Critical Reviews in Biochemistry and Molecular Biology
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The molecular era of protein S-acylation: spotlight
on structure, mechanisms, and dynamics
María-Eugenia Zaballa & F. Gisou van der Goot
To cite this article: María-Eugenia Zaballa & F. Gisou van der Goot (2018) The molecular era
of protein S-acylation: spotlight on structure, mechanisms, and dynamics, Critical Reviews in
Biochemistry and Molecular Biology, 53:4, 420-451, DOI: 10.1080/10409238.2018.1488804
To link to this article: https://doi.org/10.1080/10409238.2018.1488804
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CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
2018, VOL. 53, NO. 4, 420–451
https://doi.org/10.1080/10409238.2018.1488804
REVIEW ARTICLE
The molecular era of protein S-acylation: spotlight on structure, mechanisms,
and dynamics
Marıa-Eugenia Zaballa
and F. Gisou van der Goot
Global Health Institute, School of Life Sciences, Ecole Polytechnique Federale de Lausanne, Lausanne, Switzerland
ABSTRACT
ARTICLE HISTORY
S-Acylation (commonly referred to as S-palmitoylation) is a post-translational modification consisting in the covalent attachment of an acyl chain to a cysteine residue of the target protein.
The lability of the resulting thioester bond gives S-acylation an essential characteristic: its reversibility. S-acylation dynamically regulates different aspects in the life of a protein (including stability, localization, interactome, and function) and, thus, plays critical roles in cellular physiology.
For long, the reversibility of S-acylation has been neglected and thereby its potential as a regulatory mechanism for protein function undervalued. Thanks to technological advances, the field
has now entered its golden era. A great diversity of interesting targets is being identified, the
physio-pathological importance of the modification is starting to be revealed, structural information on the enzymes is becoming available, and the regulatory dynamics are gradually being
understood. Here we will review the most recent literature in the S-acylation field, with a special
focus on the molecular aspects of the modification, its regulation, and its consequences.
Received 14 May 2018
Revised 11 June 2018
Accepted 11 June 2018
Introduction
Protein function can be modulated by a plethora of
post-translational modifications. Some of these modify
physical–chemical properties, such as the addition of
charges by phosphorylation or increasing the hydrophobicity by the addition of lipids. This review focuses
on S-acylation, a modification conserved among eukaryotes that consists in the covalent attachment of a long
acyl chain (typically but not exclusively palmitoyl-CoA)
to specific cysteine residues on the target protein.
While most other protein long-chain lipidations (e.g.
N-terminal myristoylation, prenylation, and O-acylation)
result in stable bond formation, S-acylation is reversible,
allowing dynamic fine-tuning the properties of the
modified protein (Jiang et al. 2018). Chemically, the
thiol group of the target cysteine residue reacts with
the carbonyl carbon of the lipid moiety forming a
thioester bond, which is more labile than its oxyester
counterpart (Castro 1999). In a biological context, the
S-acylation reaction is catalyzed by a family of enzymes
named protein S-acyltransferases (PATs) whereas the
reverse reaction, deacylation, is catalyzed by the socalled acyl protein thioesterases (APTs) or deacylases
(Figure 1(A)) (Tsutsumi et al. 2008; Won et al. 2018).
KEYWORDS
Post-translational modification; S-acylation; dynamics;
protein S-acyltransferases
(PATs); thioesterases;
substrate recognition
Recently, long-chain N-acylation on lysine residues has
also been shown to be reversible (with members of the
sirtuin family catalyzing the deacylation reaction), and
thus to have the potential of dynamically regulate protein properties (Feldman et al. 2013; Bheda et al. 2016;
Jing et al. 2017). This review focuses exclusively on longchain cysteine S-acylation, commonly referred to as Spalmitoylation. Other types of protein lipidation are
reviewed elsewhere (Lin et al. 2012; Jiang et al. 2018).
