Exploring the prime site in caspases as a novel chemical strategy for understanding the mechanisms of cell death: a proof of concept study on necroptosis in cancer cells

Article abstract of DOI:10.1038/s41418-019-0364-z

Caspases participate in regulated cell death mechanisms and are divided into apoptotic and proinflammatory caspases. The main problem in identifying the unique role of a particular caspase in the mechanisms of regulated cell death is their overlapping sub

Full text of DOI:10.1038/s41418-019-0364-z

Cell Death & Differentiation
https://doi.org/10.1038/s41418-019-0364-z
ARTICLE
Exploring the prime site in caspases as a novel chemical strategy for
understanding the mechanisms of cell death: a proof of concept
study on necroptosis in cancer cells
Katarzyna Groborz1 Monica L. Gonzalez Ramirez2 Scott J. Snipas2 Guy S. Salvesen2 Marcin Drąg1,2
Marcin Poręba1,2
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Received: 19 July 2018 / Revised: 2 May 2019 / Accepted: 23 May 2019
© The Author(s), under exclusive licence to ADMC Associazione Differenziamento e Morte Cellulare 2019
Abstract
Caspases participate in regulated cell death mechanisms and are divided into apoptotic and proinflammatory caspases. The
main problem in identifying the unique role of a particular caspase in the mechanisms of regulated cell death is their
overlapping substrate specificity; caspases recognize and hydrolyze similar peptide substrates. Most studies focus on
examining the non-prime sites of the caspases, yet there is a need for novel and more precise chemical tools to identify the
molecular participants and mechanisms of programmed cell death pathways. Therefore, we developed an innovative
chemical approach that examines the prime area of the caspase active sites. This method permits the agile parallel solid-
phase synthesis of caspase inhibitors with a high yield and purity. Using synthesized compounds we have shown the
similarities and differences in the prime area of the caspase active site and, as a proof of concept, we demonstrated the
exclusive role of caspase-8 in necroptosis.
Introduction
activation in the cytoplasm. Of the 11 human caspases [2],
members of the inflammatory group (caspase-1, caspase-4,
caspase-5) are vital for the activation of specific cytokines
and inflammatory cell death or pyroptosis, while members
of the apoptotic group (caspase-3, caspase-6, caspase-7,
caspase-8, caspase-9, and caspase-10) participate in the
activation and progression of apoptosis.
Caspases, the main proteolytic enzymes that mediate regu-
lated cell death pathways, are a structurally and functionally
related family of cysteine proteases that owe their name to
the stringent requirement for aspartic acid in the P1 position
of their peptide substrates [1]. Caspases are found in all
metazoans and expressed as inactive zymogens that await
The other type of allegedly programmed cell death is
necroptosis, however, this process has been investigated
much less extensively that above-mentioned apoptosis and
pyroptosis. According to recent recommendation, necrop-
tosis means “necrotic cell death dependent on receptor-
interacting protein kinase-3 (RIPK3)” [3]. This process is
caspases-independent, as their activity has to be silenced so
that the cell could be destroyed in programmed fashion.
Necroptosis was first recognized as a type of cell death that
can be provoked by treatment with TNF (tumor necrosis
factor), exclusively in the presence of caspase inhibitors
such as zVAD-fmk which inhibits or disrupts the function
of caspase-8 [4]. Apoptosis and necroptosis share several of
the upstream signaling factors and the execution of either
death pathway is coordinated by the array of regulatory
molecules, including the FLIP protein [5], the cellular
inhibitors of apoptosis [6], deubiquitinases A20 and CYLD
[7]. So far the key caspase that needs to be blocked in order
Edited by A. Ashkenazi
Supplementary information The online version of this article (https://
doi.org/10.1038/s41418-019-0364-z) contains supplementary material,
which is available to authorized users.
* Marcin Drąg
marcin.drag@pwr.edu.pl
* Marcin Poręba
marcin.poreba@pwr.edu.pl
1
Department of Bioorganic Chemistry, Faculty of Chemistry,
Wroclaw University of Science and Technology, Wyb.
Wyspianskiego 27, 50-370 Wroclaw, Poland
2
NCI Designated Cancer Center, Sanford Burnham Prebys Medical
Discovery Institute, 10901 North Torrey Pines Road, La Jolla, CA
92037, USA

K. Groborz et al.
to trigger necroptosis is caspase-8, however, there is
growing body of evidence, that the inhibition of caspase-10,
another apical protease, can be important to this process [8].
There exists a substantial problem with defining the role
of individual caspases in the control of cell death and sur-
vival—namely their cross-reactivity with available chemical
tools [9]. Even with the development of highly selective,
well-defined substrates, there is a need for tools that dis-
criminate between the individual caspases. This difficulty is
a consequence of the overlap in consensus sequences
recognition by caspases [10–12], i.e., one caspase can
efficiently hydrolyze a substrate designed for another cas-
pase. To address this obstacle, chemical biology strategies
in which peptide substrates and inhibitors are screened
against individual caspases have been utilized [13]. While
this delivers distinguishing characteristics, current meth-
odologies are restricted to half of the available binding
interactions. The substrate binding cleft of a protease is
defined by the surfaces that interact with the substrate
side chains. The cleft is divided into a non-prime region that
binds substrate amino acids N-terminal to the scissile bond,
and a prime region that binds amino acids C-terminal to the
scissile bond [14]. Numerous studies on the kinds of cas-
pase substrates that can be used to explore the prime and
non-prime active sites have been published [13, 15].
However, in the case of inhibitors, the prime region is
occupied by a reactive warhead [16] leaving the non-prime
region to explore for inhibitor design.
There have been very few attempts to explore the prime
region using inhibitors [17], and none have adopted wide-
ranging library-based approaches. To provide an alternative
caspase inhibitor design we first needed to employ a war-
head that permitted amino acid extension into the prime
region. We then crafted a simple, straightforward method
based on solid-phase peptide synthesis and generated two
libraries of peptidyl inhibitors intended to explore this
region. We screened apoptotic caspases-3, -7, -8, -9, and -10
with the libraries, aiming specifically to generate diagnostic
agents that could distinguish between the closely related
caspase-8 and caspase-10. We selected the best hits to
analyze paradigms of apoptosis, whose mechanisms depends
on caspases [18–20], and necroptosis, which is negatively
mediated by caspases [21]. Our compounds allowed us to
differentiate the biological outcomes of caspase inhibition in
cancer cells stimulated with various cell death inducers.
synthesis of the inhibitor libraries and individual substrates,
we used Rink Amide AM resin (loading 0.74 mmol/g, Iris
Biotech GmbH), Fmoc-protected amino acids (purity
>98%, Iris Biotech GmbH, Bachem, Creosalus, P3 Bio-
Systems, QM Bio), N-hydroxybenzotriazole (HOBt mono-
hydrate purity >98%, Creosalus), diisopropylcarbodiimide
(DICI, peptide grade, Iris Biotech GmbH), HATU and
HBTU (peptide grade, ChemPep Inc.), 2,4,6-trimethylpyr-
idine (2,4,6-collidine, peptide grade, Sigma-Aldrich), N,N-
dimethylformamide (DMF, peptide grade, WITKO), acet-
onitrile (ACN, HPLC grade, WITKO), piperidine (PIP,
peptide grade, Iris Biotech GmbH), trifluoroacetic acid
(TFA, purity 99%, Iris Biotech GmbH), triisopropylsilane
(TIPS, purity 99%, Sigma-Aldrich), methanol (MeOH,
analytical grade, POCh), dichloromethane (DCM, analytical
grade, POCh), diethyl ether (Et2O, analytical grade, POCh),
acetic acid (AcOH, purity 98%, POCh) and phosphorus
pentoxide (P2O5, purity 98%, POCh). 2-chlorotrityl chloride
resin (100–200 mesh, 1.59 mmol/g) was purchased from Iris
Biotech GmbH. N,N-diisopropylethylamine (DIPEA) was
purchased from VWR International (Gdansk, Poland).
2,2,2-trifluoroethanol (TFE), HBr (30% wt. in AcOH),
anhydrous tetrahydrofuran (THF), potassium fluoride (KF,
purity >99,97%), 4-methylmorpholine (NMM), isobutyl-
chloroformate (IBCF), tere-phthalic acid (TPA) and iso-
phthalic acid (IPA) were purchased from Sigma-Aldrich.
Diazomethane was generated according to the Aldrich
Technical Bulletin (AL-180) protocol. For the apoptosome
activation in the HEK293F cell-free system, horse cyto-
chrome c and dATP were purchased from Sigma-Aldrich
and the proteasome inhibitor MG-132 was purchased from
ApexBio (Houston, USA). Methylated zVAD(O-Me)-fmk
and non-methylated zVAD-fmk were purchased from
Cayman Chemical (Michigan, USA). Recombinant human
TNF-α was purchased from R&D Systems (cat. no. 210-
TA, Minneapolis, USA). Necrostatin-1 (Nec-1) was
obtained from Santa Cruz Biotechnology (Dallas, USA).
The SMAC mimetic birinapant (TL32711) was purchased
from Active Biochem (Maplewood, USA). Individual
inhibitors and substrates were purified by HPLC on a
Waters M600 solvent delivery module with a Waters
M2489 detector system using a semi-preparative Waters
Spherisorb S10ODS2 column and a semi-preparative Dis-
covery® C8 HPLC column (particle size 10 µm). The solvent
consisted of phase A (water/0.1% TFA) and phase B
(acetonitrile/0.1% TFA). The purity of each compound was
confirmed by an analytical HPLC system using a Waters
Spherisorb S5ODS2 column and analytical Discovery® C8
column (particle size 5 µm). The molecular weight of each
inhibitor and substrate was confirmed by high-resolution
mass spectrometry on a High Resolution Mass Spectrometer
WATERS LCT premier XE with Electrospray Ionization
(ESI) and a Time of Flight (TOF) module.
Materials and methods
Reagents
All chemicals and reagents were purchased from commer-
cial suppliers and used without further purification. For the

