20 minutes with 5
␮g/mL propidium iodide in PBS and analyzed by a FACScan cytofluorimeter (Becton Dickinson) using CellQuest software. In some experiments,
cells were subsequently stained with FITC–annexin V in the appropriate
staining buffer (Pharmingen).
In vitro translation and cleavage assay
Full-length human poly(ADP-ribose) polymerase (PARP) and pro-IL1␤
cDNAs, cloned in pGem vector, were used to synthesize [35S]methioninelabeled products by coupled T7 RNA polymerase-mediated transcription
and translation in a reticulocyte lysate system (Promega). Cell pellets were
resuspended in 100 ␮L lysis buffer (50 mM NaCl, 2 mM MgCl2, 40 mM
␤-glycerophosphate, 5 mM EGTA, and 10 mM HEPES, pH 7.0). Cleavage
reactions were performed in a volume of 36 ␮L containing 25 mM HEPES,
pH 7.5; 100 mM NaCl; 2 mM MgCl2; 5 mM dithiothreitol; 0.1% Triton
X-100; 1 mM phenylmethylsulfonyl fluoride; and 2 ␮g/mL aprotinin,
leupeptin, and pepstatin with 3 ␮L [35S]methionine-labeled products and
100 ␮g cell lysates. The reaction was incubated for 1 hour at 37°C for
pro-IL␤ and up to 2 hours at 37°C for PARP. Samples were analyzed by
SDS-PAGE. Gels were fixed (acetic acid 60%, methanol 40%, and glycerol
5%), treated with Amplify solution (Amersham, Arlington Heights, IL), and
dried. Cleavage products were visualized by autoradiography.
Results
LPS induces early caspase activation in immature DCs
To investigate whether caspases participate in LPS-induced signaling in human DCs, we labeled immature DCs with the fluorescent
caspase substrate FITC-VAD-fmk and exposed them to LPS. The
tripeptide VAD can be cleaved by all known caspases. The fmk
group remains irreversibly bound to the protease catalytic pockets,
whereas FITC-VAD is retained intracellularly and fluorescent cells
can be quantitated by FACS analysis. Figure 1A-B shows that 25%
to 30% of DCs exposed to LPS trigger a caspaselike activity within
30 minutes to 1 hour, which can be completely blocked by
pretreatment with the general caspase inhibitor zVAD. By way of
comparison, the activation of caspases as detected after UVB
exposure, an event that can also be blocked by zVAD and is
associated with apoptosis of DCs,22 is also shown.
LPS is known to activate caspase 1 in myeloid cells, an
activation that can be blocked by the caspase 1–like inhibitor
YVAD.23 Yet, the early caspaselike activity induced in DCs by LPS
could not be blocked by YVAD at doses that could inhibit the
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Figure 1. Early caspaselike activation in DCs exposed to LPS. (A) DCs were
loaded with FITC-VAD and then left untreated or exposed for 30 minutes to LPS (100
ng/mL), with or without 1 hour pretreatment with zVAD (40 ␮M) or cell-permeable
YVAD (4 ␮M). FITC-VAD–loaded DCs were also exposed to UVB with or without
pretreatment with zVAD. Cell fluorescence intensity was analyzed by flow cytometry.
(B) The percentage of FITC-positive cells at different time points is shown for each of
the above mentioned conditions. Means ⫾ 1 SD from 3 independent experiments are
shown. Each experiment was performed in duplicate.
activation of caspase 1 (data not shown). This result suggests that
caspases other than caspase 1 are involved in early LPS signaling.
LPS-induced caspase activation is required for DC survival
We therefore investigated the functional relevance of LPS-induced
caspase activation. Immature DCs were pretreated with zVAD to
prevent caspase activation and exposed to LPS. Figure 2A shows
that in the presence of zVAD, LPS-stimulated DCs do not cluster
and undergo massive cell death. Loss of birifrangence and cellular
swelling by microscopic inspection, as well as propidium iodide
exclusion and FACS analysis, revealed that cell death of LPSstimulated DCs in which caspases are blocked by zVAD occurs by
necrosis (Figure 2B).
Cell death of LPS-stimulated DCs pretreated with zVAD
follows a fast kinetic, since about 30% of the cells are dead by 2
hours and about 70% within 6 to 8 hours (Figure 3A). Despite the
structural and signaling similarities between TLR4 and IL1R, IL1
does not induce cell death in zVAD-pretreated DCs, indicating that
critical caspases are selectively activated by LPS (Figure 3B).