Protein S-acylation was first reported in 1979
(Schmidt and Schlesinger 1979) and since then hundreds to thousands of protein substrates have been
identified in eukaryotes, including soluble and transmembrane proteins from most subcellular compartments (Blanc et al. 2015). S-Acylation of soluble
proteins confers increased membrane affinity and thus
controls protein localization and switches cellular
mobility of the protein from diffusion to vesicular trafficking mode. One of the best-studied examples is Sacylation of Ras proteins, which regulates their trafficking between the Golgi apparatus and the plasma membrane (Linder and Deschenes 2007). Transmembrane
proteins, despite their inherent membrane anchorage,
also frequently undergo S-acylation, which may initiate
CONTACT Marıa-Eugenia Zaballa
maria.zaballa@epfl.ch; F. Gisou van der Goot
Lausanne, EPFL SV GHI VDG, Station 19, CH-1015, Lausanne, Switzerland
gisou.vandergoot@epfl.ch
Ecole Polytechnique Fédérale de
ß 2018 The Author(s). Published by Informa UK Limited, trading as Taylor & Francis Group.
This is an Open Access article distributed under the terms of the Creative Commons Attribution-NonCommercial-NoDerivatives License (http://creativecommons.org/licenses/bync-nd/4.0/), which permits non-commercial re-use, distribution, and reproduction in any medium, provided the original work is properly cited, and is not altered, transformed,
or built upon in any way.
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
Figure 1. Protein S-acylation. (A) Scheme of the protein Sacylation reaction. The acyl moiety from an acyl-CoA is covalently bound to the thiol group of a cysteine residue through
a thioester bond. The direct reaction is catalyzed by DHHC Sacyltransferases and the reverse reaction is catalyzed by acyl
protein thioesterases. The attached acyl chain is typically
C16:0 but other chain length and saturation are possible (variable “n”). (B) Schematic representation of DHHC S-acyltransferases, highlighting their membrane topology and conserved
motifs (see text for details). While most of the members of
the family have four transmembrane domains (TMDs) as
shown here, up to six TMDs are found in some of them. (C)
Membrane topology for human zDHHC22 as predicted using
TMHMM v2.0 DTU Bioinformatics, Denmark. Similar results are
obtained using the predictor TMpred (ExPASy, SIB,
Switzerland) (not shown) (see color version of this figure at
www.tandfonline.com/ibmg).
conformational changes (e.g. tilting of transmembrane
domains), regulate recruitment of the protein to specific
membrane domains, promote association into protein
complexes, or protect from other post-translational
modifications such as ubiquitination thus preventing
degradation (Abrami et al. 2008; Blaskovic et al. 2013).
421
Membrane proteins undergoing S-acylation include
channels and pumps (e.g. BK channels and sodium/
potassium ATPase), many G protein-coupled receptors
(e.g. rhodopsin), chaperones (e.g. calnexin), plasma
membrane receptors (e.g. transferrin receptor, c-Met,
LRP6, TEM8, EGF receptor), proteins involved in synaptic
signaling (e.g. PSD-95, AMPA receptor), among others
(Chamberlain and Shipston 2015; Coleman et al. 2016;
Fukata et al. 2016; Collins et al. 2017; Howie et al. 2018).
For decades, the extent of protein S-acylation has
been underestimated in particular due to the technical
challenges associated with the identification of
enzymes and substrates. Moreover, the reversibility of
S-acylation has been neglected and thereby its potential as a regulatory mechanism for protein function
undervalued. Thanks to technological advances, the
field has now entered its golden era. A great diversity
of interesting targets is being identified, the physiopathological importance of the modification is starting
to be revealed, structural information on the enzymes is
becoming available, and the regulatory dynamics are
gradually being understood. Here we will review the
most recent literature in the S-acylation field. We will
first describe the available methods, their advantages
and drawbacks. Subsequently, we have chosen to focus
on the molecular aspects of both the modification and
its consequences, and the regulation of the modification. Excellent reviews covering earlier research and/or
focusing on specific aspects such as S-acylation in cancer or in brain function have been recently published
and thus these topics will only briefly be mentioned
here (Chamberlain and Shipston 2015; Cho and Park
2016; Fukata et al. 2016; Gottlieb and Linder 2017;
Lanyon-Hogg et al. 2017; De and Sadhukhan 2018).