Exploring the prime site in caspases as a novel chemical strategy for understanding the mechanisms of. . .
Synthesis of the inhibitor libraries
ethereal diazomethane (16.6–21.4 mmol) at 0°C and stirred
vigorously for 20 min. Then, the ice bath was removed and
the reaction was carried out for another hour at room tem-
perature. To generate Cbz-VAD(tBu)-CH2Br, 15 mL of a
1:2 solution of HBr (30% wt. in CH3COOH) and water
were slowly added to the mixture over a period of 10 min.
The reaction was monitored using analytical HPLC. After
30 min, the reaction was complete and the mixture was
diluted with ethyl acetate, transferred to a separatory funnel
and extracted three times with saturated aqueous NaHCO3,
twice with brine and once with water. The organic fraction
was dried over MgSO4 and evaporated under reduced
pressure. The product was a pale yellow solid that was used
for further synthesis without purification. The product pur-
ity, determined by HPLC, was > 90% and the overall yield
was > 85%.
(1) zVAD-bmk synthesis
The Cbz-Val-Ala-Asp(tBu)-OH peptide was synthesized on
a 2-chlorotrityl chloride resin using Fmoc-protected amino
acids. Five grams of 2-chlorotrityl chloride resin (7.95
mmol) were resuspended in 10 mL of anhydrous DCM and
gently stirred once per 5 min for 30 min. Then, the DCM
was filtered off and the resin was washed three times with
fresh portions of DCM. Next, 3 eq of Fmoc-L-Asp(tBu)-OH
amino acid in anhydrous DCM were pre-activated with 3 eq
of DIPEA and added to the resin. The suspension was
gently agitated for 4 h at room temperature. Subsequently,
the mixture was filtered off and the resin was washed three
times with DCM and three times with DMF to dispose of
the excess reagent. The Fmoc-protecting group was
removed using a 20% solution of piperidine in DMF (three
cycles: 5 min, 5 min and 25 min). After the Fmoc group de-
protection, a ninhydrin test was carried out. Next, the resin
was washed six times with DMF and 3 eq of Fmoc-L-Ala-
OH, pre-activated with 3 eq of HOBt and then, 3 eq of DICI
in DMF was added to the reaction vessel. The reaction was
carried out for 3 h at room temperature. After coupling the
second amino acid, the Fmoc-protecting group was
removed in the same manner as previously described and
the last amino acid, Cbz-L-Val-OH, was coupled using
HOBt and DICI. After solvent removal, the resin was
washed three times with DMF, three times with DCM and
three times with MeOH, then dried overnight over P2O5.
The tripeptide was cleaved from the resin using the mild
acidic condition of a mixture of TFE/AcOH/DCM (%, v/v/v
10:10:80), thus preserving the t-Bu protecting group on the
side chain residue of aspartic acid. The liquid product was
then mixed with hexane and the solvents were evaporated
under reduced pressure. The resulting product was dis-
solved in an ACN/H2O mixture (%, v/v 75:25) and lyo-
philized. The purity of the resulting compound was
confirmed by analytical HPLC and its molecular weight was
determined by HR-MS. The crude tripeptide was then con-
verted into a bromomethyl ketone (BMK) using a synthetic
procedure described by Kato et al. [22]. Briefly, a 0.2 M
solution of Cbz-VAD(tBu)-OH (5 mmol, 1 eq) in anhydrous
THF was stirred in an ice/acetone bath at −10°C for 20 min.
Then, 4- methylmorpholine (6.25 mmol, 1.25 eq) and iso-
butyl chloroformate (5.75 mmol, 1.15 eq) were added to the
mixture. Immediately after adding the isobutyl chlor-
oformate, a white precipitate formed that indicated the
formation of an anhydride. The reaction was carried out at
−10°C for 45 min. In a parallel experiment, an ethereal
solution of diazomethane was generated according to the
Aldrich Technical Bulletin protocol (AL-180). Next, the
solution of mixed anhydrides was added dropwise to the
(2) zVAD-iso-Pht-X-NH2 and zVAD-tere-Pht-X-NH2 library
synthesis
0.3 g of Rink Amide Resin (0.222 mmol, 1 eq) was suspended
in 5 mL of DCM and gently stirred once every 5 min for 30
min. The DCM was removed by filtration and the resin was
washed three times with DMF. Next, the Fmoc group was de-
protected using 20% piperidine in DMF (three cycles of 5 min,
5 min and 25 min). The resin was washed six times with DMF,
three times with DCM, and three times with MeOH then dried
over P2O5. The resin was divided into 19 equal portions, each
of which was added to 19 wells of a solid-phase peptide
synthesis reactor. To swell the resin, 1 mL of DCM was added
to each well and left there for 30 min, shaking gently once
every 5 min. Afterward, the DCM was filtered and the resin
was washed three times with DCM. Next, 3 eq of each of the
18 natural amino acids and norleucine were pre-activated with
HATU and 2,4,6-collidine in a minimal amount of DMF and
added to separate wells of the solid-phase peptide synthesis
reactor (Asp amino acids were omitted due to synthesis issues).
The reaction was carried out for 4 h, followed by three DMF
washes of the solid-phase. The Fmoc-protecting group was
removed and the resin was washed six times with DMF. Iso-
phthalic acid was coupled to the free amine group of each of
the 18 natural amino acids and norleucine with the use of the
same coupling reagents (3 eq of HATU and 3 eq of 2,4,6-
collidine). After 3 h of the coupling reaction, the resin was
washed three times with DMF. In the subsequent step, 3 eq of
Cbz-VAD(tBu)-BMK were dissolved in a minimal amount of
DMF, pre-activated with 10 eq of KF and added to each well
of the reactor. The reaction was carried out for 24 h with gentle
agitation. The resin was then washed ten times with DMF,
three times with DCM, and three times with MeOH then dried
over P2O5. Finally, the inhibitors were cleaved from the resin
using a mixture of TFA:TIPS:H2O (%, v/v/v 95:2.5:2.5) for 2 h
at room temperature, shaking once per 10 min. Supernatants