Blocking caspases with zVAD also does not interfere with IL1induced maturation of DCs (data not shown). LPS-induced cell
death can be augmented by zVAD in a dose-dependent fashion
(Figure 3C) and can be induced, in the presence of zVAD, at very
low doses of LPS (Figure 3D).
The cathepsin B inhibitor zFA, which is structurally similar to
zVAD, and the caspase 1–like inhibitor YVAD have no influence on
survival of LPS-stimulated DCs (Figure 3E). The calpain inhibitor
zVF causes per se extensive apoptosis of DCs (data not shown),
suggesting that zVAD is not acting by inhibiting calpain in this
system. However, the caspase 4– or 5–like inhibitor WEHD and, more
efficiently, the caspase 8– or 10–like inhibitor IETD, significantly
inhibit survival of LPS-treated DCs, further indicating the requirement for caspases during LPS signaling in DCs (Figure 3E).
A very brief exposure to LPS is sufficient to activate cytoprotective caspase activation. Washing away excess LPS as early as 5
minutes after exposure to DCs does not substantially prevent cell
death if DCs are subsequently treated with zVAD (Figure 3F).
Importantly, blocking caspase activation hours after the exposure to
LPS still results in extensive death of DCs. Figure 3G shows that
even if caspase activation is blocked by zVAD up to 12 hours after
LPS exposure, more than 60% of DCs undergo cell death. When
zVAD is added after 24 to 36 hours, almost no changes in cell
viability can be observed, excluding the possibility of generic
toxicity in LPS-treated cells (Figure 3G). Cytoprotective caspase
activation requires an active metabolic state, since LPS-induced cell
death in the presence of zVAD does not occur at 4°C (Figure 3H).
BLOOD, 15 OCTOBER 2003 䡠 VOLUME 102, NUMBER 8
Finally, we considered the possibility that TNF, secreted in
response to LPS treatment, could be responsible for the observed
effects. To investigate the possible contribution of TNF, we
exposed DCs to zVAD and LPS in the presence of anti-TNF–
blocking antibodies, which we previously tested for the ability to
completely suppress TNF-induced cell death in U937 cells (not
shown). Figure 4 shows that TNF does not contribute significantly
to LPS-induced caspase activation. Together these data indicate
that caspase activation, which follows LPS exposure, is specific,
sustained over time, and required for DC survival throughout the
execution of the maturation program.
LPS does not appear to activate initiator caspases 8 and 10
In an attempt to identify the caspase(s) involved, we investigated
whether the activity of caspases 8 and 10 is triggered by LPS in
DCs. This was suggested by the finding that the caspase inhibitor
IETD significantly affects DC viability in response to LPS (Figure
3E) and by the fact that the signal transduction machinery of TLRs
allows for the recruitment of DD-containing proteins, including the
adaptor FADD (Fas-associated death domain), known to, in turn,
promote the activation of caspases 8 and 10. Interestingly, in fact,
an apoptotic pathway initiated by TLR2 and sequentially involving
MyD88, FADD, and caspase 8 has been described.24 Moreover,
although initiator caspases 8 and 10 have primarily been attributed
to apoptotic pathways, some evidence suggests that they can
participate in entirely different programs.21,25 We therefore analyzed whether caspases 8 and 10 were responsible for LPS-induced
caspase activity in DCs. Figure 5A shows that both caspases are
expressed and functional in immature human DCs, since they can
be triggered by staurosporine exposure, but they are not activated
after LPS exposure as detectable by Western blot analysis of
proteolytic products in DC lysates. Moreover, cell lysates from
LPS-stimulated DCs do not contain proteolytic activity able to
cleave in vitro–translated PARP, a suitable in vitro substrate for
caspases 8 and 10, whereas PARP-cleaving activity can be
recovered from DCs exposed to staurosporine (Figure 5B). By
contrast, LPS can readily trigger caspase 1 activity in DCs, as
detected by the ability of DC lysates to cleave in vitro–translated
pro-IL1␤ (Figure 5C). These data indicate that caspase 8 and 10 are
not likely to play a significant role in this process. Moreover, since
we also observed a limited effect of the WEHD inhibitor on
survival of LPS-treated DCs (Figure 3E), we tested the possible
activation of caspases 4 and 5 in response to LPS. Both caspases
are expressed in DCs but no convincing evidence for their
activation could be obtained (not shown).
Figure 2. Caspaselike activation is required for DC survival. (A) DCs were left
untreated, pretreated with zVAD (1 hour, 40 ␮M), exposed to LPS (100 ng/mL), or
pretreated with zVAD and exposed to LPS. After 16 hours cells were visualized by
microscopy (Olympus IX50; Olympus Optical, Tokyo, Japan). Original magnification,
⫻ 40. Six independent experiments gave similar results. (B) DCs were left untreated,
treated with zVAD (40 ␮M), exposed to LPS (100 ng/mL), or pretreated with zVAD
and exposed to LPS. After 16 hours the cells were analyzed for propidium iodide (PI)
uptake by flow cytometry. Five independent experiments gave similar results.