Likewise, this review focuses on S-acylation in mammals. The reader is referred to other reviews focused on
S-acylation in plants (Hemsley 2009; Hurst and Hemsley
2015), parasites (Brown et al. 2017; Ritzefeld et al. 2017),
or fungi (Santiago-Tirado and Doering 2016).
Tools to study protein S-acylation
A variety of different methods to study S-acylation have
been developed, each addressing different aspects of
the modification. In this review, we provide a description of each method highlighting the main pitfalls, limitations, and common misinterpretations.
Radiolabeling
The oldest and probably most accepted method to
study protein S-acylation is metabolic labeling using
422
M.-E. ZABALLA AND F. G. VAN DER GOOT
radioactive palmitic acid (O’Brien and Zatz 1984;
Hancock 1995). It consists of incubating tissue culture
cells with radioactive palmitic acid in starvation
medium for a certain period that can be adjusted
according to the aims of each experiment. Both [3H]palmitic acid and [14C]-palmitic acid can be used but
the latter has remained limited given biosafety issues.
After labeling, cell lysis is followed by immunoprecipitation of the protein of interest (endogenous or ectopically
expressed), SDS-PAGE, and fluorography (Waterborg and
Matthews 1994). To confirm that the acyl chain is
bound to the protein through a thioester bond (i.e. that
the modification being considered is S-acylation and
not N- or O-acylation), a lane including a comparable
amount of protein sample treated with hydroxylamine
solution (to selectively hydrolyze the thioester bond
and thus lose the radioactive signal) needs to be
included in the same gel as a control. The time of
exposure by fluorography varies from a few days up to
months depending on the abundance of the protein,
and the efficiency of the labeling and immunoprecipitation. The required time of exposure and the use of
radioactive material are the most obvious disadvantages of this technique. It is important to realize that
metabolic labeling only reports on newly or dynamically S-acylated proteins. That is, long-lived, stably S-acylated proteins that do not exchange the endogenous
fatty acid by the exogenously added labeled one will
remain invisible to this technique. Also, when adding
radioactive palmitate (C16:0), S-palmitoylation will preferentially be monitored even though cells may metabolize the labeled fatty acid. No conclusion can be
drawn regarding other chain lengths, thus it cannot be
excluded that modification with other acyl chains (e.g.
C18:0 or C16:1) would be more efficient. Besides, metabolic labeling does not report on the fraction of the
total protein population that is S-acylated, nor does it
provide information on the number of acylation sites
and their occupancy (see below). The true power of this
method resides in its ability to study dynamics. Indeed
(i) it allows quantitative determination of the kinetics of
fatty acid incorporation and it is compatible with pulsechase experiments; and (ii) it allows quantitative comparisons of different conditions such as silencing or overexpression of S-acylation-related enzymes, cysteine point
mutations, or treatment with inhibitors or other drugs.
Metabolic labeling using bioorthogonal chemistry
A non-radioactive, faster alternative to radiolabeling is
based on the use of click-chemistry analogs of fatty
acids. The metabolic labeling is performed using azide-
or alkyne-modified fatty acids (e.g. 17-octadecynoic
acid) (Kostiuk et al. 2008; Charron et al. 2009; Hannoush
and Arenas-Ramirez 2009; Martin and Cravatt 2009).