K. Groborz et al.
were collected in 15 mL Falcon conical-bottom centrifuge
tubes and the resin was washed once again with TFA:TIPS:
H2O (%, v/v/v 95:2.5:2.5). For each compound, both super-
natant fractions were combined and precipitated in cold Et2O
for 30 min at −20°C. Then, the samples were centrifuged
(5 min, 4°C, 4400 rpm), the supernatants were decanted and the
precipitates were washed with 10 mL of Et2O, then centrifuged
(5 min, 4°C, 4400 rpm) and the supernatant was discarded. The
precipitated compounds were dissolved in 0.8 mL of DMSO
and purified using HPLC (Waters, column: Spherisorb, 5 µm
particle size, L × I.D. 25 cm × 4.6 mm) with an H2O:ACN
gradient and then lyophilized. The purified white/pale yellow
products were dissolved in DMSO to a final concentration of
10 mM and stored at −80°C until used. The second library
with tere-phthalic acid and the zVAD-2,6-dimethylter-
ephthalic-X-NH2 inhibitors was synthesized in an analogous
manner. The purity of each inhibitor was confirmed by ana-
lytical HPLC and analyzed using HRMS (Table S4, Fig. S5).
mixture was filtered and the resin was washed six times with
DMF. The Fmoc-protecting group was removed using 20%
piperidine in DMF. A ninhydrin test was carried out after
each round of coupling and Fmoc de-protection. A solution
of 2.5 eq of Fmoc-P3-OH, 2.5 eq of HOBt and 2.5 eq of DICI
in DMF was added to the resin and the resin was agitated for
3 h. After removal of the solution, the resin was washed with
DMF and coupling and de-protection of Fmoc-P4-OH was
performed in the same manner as for the P2 position. The N-
terminus was protected with an acetyl group using 5 eq of
AcOH, 5 eq of HBTU and 5 eq of DIPEA in DMF. After
solvent removal, the resin was washed six times with DMF,
three times with DCM and three times with MeOH, dried
over P2O5 and cleaved from the resin with a mixture of TFA/
TIPS/H2O (%, v/v/v 95: 2.5: 2.5). The crude product was
purified by HPLC and lyophilized. The purity of each sub-
strate was confirmed by analytical HPLC and analyzed using
HRMS. The substrates were then dissolved in peptide grade
DMSO to a concentration of 20 mM and stored at −80°C
until used.
Individual substrate synthesis
To determine the kinetic parameters of the obtained inhibitors
we synthesized broad-spectrum, ACC-labeled fluorogenic
caspase substrates using a previously published method [23].
The Fmoc-ACC-OH fluorophore was synthesized according
to Maly et al. [24]. Briefly, Rink Amide Resin was added to a
glass reaction vessel containing DCM and was stirred gently
every 10 min for 1 h then filtered and washed with DMF. The
Fmoc-protecting group was removed using 20% piperidine in
DMF and the resin was washed six times with DMF. Next,
2.5 eq of Fmoc-ACC-OH were pre-activated with HOBt
monohydrate and DICI in DMF and the mixture was poured
onto the resin. After 24 h of gentle agitation at room tem-
perature, the Fmoc-ACC-OH coupling reaction was repeated
to improve the amount of ACC coupling to the resin. When
the reaction was complete, the resin was washed with DMF
and the Fmoc-protecting group was removed with 20%
piperidine in DMF. Next, 2.5 eq of Fmoc-L-Asp(tBu)-OH,
2.5 eq HATU and 2.5 eq of collidine in DMF were activated
for 3 min and added to the reaction vessel containing H2N-
ACC-resin and the reaction was carried out for 24 h. The
resin was washed four times with DMF and the reaction was
repeated using 1.5 eq each of Fmoc-L-Asp(tBu)-OH, HATU,
and collidine in DMF. After the resin was washed with DMF,
the Fmoc-protecting group was removed using 20% piper-
idine in DMF. The resin was subsequently washed three
times with DMC and three times with MeOH and dried over
P2O5. Next, the resin was divided and individual substrates
were synthesized by solid-phase peptide synthesis. The 2.5
eq of Fmoc-P2-OH were pre-activated with 2.5 eq of HOBt
and 2.5 eq of DICI in DMF and poured into the cartridge
with 1 eq of NH2-Asp(tBu)-ACC-resin, followed by gentle
agitation for 3 h at room temperature. Then, the reaction
Recombinant caspase assay conditions
Recombinant human apoptotic caspase-3, caspase-7, cas-
pase-8, caspase-9, and caspase-10, recombinant human
proinflammatory caspase-1, caspase-4, and caspase-5 and
recombinant mouse proinflammatory caspase-1, and
caspase-11 were expressed and purified as previously
described [25, 26]. All the enzymes were active site-titrated
using the zVAD(OH)-fmk inhibitor (cat. no. 14467, Cay-
man Chemical Company) as previously described [25]. The
caspase buffer consisted of 20 mM Pipes, 10 mM NaCl, 1
mM EDTA, 10% sucrose (w/v) and 10 mM DTT (pH 7.2).
To allow dimerization and to maintain the maximal activity,
the buffer for the apical caspases (8, 9, and 10) and the
proinflammatory caspases (1, 4, 5, and 11) was supple-
mented with 0.75 M sodium citrate.
Determination of the kinetic parameters for the
inhibitors
We measured the Ki parameters for the reversible inhibitors
using the Morrison equation [27]. For each enzyme, an
appropriate fluorogenic substrate was used to monitor sub-
strate hydrolysis (mCasp-1, Ac-LEHD-ACC KM = 36 μM;
mCasp-11, Ac-LEHD-ACC KM = 165 μM; hCasp-1, Ac-
LEHD-ACC KM = 49 μM; hCasp-3, Ac-DEVD-ACC KM
= 21 μM; hCasp-4, Ac-LEHD-ACC KM = 112 μM; hCasp-
5, Ac-LEHD-ACC KM = 124 μM; hCasp-7, Ac-DEVD-
ACC KM = 57 μM; hCasp-8, Ac-LEHD-ACC KM = 15 μM;
hCasp-9, Ac-LEHD-ACC KM = 106 μM; hCasp-10, Ac-
LEHD-ACC KM = 24 μM). To meet the criteria for rever-
sible inhibitor kinetics, the lowest inhibitor concentration

Exploring the prime site in caspases as a novel chemical strategy for understanding the mechanisms of. . .
was at least 4-fold higher than the caspase concentration
used in the assay. The caspase was first pre-warmed in the
assay buffer in the 96-well plate for 15 min at 37°C and then
preincubated with the inhibitor for another 15 min at 37°C.
Next, the substrate was added to the enzyme-inhibitor
mixture and the progress of the reaction was monitored
using CLARIOstar plate reader (BMG Labtech) operating
in fluorescence kinetic mode (355 nm excitation and 460 nm
emission). The kinetic parameters (kobs/I) for the irreversible
inhibitors (tagged with an AOMK warhead) were deter-
mined under pseudo-first order kinetic conditions, where the
lowest inhibitor concentration was at least 5 times higher
than the caspase concentration. The inhibitor was diluted
into the wells of the 96-well plate, mixed with the fluor-
escent substrate and pre-warmed for 15 min at 37°C. In
parallel, the caspase was incubated at 37°C for 15 min,
poured onto the plate and the reaction was started imme-
diately under the same fluorescence setting as that of the
reversible inhibitors. The reaction was monitored until the
enzyme was completely inhibited. All the measurements for
the reversible and irreversible inhibitors were performed at
least three times and the data were analyzed using Graph
Pad Prism 7.0 software (La Jolla, CA, USA).
was preincubated at either low or high concentrations with an
excess of FLIP protein and subjected to kinetic analysis using
the Ac-LEHD-ACC substrate. Next, to optimize the kinetic
assay conditions, caspase-8 was titrated with FLIP for 15 and
30 min and then reacted with Ac-LEHD-AFC.
Determination of the inhibition mode of selected
inhibitors for caspase-8, caspase-10, and caspase-8:
FLIP
To test whether the inhibitors were reversible or irrever-
sible, we competitively labeled caspase-8, caspase-8:FLIP,
and caspase-10. Briefly, 50 nM of active site-titrated cas-
pase was preincubated for 30 min with 1 μM selected cas-
pase inhibitor, followed by a 15 min incubation with a
potent and irreversible caspase activity-based probe (100
nM, biotin-ahx-LEHD-AOMK, total reaction volume—100
μL). Next, the reaction products were precipitated using
100 μL of ice-cold 30% trichloroacetic acid (TCA) in water.
TCA was added to each sample, incubated on ice for 30 min
and centrifuged for 30 min (20,000 × g, 4°C). The super-
natants were gently aspirated using a vacuum device, the
pellet was resuspended in 200 μL of ice-cold 10% TCA and
the samples were centrifuged for an additional 15 min
(20,000 × g, 4°C). The supernatants were again removed
and the pellet resuspended in 200 μL of acetone. The
samples were centrifuged for 10 min (20,000 × g, 4°C) and
the supernatant was removed. The samples were dried at
37°C for 15 min and then 80 μL of 1× SDS/DTT buffer was
added to each sample, followed by denaturation at 95°C for
5 min. Then, 30 μL of each sample was subjected to SDS-
PAGE (30 min, 200 V), followed by transfer to a nitro-
cellulose membrane (60 min, 10 V) and blocking with 2%
BSA in TBS-T (60 min, RT). The biotin-labeled probe/
caspase complex was detected by probing the membrane
with fluorescently labeled streptavidin (30 min, RT,
1:10,000, IRDye® 800CW, LI-COR), followed by scanning
in a LI-COR system (800 nm channel, Odyssey® CLx,
Lincoln, NE, USA). The data were analyzed using Image
Studio Lite software (Lincoln, NE, USA). The use of 1 μM
(inhibitor-enzyme 20x excess) of zVAD-Bzl-X and zVAD-
AOMK-X inhibitor ensured that all caspase activity was
blocked before adding the LEHD activity-based probe. In
the second experiment, caspase-8, caspase-8:FLIP and
caspase-10 (100 nM) were preincubated with select inhibi-
tors (1 µM) for 30 min and then the caspase-inhibitor
complex was diluted 20× in assay buffer (final caspase
conc. 5 nM, final inhibitor conc. 50 nM), the Ac-LEHD-
AFC (100 μM) was added and the fluorescence was mon-
itored immediately for 30 min. The dissociation of the
caspase-inhibitor complex and the recovery of the enzyme
activity indicated reversible inhibition. In a separate
experiment, 5 nM of active site-titrated caspase was mixed
Specificity in the prime region of the caspases
The Ki parameters of all 39 inhibitors for the individual
recombinant caspases were used to create specificity profiles
(heat maps) in the prime areas of the active sites of the cas-
pases. For better data presentation, the Ki for the reference
inhibitor (zVAD-Bzl) for each caspase was separately set at 1
and the Ki parameter for the other inhibitors was adjusted
accordingly. The heat maps were created in Graph Pad
Prism 7.0.
Correlation of inhibition profiles between caspases
To identify the differences between inhibitor specificities across
caspases, we generated correlation matrices for the proin-
flammatory caspases and the apoptosis caspases, where we
calculated the Pearson correlation coefficient (ρ) for each cas-
pase pair. All calculations were performed using Graph Pad
Prism 7.0 and presented either as a heat map or a scatter plot.
Caspase-8:FLIP complex formation and activity
assays
FLIP protein was expressed and purified according to the
procedure described by Boatright et al. [28]. The caspase-8:
FLIP complex was assembled in regular caspase buffer sup-
plemented with 0.75 M of sodium citrate. Caspase-8 mono-
mers dimerize easily in citrate buffer, therefore, to avoid the
proteolytic background from dimerized caspase-8, caspase-8