Horizontal bars indicate necrotic cells.
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CASPASE ACTIVATION IN DENDRITIC CELLS
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Figure 3. LPS-induced caspaselike activation is specific, rapid, and sustained in time. (A) Untreated (E), zVAD (40 ␮M) treated (F), LPS (100 ng/mL) stimulated (‚), or
zVAD pretreated (1 hour) and LPS-stimulated (100 ng/mL) (Œ) DCs were analyzed for PI uptake by flow cytometry at the indicated time points. Means ⫾ 1 SD from 5 different
experiments are shown. (B) Untreated (E), zVAD pretreated (F), IL1 (100 ng/mL) stimulated (‚), or zVAD pretreated and IL1-stimulated (Œ) DCs were analyzed for PI uptake by
flow cytometry at the indicated time points. Means ⫾ 1 SD from 3 different experiments are shown. (C) DCs were pretreated with different doses of zVAD before LPS
stimulation. Cells were analyzed after 8 hours for PI uptake by flow cytometry. (D) DCs were pretreated with 40 ␮M zVAD and then stimulated with different doses of LPS. After
8 hours cells were analyzed for PI uptake by flow cytometry. (E) DCs were pretreated with YVAD (up to 160 ␮M), IETD (up to 40 ␮M), zFA (up to 160 ␮M), WEHD (up to 20 ␮M),
or zVAD (40 ␮M) and exposed to LPS. Cell death was analyzed by FITC–annexin V staining and flow cytometry after 24 hours. Means ⫾ 1 SD from 4 different experiments are
shown. (F) DCs were exposed to LPS. At the indicated time points, cells were washed 3 times and put back in colture with or without zVAD. After 12 hours cells were analyzed
for PI uptake by flow cytometry. Means ⫾ 1 SD from 3 different experiments are shown. (G) DCs left without additional treatment (E) or treated with zVAD (F) at different time
points after LPS stimulation. Eight hours after zVAD addition, cells were analyzed for PI uptake by flow cytometry. Means ⫾ 1 SD from 3 different experiments are shown. (H) A
typical experiment was performed in parallel at 37°C and at 4°C. Twenty-four hours after LPS exposure zVAD pretreated DCs were analyzed for PI uptake by flow cytometry.
Means ⫾ 1 SD from 3 different experiments are shown.
Caspase activation is required for LPS-induced ERK activation
and cFLIP induction
Signals that promote DC maturation and survival may follow
different pathways. LPS induces an early ERK activation, which
appears necessary for DC survival.16,17 It has been shown moreover
that LPS exposure rapidly up-regulates the expression of cFLIP in
human DCs, an event that confers significant survival advantage to
mature DCs.22,26 We therefore investigated whether caspasedependent survival signals controlled ERK activation and cFLIP
induction. Figure 6A shows that 30 minutes after LPS exposure,
ERK phosphorylation can be detected in immature DCs by Western
blot analysis of cellular lysates. LPS-induced ERK activation is
completely blocked by preventing caspase activation with zVAD,
as well as by pretreating DCs with the specific ERK inhibitor
PD98059, suggesting that LPS-activated caspases control ERK
activation. Similarly, blocking caspase activation with zVAD
prevented cFLIP up-regulation observed in DCs 2 hours after LPS
exposure. Importantly, zVAD could not block IL1-induced cFLIP
up-regulation (Figure 6B), again indicating that caspase activation
specifically belongs to LPS-dependent signaling. LPS-induced
Figure 4. TNF does not significantly contribute to LPS-induced caspaselike
activation. DCs were exposed to LPS (100 ng/mL), with or without pretreatment with
zVAD (1 hour, 40 ␮M) and in the presence of the indicated neutralizing concentrations
(2.5-20 ␮g/mL) of anti-TNF–blocking antibodies. After 16 hours cells were analyzed
for PI uptake by flow cytometry. Means ⫾ 1 SD from 3 different experiments are
shown.
caspase activation therefore controls molecular pathways that
critically affect survival of human DCs.