Cell labeling and lysis can be directly followed by
immunoprecipitation of the protein of interest or by
the so-called click reaction (a copper-catalyzed 1,3dipolar cycloaddition). One important advantage of this
technique compared to radiolabeling is that the click
handle not only allows for detection of the protein of
interest (based on fluorescence or chemiluminescence)
but also for affinity pull-down of the cellular pool of
labeled proteins (usually based on a biotin–streptavidin
interaction) and global analysis of protein S-acylation
when combined with mass spectrometry (MS)-based
proteomics. Metabolic labeling with clickable fatty acids
has also been followed by fluorescent labeling on cells
and visualization by microscopy (Gao and Hannoush
2014). A detailed review on the use of click-chemistrybased probes for protein S-acylation has been recently
published (Gao and Hannoush 2018). While a recent
study has elegantly shown that fatty acid azides are
good mimics of endogenous fatty acids and can be efficiently taken up and metabolized by cells (Greaves
et al. 2017), this might not be true for all click-chemistry
analogs and all tested conditions, so conclusions should
be drawn with additional care.
Metabolic labeling using click-chemistry analogs, of
course, faces the same limitations as the above-described
radiolabeling approach. Therefore, it is highly recommended to combine these “lipid-centric” methods with
the so-called “cysteine-centric” (Chamberlain and Shipston
2015) or “hydroxylamine-switch” (Won et al. 2018) methods described below.
Acyl-biotin exchange (ABE) and acyl resin-assisted
capture (Acyl-RAC)
ABE and Acyl-RAC are two related, extensively used
methods to study protein S-acylation (Drisdel and
Green 2004; Forrester et al. 2011). These cysteine-centric methods provide a snapshot of the cellular pool of
S-acylated proteins. Briefly, these methods are based on
three main steps, once the protein sample is obtained:
(1) blocking of all free thiol groups, (2) selective cleavage of thioester bonds using a hydroxylamine solution
at neutral pH, and (3) capture of the proteins with
newly freed thiol groups either by biotinylation and
subsequent streptavidin-based capture (ABE) or directly
by thiopropyl beads-mediated pull-down (Acyl-RAC).
These methods allow for western blot-based detection
of the proteins of interest (notably more than one protein can be analyzed at the same time) or can be easily
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
coupled to MS-based proteomics. Together with bioorthogonal labeling, these methods have proved to be a
valuable source of information towards the global analysis of protein S-acylation, as described below (Blanc
et al. 2015; Sanders et al. 2015). Moreover, these methods can be applied to all kind of protein samples,
including tissues from animal models and other biological samples. Several caveats need, however, to be
highlighted. A main drawback of these methods is that
the capture of proteins is based on the selective cleavage of thioester bonds; thus, enzymes forming non Sacylation-related thioester bonds will be picked up as
false positives for S-acylation. These include ubiquitin
conjugases and activating enzymes, subunits of the
pyruvate dehydrogenase, fatty acid synthase, and other
metabolic enzymes utilizing phosphopantetheine prosthetic groups or acting on acyl-CoA substrates (Roth
et al. 2006; Kang et al. 2008). Additional false positives
can be detected due to incomplete blockade of free
thiol groups during the first step of the protocol. It is,
moreover, important to note that this method does not
discriminate a protein with one S-acylation site from
one with multiple S-acylation sites. Thus, for a protein
with several S-acylated residues, mutation of individual
cysteines (or downregulation of cysteine-selective
PATs) might not be translated into a reduced signal as
the remaining cysteines would still be available for pulldown. Besides, capture efficiency might be influenced
by a variety of factors, such as protein relative abundances and hydrophobicity. Therefore, these methods provide only semi-quantitative information. Finally,
quantification requires the parallel analysis of a known
amount of input sample. All in all, these thioester-based
methods are very powerful and allow for a relatively
easy and fast identification of S-acylation candidates in
a variety of samples but careful analysis of the results
should be performed to avoid overinterpretations when
intended to obtain quantitative information.