K. Groborz et al.
with 50 nM of the selected inhibitor and 100 μM Ac-LEHD-
AFC and the fluorescence of the reaction was measured.
zVAD-Bzl inhibitors were added to wells and preincubated
for 2 h prior to TRAIL (100 ng/mL) and CHX (5μg/mL)
addition. Cell were harvested at various timepoints, and
subjected for SDS-PAGE and Western blotting. Caspase-8
processing was detected with an anti-caspase-8 mouse
serum as described above.
Apoptosis and necroptosis assays
We used HT-29 human colon cancer cells, which respond
well to apoptosis and necroptosis stimulants, to test whether
we could use our inhibitors to identify caspases’ involve-
ment in apoptosis vs. necroptosis. HT-29 cells were seeded
on a 96-well cell culture plate (20,000 cells per well in 100
μL of phenol red-free McCoy’s media) and allowed to
attach overnight. The next day, the cells were preincubated
for 6 h with or without caspase inhibitors (25 μM) and with
or without Nec-1 (50 μM). After this, the cells were sti-
mulated with TRAIL (100 ng/mL) + CHX (5μg/mL)
[29, 30] or TNF-α (100 μg/mL) + birinapant (2 μM) to
induce apoptosis [31, 32]. After 24 h, the HT-29 cell via-
bility was determined using MTS assay according to the
manufacturer’s protocol (CellTiter 96® Aqueous One Solu-
tion Cell Proliferation Assay, G3582, Promega). The
number of living cells was reported as a survival ratio cal-
culated based on the number of un-stimulated control cells.
Apoptotic cells (%) were calculated in the presence of 50
μM of Nec-1, and necroptotic cells (%) were calculated
using the equation: [100% − (apoptotic cells + viable
cells)] = necroptotic cells (in the absence of Nec-1)
(Fig. S4).
Western Blot analysis of siRNA-induced knockdowns
of caspase-8 and caspase-10
HT29 cells were seeded on 12-well cell culture plate
(200,000 per well) and allowed to attach overnight. The
next day caspase-8 and caspase-10 siRNA lyophilizate was
dissolved in nuclease-free water to a final concentration of
26.7 μM, combined with Lipofectamine® 2000 Reagent in
Optimem medium and incubated for 30 min. Next, the
mixture was dissolved in McCoy’s medium (10%FBS, 5
mM glutamine, 5 mM penicillin/streptavidin) to a final
concentration of 100 nM siRNA and 2 ug/mL lipofecta-
mine. 1 mL of siRNA/lipofectamine solution was added to
three wells and the medium to three control wells. Control
and siRNA-treated cells were harvested after 24, 48, and 72
h of incubation, SDS-PAGE electroporated and subjected to
Western blot analysis. Membranes were incubated over-
night with anti-caspase-8 mouse serum (1:250) at 4°C or
mouse anti-caspase 10 antibody (1:500, MBL; cat. no
M059-3). Primary antibodies were detected with secondary
goat anti-mouse antibody (30 min, RT, 1:10,000, Alexa
Fluor 680) and membranes were scanned using Azure
system (658 nm channel). The siRNA-induced knockdowns
were analyzed using Image Studio Lite software (Lincoln,
NE, USA) and the protein depletion was calculated based
on the fluorescence intensity of detected bands.
Caspases activation in TRAIL/CHX-treated HT29 cells
500,000 cells per well were seeded on 12-well plates and
allowed to attach overnight. Cells were treated with TRAIL
(100 ng/mL) and CHX (5 μg/mL) for various timepoints,
harvested and resuspended in 100 μL of 1xSDS/DTT. After
sample denaturation at 95°C cells were sonicated and sub-
jected to western blot analysis in the same manner as
described above (SDS-PAGE: 30 min at 200 V; transfer to
nitrocellulose membrane: 60 min at 10 V; blocking-2%
BSA in TBS-T, 60 min at room temperature). Membranes
were separately incubated overnight with anti-caspase-8
mouse serum (1:250), anti-caspase-3 (1:1000, cat no. 9662,
Cell Signaling) or mouse anti-caspase 10 antibody (1:500,
MBL; cat. no M059-3) at 4°C, incubated with fluorescently
labeled secondary antibody (30 min, RT, 1:10,000, goat
anti-mouse IgG, AlexaFluor 680), followed by scanning in
Azure system (Sapphire Biomolecular Imager). The data
were analyzed using Azure Spot Analysis software.
Apoptosis and necroptosis assays using caspase-8
and caspase-10 knockdown cells
HT29 cells were seeded on two 96-well plates (10,000 cells/
well) and subjected to siRNA-induced knockdown of
caspase-8 or caspase-10. Lipofectamine® 2000 Reagent was
incubated with caspase-8 or caspase-10 siRNA in Optimem
medium for 30 min, then diluted in McCoy’s medium
supplemented with 10%FBS, 5 mM penicillin/streptavidin
and 5 mM glutamine to final concentration of 100 nM
siRNA and 2 ug/mL lipofectamine. Next, 100 μL of pre-
pared solution was added to each well and plates were
incubated for 72 h at 37°C (5% CO2). After this time,
siRNA solution was aspirated, and cells were preincubated
for 2 h with various inhibitors in two different concentra-
tions (5 and 25 μM). Next, extracellular death stimulant
TRAIL (100 ng/mL) and protein synthesis inhibitor-CHX
(5μg/mL) were added to induce apoptosis. To rescue cells
from necroptosis, Nec-1 (50 μM) was added to each well.
Monitoring caspases (auto)cleavage in the presence
of zVAD-fmk and zVAD-Bzl inhibitors
500,000 cells per well were seeded on 12-well plates and
allowed to attach overnight. Next, 25 μM zVAD-fmk or

Exploring the prime site in caspases as a novel chemical strategy for understanding the mechanisms of. . .
After 24 h, the HT-29 cell viability was determined using
MTS assay according to the manufacturer’s protocol
(CellTiter 96® Aqueous One Solution Cell Proliferation
Assay, G3582, Promega).
determining the mechanisms by which the caspases are
involved in apoptosis [33], therefore, we decided to use the
zVAD (also called Cbz-VAD) caspase recognition motif as
a scaffold for the addition of warheads that allow extension
into the prime region. To determine the significance of the
next building block position for compound activity, we used
iso- and tere-isomers of phthalic acid (Pht) in the synthesis,
since these isomers allow extension from the meta and para
positions into prime region space. In the prime area of the
inhibitor backbone, we embedded one of twenty natural
amino acids to see whether they would enhance the potency
and specificity of the inhibitor toward individual caspases.
Next, caspase broad-spectrum peptide (zVAD) was trans-
formed into reactive bromomethyl ketone (zVAD-BMK)
which was subsequently coupled to HO-iso/tere-Pht-X-resin
moiety (X stands for amino acid) on solid support, followed
by HPLC purification (Fig. 1c). This solid-phase/solution
phase parallel block chemical synthetic strategy allowed us
to obtain a large number of highly pure caspase inhibitors in
relatively short time (Fig. 1, Table S3).
Results and discussion
Inhibitor library design
We developed a comprehensive method for investigating
the prime region of the active site in caspases through direct
inhibitor library screening as this approach has not been
previously taken. The concept of the library design was
based on (1) the general caspase requirements for aspartic
acid in the P1 position, (2) the broad-spectrum peptide
sequence and (3) the electrophilic binding group that inhi-
bits caspase activity and can be used to explore the prime
region of the active site in caspases (Fig. 1a, b). zVAD-fmk
is the most widely used pan-caspase inhibitor for
Fig. 1 A general outline of the solid-phase synthesis of the caspase
inhibitor library. a The general architecture of caspase inhibitors for
the investigation of the prime site specificity. b We selected the zVAD
peptide motif as a general sequence recognized by all caspases and
combined it with either iso-/tere-phthalic acid to allow for the gen-
eration of a library with various amino acids to explore the prime site.
We used the same 20 amino acids in the iso-phthalic and tere-phthalic
isomers. c A general scheme of the solid-phase library synthesis.
Briefly, we transformed the z-VAD-OH peptide into a reactive bro-
momethyl ketone and coupled it to an electrophilic moiety (Pht) and
one of twenty amino acids that were previously attached to the resin.
Next, we cleaved all thirty-eight reaction products (19 iso-isomers and
19 tere-isomers) from the resin and purified them using HPLC