Discussion
Here we provide evidence that a caspaselike activity triggered by
LPS is required for survival of human DCs and critically controls
Figure 5. Caspases 8 and 10 are not activated by LPS. (A) DCs were exposed to
LPS (100 ng/mL) or staurosporine (10 ␮M), with or without pretreatment with zVAD
(1 hour, 40 ␮M), and activation of caspases 8 and 10 was analyzed by Western
blotting at the indicated time points. Three independent experiments gave similar
results. (B) DCs were exposed to LPS (100 ng/mL) or staurosporine (1 ␮M) for the
indicated time points. Cell lysates were then incubated with in vitro–translated
[35S]PARP, with or without 100 ␮M zVAD, and the products of the cleavage reaction
were analyzed by autoradiography. Three independent experiments gave similar
results. (C) DCs were left untreated or exposed to LPS (100 ng/mL) for 2 hours. Cell
lysates were then incubated with in vitro–translated pro-IL1␤, with or without 100 ␮M
zVAD or YVAD, and the products of the cleavage reaction were analyzed by
autoradiography. Two independent experiments gave similar results.
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Figure 6. Caspaselike activation is required for the generation of LPS-induced
survival signals. (A) DCs were left untreated, stimulated with LPS (100 ng/mL),
pretreated for 1 hour with PD98059 (30 ␮M) and stimulated with LPS, or pretreated
for 1 hour with zVAD (40 ␮M) and stimulated with LPS. Thirty minutes after LPS
stimulation, cell lysates were analyzed by Western blotting with a specific anti–P-ERK
antibody. Anti-ERK antibody was used to control loading. Three independent
experiments gave similar results. (B) DCs were left untreated or stimulated with IL1
(100 ng/mL) or LPS, with or without zVAD pretreatment. After 2 hours, cell lysates
were analyzed by Western blotting with an anti-FLIP antibody. Antitubulin antibody
was used to control loading. Five independent experiments gave similar results.
the LPS-induced activation of ERKs and the LPS-induced upregulation of cFLIP.
Proteolytic processing by caspases is a defined modification of
proteins that enables specific cleavage products to activate/lose
functions, acquire new functions, or simply act as physical signal
transducers. As key players in signal generation and transduction,
caspases have been shown to participate in diverse cellular
programs from cell differentiation to adaptation, proliferation, and
apoptosis. During apoptosis, they also accomplish most of the
ordered degradation of structural cellular proteins, which is required for the making of apoptotic bodies. Caspases are expressed
as inactive zymogens, or procaspases, and require proteolytic
processing to allow for the generation of active subunits and
assembly of functional enzymes. Specific structural features in the
prodomain of the zymogens allow the grouping of different
caspases into distinct subfamilies, which may correspond to the
ability to carry out specialized tasks. The presence of protein
interaction death effector domains (DEDs), for instance, distinguishes “initiator” caspases, such as caspases 8 and 10. Initiator
caspases become activated after oligomerization induced by DEDmediated clustering with the adaptor FADD, which in turn is
recruited via its death domain (DD) to DD-expressing surface
receptors. Procaspases 8 and 10 are therefore activated in membraneassociated multiprotein signaling complexes and initiate receptordependent signaling.
Evidence has shown that caspases can be activated following
the engagement of TLRs. Caspase 1 can be activated in human
monocytoid THP1 cells via engagement of TLR2 by bacterial
lipoproteins (BLP), resulting in pro-IL1 proteolytic processing and
IL1 release.25 More importantly, a proapoptotic pathway can also
be initiated by TLR2 in cycloheximide-treated THP1 cells. This
pathway may sequentially involve MyD88, FADD, and caspase 8
and be counteracted by a parallel MyD88-dependent pathway,
which includes TRAF6 and leads to NF␬B activation.24,27 LPS, a
ligand for TLR4, can induce apoptosis in multiple cellular systems.
A TLR4-mediated apoptotic pathway can be triggered by LPS in
cycloheximide-treated endothelial cells and can be blocked by
zVAD.28,29 Concomitantly, LPS triggers survival pathways in
several cell types, including human DCs, which result in the
up-regulation of cFLIP, an inhibitor of the proapoptotic initiator
caspases.22,26 Suppression of the protein synthesis–dependent upregulation of cFLIP greatly enhances LPS-induced apoptotic cell
death,30 indicating that apoptotic signaling by TLRs runs in parallel
with survival pathways, the final outcome depending on reciprocal
inhibition and general integration.
Proapoptotic caspases can therefore be triggered by microbial
products via TLRs, together with molecular strategies designed to
block their activation. Quite differently, our data show that LPS, the
primary component of the outer membrane of Gram-negative
BLOOD, 15 OCTOBER 2003 䡠 VOLUME 102, NUMBER 8
bacteria, triggers a caspaselike activity that is required for survival
of human DCs. No information is currently available concerning
the involvement of caspases in TLR-dependent survival pathways
in mammals.