K. Groborz et al.
Determination of the kinetic parameters of the
inhibitors
structures of obtained compounds are quite rigid, prime
region of particular caspases has to be somehow adjusted to
be pro- iso or pro- tere. Furthermore, individual amino acids
situated in the prime area of the compound backbone were
able to distinguish between individual enzymes, even the
ones with strong, structural and functional similarities such
as caspase-8 and caspase-10. We were able to determine the
best and worst inhibitors for other caspases (Table S1, S2).
Table 1 shows the kinetic parameters (Ki) of selected
reversible inhibitors for the proinflammatory and apoptotic
caspases. In some cases, a change of only one amino acid in
the prime area allowed for significant increases or decreases
in the activity of the inhibitor for a particular enzyme, as
shown for caspases-4, -5, and -11. The analysis of apoptotic
caspases demonstrated that although the differences
between Ki values for the opposite inhibitors are smaller
than for proinflammatory caspases, we were still able to
obtain good inhibitory discrimination with our approach.
The best example here is caspase-8, for which the difference
between the best (isoPht-Trp) and the worst (terePht-Phe)
inhibitors was over 80-fold.
All the synthesized inhibitors containing the Pht binding
group demonstrated a rapid reversible mode of inhibition,
precluding the calculation of a kobs/I parameter, which is the
rate of enzymes inactivation via covalent modification of
their active site (Cys residue in caspases) by an irreversible
inhibitor (Fig. S1). This may be related to the fact that Pht
does not represent a good leaving group, because its
negative charge is not stabilized adequately. Therefore, to
test the activity and selectivity of the inhibitors for the
caspases, we calculated the equilibrium constant (Ki, tight
inhibition) across ten recombinant caspases (Fig. 2a). By
measuring Ki parameter we directly determined the strength
of the inhibitor-enzyme complex. (Fig. S1, Table S1, S2).
The most relevant differences between the examined cas-
pases are shown as a heat map (Fig. 2b). We used zVAD-
Bzl as a control compound and all the calculated values
were normalized to its Ki value (Fig. 2c). There were strong
differences between the recognition model of the particular
caspases, their preferences for the inhibitor isomers (iso- or
tere-) and the arrangement of the amino acid in the prime
region. For the proinflammatory caspases-4, -5, and -11 and
apoptotic caspases-9 and -10, inhibitors containing the tere-
isomer of phthalic acid were much more potent than the
ones with the iso-isomer. On the other hand, caspases-1
(both human and mice) and human caspases-3, -7, and -8
displayed general kinetic preferences for iso-isomers. As the
As apoptotic and proinflammatory caspases are involved in
different processes we correlated the global features in their
specificities to see whether we could extract a pattern showing
similarities between closely related enzymes, even if they
were from different organisms. In Fig. 3 we show the most
interesting outcomes of the Ki correlation for proinflammatory
caspases, which depicts the strong similarities in their prime
site architecture. Moreover, by using a simple functional/
Fig. 2 The inhibition parameters
(Ki) of the individual
compounds tested on individual,
recombinant caspases. a
Fluorogenic substrates (Ac-
peptide-ACC) used for the Ki
calculations. b The inhibition
parameters (Ki) of all 39
inhibitors presented as a heat
map where red indicates strong
inhibitor binding (low Ki) and
blue (and green) indicates weak
inhibitor binding (high Ki). For
better presentation, all kinetic
values (Ki) across the individual
caspases were presented as fold-
change comparing to zVAD-bzl
(value = 1, marked gray). c The
Ki values of zVAD-Bzl for
individual caspases

Exploring the prime site in caspases as a novel chemical strategy for understanding the mechanisms of. . .
Table 1 The kinetic parameters
Proinflammatory caspases
(Ki) of the broad-spectrum
reversible inhibitor of caspases,
zVAD-Bzl, contrasted with the
most and least potent inhibitors
from the iso-and tere-Pht-X
library, determined for
Ki, nM/enzyme hCasp-1
hCasp-4
hCasp-5
mCasp-1
mCasp-11
zVAD-Bzl 3.71
24.8
14.7
0.623
57.9
Best inhibitor isoPht-Trp 0.79 terePht-Trp 2.60 isoPht-Trp 2.3 terePht-Asn 0.178 terePht-Tyr 4.81
Worst inhibitor isoPht-Arg 12.0 isoPht-Arg 139 isoPht-Arg 116 isoPht-Arg 4.37
isoPht-Arg 403
individual caspase
Apoptotic caspases
Ki, nM/enzyme hCasp-3
hCasp-7
hCasp-8
hCasp-9
hCasp-10
zVAD-Bzl
1110
740
4.69
10.2
17.1
Best inhibitor
isoPht-Phe 295 isoPht-Phe 233 isoPht-Trp 1.83 terePht-Ser 1.13 terePht-Tyr 1.60
Worst inhibitor isoPht-Arg 3700 isoPht-Glu 1780 terePht-Phe 149 isoPht-Arg 29.4 isoPht-Gly 77.8
The Ki values for each inhibitor are displayed in bold font
Fig. 3 The correlation of the Ki parameters of the zVAD iso- and tere-
Pht-X inhibitors among select caspases. a A matrix of Ki correlations
for the proinflammatory caspases (left) and the detailed correlation
plots between human caspase-4 and mouse caspase-11 (right). These
data demonstrate that there is a significant similarity in the prime site
architecture of the proinflammatory caspases. b A matrix of the Ki
correlation for human apoptotic caspases (left) and the detailed
correlation plots between human caspase-8 and human caspase-9.
These data demonstrate that the prime site architecture differs across
the caspases. For example, there is a strong linear correlation between
the tere-phthalic inhibitors for caspase-8 and -9 but no correlation for
the iso-phthalic inhibitors. For each caspase pair, the Pearson corre-
lation coefficient was calculated and the results were presented either
as heat maps or scatter plots
activity approach, we demonstrated that human caspase-4 and
caspase-5 are equivalent to mouse caspase-11, as the Pearson
correlation coefficients for both iso- and tere-isomers were
very high, 0.962 and 0.780 respectively (for caspase-4/-11
pair), and 0.829 and 0.833 (for caspase-5/-11 pair). This
observation is in the line with a genetic evidence that in
humans caspases-4 and -5 are both orthologues to murine
caspase-11 (Fig. 3a) [34, 35]. Subsequently, we also made
this analysis for five human apoptotic caspases, demonstrating
that in general all these enzymes display much less conserved
prime region architecture than inflammatory caspases
(Fig. 3b). As an example, we investigated caspase-8 and

K. Groborz et al.
Table 2 The kinetic parameters
(Ki) of the most selective
inhibitors for the human
Apoptotic caspases
Target enzyme
Inhibitor, Ki nM
Caspase-3
Caspase-8
Caspase-9
Caspase-10
apoptotic caspases (-8, -9, -10,
upper panel) and the human
proinflammatory caspases
(-1, -4, -5, lower panel)
Caspase-8
Caspase-9
Caspase-10
zVAD-isoPht-Trp
zVAD-terePht-Ser
zVAD-terePht-Tyr
388
1.83
11.9
59.1
17.4
1.13
8.22
35.3
9.12
1.63
1315
1421
Proinflammatory caspases
Target enzyme
Inhibitor, Ki nM
Caspase-1
Caspase-4
Caspase-5
Caspase-1
Caspase-4
Caspase-5
zVAD-isoPht-Lys
zVAD-terePht-Leu
zVAD-terePht-Trp
2.82
8.43
5.21
60.4
4.64
8.44
65.6
9.32
3.21
The most selective inhibitor for each particular caspase is displayed in bold font
caspase-9 in details, as they are the key players in two dif-
ferent pathways in apoptosis. There was no correlation
between caspase-8/-9 with the iso-phthalic isomers of our
compounds (ρ = 0.024), thus these inhibitors could be used to
distinguish between the caspases by analyzing their prime
area preferences (Fig. 3b). The analysis of tere-phthalic iso-
mers showed that the inhibitor binding preferences of these
caspases at this orientation are alike. Interestingly, we
observed almost no correlation in inhibitors potency between
caspase-8 and caspase-10, which might suggest that although
these enzymes participate in the same pathway of apoptosis
initiation, their substrate preferences are different, and could
be possible to separate these proteases by selective inhibitors.
The correlation analysis for the other caspases showed that
our method can distinguish between closely-related pairs of
caspases (i.e., caspase-4 and caspase-5; caspase-8 and cas-
pase-10; and caspase-3 and caspase -7) (Figs. S1, S2).
Table 2 summarizes the kinetic parameters of the most
selective inhibitors of the apoptotic and proinflammatory
caspases. The presented values are the exact Ki values for
each enzyme and the Ki compared with that of human
caspase-3. The highest kinetic discrimination within apop-
totic caspases we obtained was for caspase-8 (zVAD-
isoPht-Trp), however for caspase-9 and -10 we also were
able to develop selective inhibitors (zVAD-terePht-Ser and
zVAD-terePht-Tyr). For the inflammatory caspases the best
selectivity we obtained was for caspase-1 with zVAD-
isoPht-Lys inhibitor which was 21- and 23-fold more potent
over caspase-4 and -5 respectively. The most selective
caspase-4 and -5 inhibitors displayed only poor dis-
crimination between the enzymes. However, it must be
stressed here, that the selectivity for each caspase can be
significantly improved by applying the optimal peptide
sequence in the non-prime position. Here, we analyzed only
the caspases non-prime site, while the prime region was
occupied by zVAD sequence that is known to be selective
for caspase-1 and caspase-8. This explains why the absolute
selectivity factors for these two enzymes were the most
pronounced.
Determination of the inhibition mode of the
selected inhibitors
Since the ultimate goal of our studies was to apply caspase
inhibitors in cell-based assays, we decided to transform
selected zVAD-Pht-X compounds into irreversible inhi-
bitors, as the latter ones are generally more potent in cel-
lulo. To do so we replaced the Pht group with a widely
used acyloxymethyl ketone (AOMK; 2,6-dimethylter-
ephthalic acid). We synthesized irreversible inhibitors with
glycine, serine, and phenylalanine in the prime area to be
able to distinguish between closely-related caspases such
as caspase-8, caspase-9 and, caspase-10 and compared
them with the reversible analogs (Fig. 4a). We examined
the differences in the mechanism of action of Pht- and
AOMK-based compounds on recombinant caspases-8 and
-10, as both are involved in the extrinsic pathway of
apoptosis (Fig. S5). As we expected, the zVAD-Pht-X
inhibitors had reversible kinetics and the zVAD-AOMK-X
inhibitors had irreversible kinetics with recombinant
caspase-10 (Fig. 4b). Surprisingly, inhibitors with either
warhead (Pht or AOMK) demonstrated classic reversible
kinetics on recombinant caspase-8 within initial assay
timeframe and tended to form an irreversible complex after
prolonged incubation (Fig. 4b, Fig. S5). To explore this
phenomenon, we incubated 50 nM of recombinant
caspase-8 and caspase-10 with 1 μM of selected inhibitors
for 30 min to fully inhibit caspase activity. Next, we added
the pan-caspase irreversible biotin-LEHD-AOMK probe to
detect enzymatic activity using fluorescence streptavidin
(Fig. 4c). When caspases were inhibited with irreversible
inhibitor (for example zVAD-fmk) no enzymatic activity
was detected with biotin-labeled probe. However, when
enzyme was inhibited in reversible fashion, biotin-LEHD-

Exploring the prime site in caspases as a novel chemical strategy for understanding the mechanisms of. . .
Fig. 4 Dissecting the mechanisms of the inhibition of caspases by
zVAD-tere-Bzl-X and zVAD-iso-Bzl-X inhibitors. Panel a The
structures of zVAD inhibitors with tere-phthalic (Bzl) or 2,6-dime-
thylterephthalic (AOMK) warheads. Panel b The inhibition curves of
caspase-8 and caspase-10 by zVAD-Bzl and zVAD-AOMK inhibitors
indicated that only caspase-10 vs. zVAD-AOMK had classic irrever-
sible inhibitor kinetics. Panel c Reversible/irreversible inactivation of
caspases-8 and -10 using selected inhibitors followed by the use of
irreversible pan-caspase biotin-LEHD-AOMK probe. Panel d zVAD-
Bzl and zVAD-AOMK inhibitors were shown to display reversible
inhibition kinetics towards caspase-8 in the initial timeframes of the
assay. In brief, caspase-8 was either preincubated with inhibitors
(green line) or the inhibitor was added right before the assay (orange
line). Next, the caspase-8-inhibitor complex was diluted, Ac-LEHD-
AFC substrate was added to the reaction mixture and the fluorescence
was read immediately. The recovery of enzyme activity demonstrated
reversible inhibitor kinetics. The blue line serves as a control (enzyme
plus substrate, no inhibitor)
AOMK probe competed for the active site with such
inhibitor, which resulted in the covalent enzyme mod-
ification (Fig. 4c). With this experiment, we showed that
even compounds considered to be irreversible inhibitors of
caspase-8, such as zVAD-AOMK-Gly/Ser/Phe, turned out
to form reversible complexes under tested conditions.
However, on the other hand, non-substituted zVAD-Bzl
and zVAD-AOMK appeared to display bimodal inhibition.
To further explore this, we preincubated caspase-8 with
selected inhibitors for 60 min, allowing the enzyme-
inhibitor complex to mature, then diluted it, and mea-
sured the recovery of enzyme activity in the presence of
fluorogenic substrate (Ac-LEHD-ACC). The rate of sub-
strate cleavage was lower compared to the one where the
inhibitor and caspase-8 were mixed right before assay (no
pre-incubation step), indicating that some portion of the
enzyme was irreversibly inhibited (Fig. 4d). Caspase-10,
which has a strong structural and functional likeness to
caspase-8, was irreversibly inhibited by both fmk and
AOMK-based warheads and displayed reversible inhibi-
tion toward Pht-inhibitors (Fig. 4b, d).
Dissecting cell death pathways in human colon
cancer cells
To explore the utility of our inhibitors we dissected the roles
of caspase-8 and caspase-10, which are involved in the
interplay of apoptosis and necroptosis (Fig. 5a). If the
activity of caspase-8 is blocked or disrupted, the cell trig-
gers necroptosis, which can be blocked by necrostatin-1
(Nec-1), an inhibitor of RIPK-1, [36]. Another important
protein is FLIPL which forms a heterodimer with caspase-8
that blocks the necroptosis pathway [37, 38]. Therefore, we
assembled a catalytically active caspase-8:FLIPL complex
(Fig. S6) and found that although the Ki values were higher
than the ones for caspase-8 alone, the specificity remained
unchanged (Fig. 5d, e). Next, we checked whether the
outcomes of apoptosis and necroptosis (cell viability)

K. Groborz et al.
Fig. 5 Necroptosis in HT29 cells was mediated by caspase-8 and not
by caspase-10. a The extrinsic pathway of apoptosis leads to the
activation of the death-inducing signaling complex (DISC) containing
active caspase-8 and caspase-10, which further leads to the activation
of downstream caspase-3 and caspase-7. When caspase-8 activity is
blocked, an alternative pathway is triggered, leading to a type of cell
death called necroptosis. Cells can be rescued from necroptosis when
treated with necrostatin-1 (Nec-1), a potent RIPK inhibitor. b Time-
course of the activation of caspases-8, -10, and -3 in TRAIL/CHX-
stimulated HT29 cells via Western blotting analysis. c The inhibition
of caspase-8 activation in apoptotic HT29 cells using irreversible
zVAD-fmk and reversible zVAD-Bzl inhibitors. zVAD-fmk was
shown to completely prevent caspase-8 activation, while zVAD-Bzl
only partially attenuates its auto-cleavage (top panel). Nevertheless,
both inhibitors were able to block apoptosis, and induce necroptosis
(bottom panel) Panel d and e. HT-29 cells were preincubated with
selected inhibitors, followed by induction of apoptosis (TRAIL
+CHX) or necroptosis (TRAIL+CHX+inhibitors). To inhibit
necroptosis, Nec-1 inhibitor was used. zVAD-fmk, a potent, broad-
spectrum caspase inhibitor rescued cells from apoptosis but not
necroptosis. When the cells were pretreated with both zVAD-fmk and
Nec-1, both processes were inhibited. The use of selective inhibitors
for caspase-8 and caspase-8:FLIPL (zVAD-Bzl-X) or caspase-10
(zVAD-AOMK-X) demonstrated that caspase-8 but not caspase-10
was involved in necroptosis in HT-29 cells. zVAD-AOMK-Phe, a
potent and selective caspase-10 inhibitor had no effect on either
apoptosis or necroptosis, while zVAD-Bzl (a selective and potent
caspase-8 inhibitor) partially rescued HT29 cells from apoptosis and
induced necroptosis
correlated with the activity of caspase-8, caspase-10 or both
enzymes. For example, a recent study reported that caspase-
10 can actually mediate cell death in HT-29 cells, but only if
caspase-8 is ablated [8]. To investigate this, we first per-
formed western blotting analysis, to confirm the presence
and the activation of caspase-8 and caspase-10 in response
to TRAIL + cycloheximide (CHX) stimulation at various
timepoints (Fig. 5b). The data showed that indeed, both
caspases are processed, leading to caspase-3 activation,
which in turn triggered cell death. Since caspase-8, when
inhibited, promotes necroptotic cell death, we also dissected
the influence of the most potent caspase-8 inhibitors on the
TRAIL/CHX-induced cell death (Fig. 5c). The results
demonstrated that zVAD-fmk fully inhibits caspase-8 acti-
vation, even after prolonged incubation, while reversible
autoprocessing. To test the influence of various inhibitors
on the outcomes of cell death stimulation, HT-29 cell were
preincubated with the inhibitors; TRAIL + cycloheximide
(CHX) was added to induce cell death (apoptosis and
necroptosis) and TRAIL + CHX + Nec-1 was added to
induce apoptosis. The potent broad-spectrum irreversible
caspase inhibitor (zVAD-fmk) blocked apoptosis and
induced necroptosis in the HT29 cell line (Fig. 5d, e).
However, cells pretreated with zVAD-fmk and Nec-1 sur-
vived, as both pathways were inhibited (Fig. S4). Next, we
pretreated HT-29 cells with selective inhibitors for caspase-
8 (Pht warhead) and caspase-10 (AOMK warhead) and
induced either apoptosis or necroptosis. There was a good
correlation between the potency of caspase-8 inhibitors and
the inhibition of apoptosis (and the promotion of necrop-
tosis) (Fig. 5d). The most potent caspase-8 inhibitor
zVAD-Bzl
only
partially
attenuates
caspase-8