The fruit fly Drosophila melanogaster produces a number of
antibiotic peptides to counteract microbial infections. The genes
coding for these peptides are under the control of 2 different
pathways.31 The Toll pathway, required for the immune response to
Gram-positive bacteria and fungi, is initiated by surface Toll
receptors. Similarly to mammalian TLRs, Toll interacts with either
dMyD88 via TIR or with the DD-containing adaptor Tube. The
DDs of dMyD88 or Tube allow for further interaction with Pelle, a
serine/threonine kinase homologous to IRAK. This is followed by
the downstream phosphorylation of Cactus, an NF␬B inhibitor
(I␬B) homolog expressing ankirin repeats, and its degradation and
dissociation from dorsal-related immunity factor (DIF), a transcription factor of the Rel family. Once rid of Cactus, DIF translocates to
the nucleus where it acts as the major transactivator of genes
coding for anti–Gram-positive and antifungal peptides.32
The pathway controlling the peptide response to Gram-negative
bacteria in Drosophila is less understood. Recently, PGRP-LC, a
member of the family of peptidoglycan recognition surface receptors (PGRPs), has been shown to control anti–Gram-negative
bacterial responses.33-35 Genetic analysis has revealed that this
pathway requires immune deficiency (IMD), a DD-containing
adaptor homologous to mammalian receptor interacting protein
(RIP).36 IMD acts upstream of dFADD, a DD-containing adaptor.37,38 dFADD may allow the recruitment of DREDD (deathrelated CED-3/NEDD2-like protein), a caspase 8 homolog and the
only caspase found so far involved in microbial defense in
Drosophila.39 Downstream of IMD is also dTAK1 (Drosophila
homolog of transforming growth factor-␤–activated kinase 1), a
mitogen-activated protein 3 (MAP3) kinase, homolog to human
TAK1. dTAK1 is probably involved in the activation of the I␬B
kinase (IKK) complex required for the nuclear translocation of the
transcription factor Relish.40 Relish, another member of the Rel
family, is activated by phosphorylation and by proteolytic cleavage
necessary to release an ankirin repeat–rich Cactus-like domain and
allow for nuclear translocation. Relish controls the transactivation
of genes coding for anti–Gram-negative bacterial peptides. Importantly, the PGRP-IMD-Relish pathway is also triggered by LPS.35,41
In mammals, peptidoglycans use TLR2, whereas LPS uses
TLR4 to induce DC maturation.42,43 Following LPS/TLR4 interactions, the DD of MyD88 is in principle available for interactions
with FADD allowing further recruitment and activation of initiator
as well as apoptotic caspases.24 However no apoptotic caspase
activation is detectable during LPS signaling in DCs. Survival is
therefore critically dependent on LPS-induced cytoprotective pathways, including cFLIP expression. Intriguingly, we find that
LPS-induced cFLIP up-regulation itself requires the activation of a
caspaselike activity. Moreover, our data indicate that the early ERK
activation, which also contributes to DC survival, is under the
control of upstream caspases.
It is remarkable that caspase activation is relevant for LPS but
not for IL1-dependent signaling in DCs. Blocking caspases, in fact,
does not prevent IL1-induced cFLIP up-regulation, maturation, and
survival. Similarly to TLR4, the IL1 receptor signals via MyD88 or
TOLLIP to IRAKs and TRAF6. However, MAL/TIRAP-dependent
pathways are unique to TLR4.13,14 Early caspase activation may
therefore contribute to the specificity of LPS-induced signaling.
The identity of the critical caspase(s) is currently missing, since
it does not seem to correspond to canonical initiator caspases 8 or
BLOOD, 15 OCTOBER 2003 䡠 VOLUME 102, NUMBER 8
CASPASE ACTIVATION IN DENDRITIC CELLS
10, despite its very early activation during LPS signaling. Accordingly, the caspase 8 homolog DREDD does not seem to be
implicated in early signaling along the PGRP-Relish pathway, as
defined by epistatic analysis.40 An attractive candidate could be the
human paracaspase MLT/MALT1 (mucosa-associated lymphoid
tissue lymphoma translocation gene 1),44-46 which possesses a DD
highly homologous to the DD of MyD88 and IRAKs as well as to
Drosophila Pelle and Tube.47 Further investigation is required to
identify the relevant caspase(s) involved in LPS signaling in
2915
DCs and to better clarify the intimate connections between survival and death decisions governing cellular adaptation to microbial organisms.
Acknowledgments
The authors acknowledge Dario Serio for technical help and all lab
members for insightful discussions.
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