Exploring the prime site in caspases as a novel chemical strategy for understanding the mechanisms of. . .
(zVAD-Bzl, Ki = 2.55 nM) switched HT-29 cells from
apoptosis to necroptosis, whereas the least potent caspase-8
inhibitor (zVAD-Bzl-Phe, Ki = 137 nM) did not. Similar
results were obtained for other caspase-8 selective com-
pounds (-Bzl-Gly and -Bzl-Ser), which were also able to
induce programmed necrosis in these cells. On the other
hand, the potent and irreversible caspase-10 inhibitor
Western blotting analysis revealed that indeed, transfecting
cells with caspase-8 or caspase-10 siRNA resulted in a
significant decrease of the expression of these enzymes,
however, we could not completely deplete caspase-8
(Fig. 6b, c). In the first step, caspase-8 or caspase-10
genes were silenced by administering siRNA/lipofectamine
solution for 72 h prior to experiment. Afterward, wild type
and knockdown cells were preincubated with the panel of
inhibitors and stimulated with TRAIL/CHX to induce cell
death. The data demonstrated that caspase-8 depletion
resulted in the significant increase of cell survival when
compared to wild type and caspase-10 depleted cells,
highlighting its prominent role in TRAIL/CHX-mediated
cell death (Fig. 6d). In a follow-up experiment, all HT29
cell variants (WT, Casp-8 siRNA and Casp-10 siRNA) were
preincubated with selected zVAD inhibitors containing
various binding groups (-fmk, -bzl, and -aomk) and +/−
necrostatin-1, followed by TRAIL/CHX treatment. The
differential use of Nec-1 allowed to directly measure the
level of necroptosis and apoptosis in HT29 cells. The data
show that in caspase-8 depleted cells, the necroptosis is
significantly reduced, whereas depletion of caspase-10 had
no effect on cell death, neither apoptosis, nor necroptosis
(zVAD-AOMK-Phe,
k ) did not
obs/I = 135,000 M−1 s−1
affect any of the cell death pathways, demonstrating that the
inhibition of caspase-10 is not relevant to any of the types of
cell death in the wild-type HT-29 cell line (Fig. 5e). To
further demonstrate the usability of our inhibitors to study
cell death mechanisms, we induced the apoptosis in HT-29
using TNF-α/birinapant stimulants, and triggered the
necroptosis switch by pre-incubation the cells with selected
inhibitors. By doing this we demonstrated that in this sys-
tem caspase-8, but not caspase-10 inhibition is the main
driver to induce programmed necrosis (Fig. S4).
Next, we aimed to additionally challenge our conclusions
that caspase-8, but not caspase-10 is mainly responsible for
necroptosis in HT29 cells. To do so, we performed caspase-
8 and caspase-10 siRNA-mediated knockdowns in HT29
cells, followed by the induction of cell death (Fig. 6a).
Fig. 6 Dissecting the role of caspases in necroptosis using wild type
and caspase-8/caspase-10 depleted HT29 cells. a The schematic
representation of three cell models to study necroptosis: wild type,
caspase-8 depleted and caspase-10 depleted cells. b, c Time-course of
the depletion of caspase-8 and caspase-10 in HT29 cells using siR-
NAs. The efficiency of caspase knockdowns was calculated based on
the fluorescence intensity of detected bands. d The effect of caspase-8/
caspase-10 depletion on TRAIL/CHX-induced cell death. Wild type
cells and caspase siRNA variants were preincubated with panel of
caspase inhibitors and stimulated with TRAIL/CHX for 24 h. Cells
with reduced caspase-8 expression were less susceptible for cell death
(apoptosis+necroptosis), while cells lacking caspase-10 displayed
similar “death pattern” as wild type HT29. The “−” in the x-axis
indicates that inhibitor is lacking amino acid in the prime position. e
Necroptotic pattern in three HT29 variants. All cell types were pre-
incubated with inhibitors and with/without Nec-1 followed by TRAIL/
CHX treatment. Next, the level of apoptosis and necroptosis were
calculated. Results showed that caspase-8 siRNA cells displayed sig-
nificantly lower necroptosis comparing to both, wild type and caspase-
10 siRNA cells

K. Groborz et al.
(Fig. 6e). Moreover, the reduced necroptosis in casp-8
siRNA cells is in line with the model where inhibited
caspase-8 is required for necrosome assembly [39–41].
caspase-10 from HT-29 cells did not affect necroptosis,
whereas silencing of caspase-8 resulted in increased cell
survival. We postulate that our methodology may be used to
determine which caspases are involved in other types of cell
death, i.e., pyroptosis, as we have already demonstrated that
by exploring the prime site specificity, a significant inhibitory
potency and selectivity can be achieved.
Conclusions
The development of selective tools that enable the distinc-
tion between particular caspases is of great importance since
they are of exceptional use in biochemical and biological
studies. Although there have been many attempts to find
selective compounds that can be used to suppress individual
caspases and reveal their involvement in various pro-
grammed cell death pathways, none have been sufficient
enough to distinguish between closely related cysteine
proteases that exhibit overlapping substrate specificity. To
address this issue we used a novel, chemical strategy to
examine the prime region in caspases, as its analysis is
informative. The advantage of our approach is the ease, the
speed and yield of the solid-state synthesis of bifunctional,
prime site-exploring compounds. Using our inhibitors we
showed the differences and similarities in the binding
strengths of particular caspases (expressed as Ki values) and
proved the importance of the prime region in their active
sites. Our detailed kinetic analysis demonstrated that
although all these inhibitor contain the same peptide motif
(zVAD) motif in the non-prime region, they can be fairly
distinguished using only one amino acid in the prime region
of active site groove. In addition, the selectivity toward
particular caspase can be largely improved by applying its
optimal non-prime peptide sequence.
Acknowledgements This project was funded by the European Union’s
Horizon 2020 research and innovation program under the Marie
Skłodowska-Curie grant agreement No. 661187 (to MP) and by
National Science Centre in Poland (grant OPUS, UMO-2018/29/B/
NZ102249 to MP). The Salvesen laboratory is supported by Sanford
Burnham Prebys NCI Cancer Center Support Grant P30CA030199
and NIH grant (GM99040). The Drag laboratory is supported by the
Foundation for Polish Science and National Science Centre in Poland.
Author contributions MP and MD conceived the project, KG, GS,
MD, and MP designed and KG and MP performed the experiments,
MLG and SJS contributed new reagents and tools, KG, GS, MD, and
MP drafted and edited the paper. All authors reviewed the results and
approved the final version of the paper.
Compliance with ethical standards
Conflict of interest The authors declare that they have no conflict of
interest.
Publisher’s note: Springer Nature remains neutral with regard to
jurisdictional claims in published maps and institutional affiliations.
References
1. Grutter MG. Caspases: key players in programmed cell death.
Curr Opin Struct Biol. 2000;10:649–55.
In order to test our compounds in cell-based experiments,
we transformed our reversible inhibitors into irreversible
AOMK-based analogs, since these compounds are more
potent in in cellulo studies. Surprisingly, we discovered that
these compounds exhibited only reversible inhibition kinetics
for caspase-8 and irreversible inhibition kinetics for the other
caspases, i.e., caspase-3, caspase-7, and caspase-10. We then
we used selected Bzl and AOMK inhibitors to dissect cell
death mechanisms in the HT-29 human colon cancer cell line.
Necroptosis is a type of cell death that can be initiated
exclusively by treatment with tumor necrosis factor (TNF) in
the presence of caspase inhibitors such as zVAD-fmk, which
inhibits caspase-8 function [20, 42]. Using small inhibitors,
we demonstrated that necroptosis in HT-29 human colon
cancer cells was mediated by caspase-8 (and caspase-8:
FLIPL), as its selective inhibition blocked apoptosis and
promoted necroptosis. The inhibition of caspase-10 did not
affect cell viability and we concluded that it had no measur-
able impact on apoptosis or necroptosis. In addition, we
performed caspase-8 and caspase-10 knockdown experiments
to further confirm the negligible role of caspase-10 in TRAIL/
CHX/inhibitors-mediated cell death. Indeed, depletion of
2. Rawlings ND, Waller M, Barrett AJ, Bateman A. MEROPS: the
database of proteolytic enzymes, their substrates and inhibitors.
Nucleic acids Res. 2014;42(Database issue):D503–9.
3. Sun L, Wang H, Wang Z, He S, Chen S, Liao D, et al. Mixed
lineage kinase domain-like protein mediates necrosis signaling
downstream of RIP3 kinase. Cell. 2012;148:213–27.
4. Vercammen D, Beyaert R, Denecker G, Goossens V, Van Loo G,
Declercq W, et al. Inhibition of caspases increases the sensitivity
of L929 cells to necrosis mediated by tumor necrosis factor. J Exp
Med. 1998;187:1477–85.
5. Silke J, Strasser A. The FLIP Side of Life. Sci Signal. 2013;6:pe2.
6. Silke J, Meier P. Inhibitor of apoptosis (IAP) proteins-modulators
of cell death and inflammation. Cold Spring Harb Perspect Biol
2013;5: pii: a008730.
7. Lork M, Verhelst K, Beyaert R. CYLD, A20 and OTULIN deu-
biquitinases in NF-kappaB signaling and cell death: so similar, yet
so different. Cell Death Differ. 2017;24:1172–83.
8. Tanzer MC, Khan N, Rickard JA, Etemadi N, Lalaoui N, Spall
SK, et al. Combination of IAP antagonist and IFNgamma activates
novel caspase-10- and RIPK1-dependent cell death pathways. Cell
Death Differ. 2017;24:481–91.
9. Berger AB, Sexton KB, Bogyo M. Commonly used caspase
inhibitors designed based on substrate specificity profiles lack
selectivity. Cell Res. 2006;16:961–3.
10. Benkova B, Lozanov V, Ivanov IP, Mitev V. Evaluation of
recombinant caspase specificity by competitive substrates. Anal
Biochem. 2009;394:68–74.

Exploring the prime site in caspases as a novel chemical strategy for understanding the mechanisms of. . .
11. Pereira NA, Song Z. Some commonly used caspase substrates and
inhibitors lack the specificity required to monitor individual cas-
pase activity. Biochem Biophys Res Commun. 2008;377:873–7.
12. McStay GP, Salvesen GS, Green DR. Overlapping cleavage motif
selectivity of caspases: implications for analysis of apoptotic
pathways. Cell Death Differ. 2008;15:322–31.
27. Copeland RA. Enzymes: a practical introduction to structure,
mechanism, and data analysis. 2nd edn. New York: Wiley;
2000.
28. Boatright KM, Deis C, Denault JB, Sutherlin DP, Salvesen GS.
Activation of caspases-8 and -10 by FLIP(L). Biochem J.
2004;382(Pt 2):651–7.
13. Poreba M, Szalek A, Kasperkiewicz P, Rut W, Salvesen GS, Drag
M. Small molecule active site directed tools for studying human
caspases. Chem Rev. 2015;115:12546–629.
29. Sosna J, Philipp S, Fuchslocher Chico J, Saggau C, Fritsch J, Foll
A, et al. Differences and similarities in TRAIL- and tumor
necrosis factor-mediated necroptotic signaling in cancer cells. Mol
Cell Biol. 2016;36:2626–44.
30. Voigt S, Philipp S, Davarnia P, Winoto-Morbach S, Roder C,
Arenz C, et al. TRAIL-induced programmed necrosis as a novel
approach to eliminate tumor cells. BMC Cancer. 2014;14:74.
31. Fauster A, Rebsamen M, Huber KV, Bigenzahn JW, Stukalov A,
Lardeau CH, et al. A cellular screen identifies ponatinib and pazo-
panib as inhibitors of necroptosis. Cell Death Dis. 2015;6:e1767.
32. Vince JE, Wong WW, Khan N, Feltham R, Chau D, Ahmed AU,
et al. IAP antagonists target cIAP1 to induce TNFalpha-dependent
apoptosis. Cell. 2007;131:682–93.
14. Schechter I, Berger A. On the size of the active site in proteases. I.
Papain. Biochem Biophys Res Commun. 1967;27:157–62.
15. Poreba M, Szalek A, Rut W, Kasperkiewicz P, Rutkowska-
Wlodarczyk I, Snipas SJ, et al. Highly sensitive and adaptable
fluorescence-quenched pair discloses the substrate specificity
profiles in diverse protease families. Sci Rep. 2017;7:43135.
16. Powers JC, Asgian JL, Ekici OD, James KE. Irreversible inhibi-
tors of serine, cysteine, and threonine proteases. Chem Rev.
2002;102:4639–750.
17. Ganesan R, Jelakovic S, Campbell AJ, Li ZZ, Asgian JL, Powers
JC, et al. Exploring the S4 and S1 prime subsite specificities in
caspase-3 with aza-peptide epoxide inhibitors. Biochemistry.
2006;45:9059–67.
33. Van Noorden CJF. The history of Z-VAD-FMK, a tool for
understanding the significance of caspase inhibition. Acta His-
tochem. 2001;103:241–51.
18. Elmore S. Apoptosis: a review of programmed cell death. Toxicol
Pathol. 2007;35:495–516.
34. Kamens J, Paskind M, Hugunin M, Talanian RV, Allen H,
Banach D, et al. Identification and characterization of ICH-2, a
novel member of the interleukin-1 beta-converting enzyme family
of cysteine proteases. J Biol Chem. 1995;270:15250–6.
35. Munday NA, Vaillancourt JP, Ali A, Casano FJ, Miller DK,
Molineaux SM, et al. Molecular cloning and pro-apoptotic activity
of ICErelII and ICErelIII, members of the ICE/CED-3 family of
cysteine proteases. J Biol Chem. 1995;270:15870–6.
36. Declercq W, Vanden Berghe T, Vandenabeele P. RIP kinases at
the crossroads of cell death and survival. Cell. 2009;138:229–32.
37. Pop C, Oberst A, Drag M, Van Raam BJ, Riedl SJ, Green DR,
et al. FLIP(L) induces caspase 8 activity in the absence of inter-
domain caspase 8 cleavage and alters substrate specificity. Bio-
chem J. 2011;433:447–57.
38. Oberst A, Dillon CP, Weinlich R, McCormick LL, Fitzgerald P,
Pop C, et al. Catalytic activity of the caspase-8-FLIP(L) complex
inhibits RIPK3-dependent necrosis. Nature. 2011;471:363–7.
39. Moriwaki K, Chan FK. RIP3: a molecular switch for necrosis and
inflammation. Genes Dev. 2013;27:1640–9.
40. Vandenabeele P, Declercq W, Van Herreweghe F, Vanden Berghe
T. The role of the kinases RIP1 and RIP3 in TNF-induced
necrosis. Sci Signal. 2010;3:re4.
19. McStay GP, Green DR. Measuring apoptosis: caspase inhibitors
and activity assays. Cold Spring Harb Protoc. 2014;2014:799–806.
20. Riedl SJ, Shi Y. Molecular mechanisms of caspase regulation
during apoptosis. Nat Rev Mol cell Biol. 2004;5:897–907.
21. Weinlich R, Oberst A, Beere HM, Green DR. Necroptosis in
development, inflammation and disease. Nat Rev Mol Cell Biol.
2017;18:127–36.
22. Kato D, Boatright KM, Berger AB, Nazif T, Blum G, Ryan C,
et al. Activity-based probes that target diverse cysteine protease
families. Nat Chem Biol. 2005;1:33–8.
23. Poreba M, Kasperkiewicz P, Snipas SJ, Fasci D, Salvesen GS,
Drag M. Unnatural amino acids increase sensitivity and provide
for the design of highly selective caspase substrates. Cell Death
Differ. 2014;21:1482–92.
24. Maly DJ, Leonetti F, Backes BJ, Dauber DS, Harris JL, Craik CS,
et al. Expedient solid-phase synthesis of fluorogenic protease
substrates using the 7-amino-4-carbamoylmethylcoumarin (ACC)
fluorophore. J Org Chem. 2002;67:910–5.
25. Stennicke HR, Salvesen GS. Caspases: preparation and char-
acterization. Methods. 1999;17:313–9.
26. Gonzalez ML, Poreba M, Snipas SJ, Groborz K, Drag M, Sal-
vesen GS. Extensive peptide and natural protein substrate screens
reveal that mouse caspase-11 has much narrower substrate spe-
cificity than caspase-1. J Biol Chem. 2018;293:7058–67.
41. Tummers B, Green DR. Caspase-8: regulating life and death.
Immunol Rev. 2017;277:76–89.
42. Zhou W, Yuan J. Necroptosis in health and diseases. Semin Cell
Dev Biol. 2014;35:14–23.