Monitoring multi-S-acylation using a PEG-induced
mass shift assay
None of the above methods provide information
regarding the number of S-acylation sites, nor the
abundance of the modified molecules. Therefore, the
Fuller group developed a PEG switch assay (Howie et al.
2014). This assay is a modification of the Acyl-RAC
method where protein samples are treated with 5 kDa
or 10 kDa PEG-maleimide during neutral hydroxylamine
incubation to allow for PEGylation of the freed cysteine
residues. The Hang and Fukata groups have proposed
modified versions of this method, which they called
423
acyl-PEG exchange (Percher et al. 2016) or APEGS
(Yokoi et al. 2016). In all these methods, for every
reacted cysteine, PEGylation leads to a mass shift that
can be observed by SDS-PAGE and western blotting. If
a protein has two S-acylation sites, this would allow the
detection of three bands, additionally providing some
information on the relative abundance of doubly, singly, and non-acylated species. Note that even the singly
modified species can be a mixed population (Dallavilla
et al. 2016). Also, PEGylation efficiency might be
affected by the distance between sites, and thereby
potential steric hindrance, and by labeling probabilities.
Therefore, PEGylation is particularly useful to estimate
the fraction of non-acylated protein.
Given the specific advantages and disadvantages of
the different methods available to monitor S-acylation
(Table 1), a combination of at least one lipid-centric and
one cysteine-centric methods is required to obtain a
reliable view of the S-acylation status of a given protein.
Global profiling of S-acylproteomes
The implementation of MS-based high-throughput
approaches to study protein S-acylation has significantly contributed to the development of the field by
providing S-acylation candidates for further follow-up
studies as well as statistics on the cellular processes or
compartments enriched in S-acylated proteins. To date,
a total of 40 S-acylproteomics studies on several species
(including unicellular eukaryotes, plants, and mammals)
have been published. From these, 17 studies are based
on lipid-centric approaches using click-chemistry
(Martin and Cravatt, 2009; Yount et al. 2010; Wilson
et al. 2011; Jones et al. 2012; Li et al. 2012; Martin et al.
2012; Zhang et al. 2013; Chesarino et al. 2014a;
Colquhoun et al. 2015; Foe et al. 2015; Morrison et al.
2015; Peng and Hang 2015; Santiago-Tirado et al. 2015;
ska
Serwa et al. 2015; Hernandez et al. 2016; Sobocin
et al. 2018; Thinon et al. 2016), whereas 23 studies
employ cysteine-centric approaches (mainly ABE or
Acyl-RAC) (Roth et al. 2006; Kang et al. 2008; Yang et al.
2010; Dowal et al. 2011; Emmer et al. 2011; Forrester
et al. 2011; Merrick et al. 2011; Ivaldi et al. 2012; Jones
et al. 2012; Marin et al. 2012; Hemsley 2013; Wan et al.
2013; Ren et al. 2013a; Wei et al. 2014; Gould et al.
2015; Morrison et al. 2015; Caballero et al. 2016; Fang
et al. 2016; Pinner et al. 2016; Upadhyay et al. 2016;
Collins et al. 2017; Edmonds et al. 2017; Shen et al.
2017). The comparative analysis of the available
S-acylproteomics studies represents a valuable source
of information and a powerful hypothesis-generating
engine. Particularly useful is the comparison of S-
ABE: acyl-biotin exchange; Acyl-RAC: acyl resin-assisted capture; IP: immunoprecipitation; Mass spec: mass spectrometry-based proteomics; NH2OH: hydroxylamine.
Non S-acylation-related thioester bonds; Incomplete PEGylation;
incomplete blockade of thiol groups
steric hindrance on
nearby sites
Simultaneous monitoring of multiple
S-acylated species; suitable for
all kind of samples
No
Western blot
Yes (þ)
No enrichment
PEG switch assay,
Acyl-PEG exchange,
APEGS
Potential false negatives
Suitable for all kind of samples; simple Non S-acylation-related thioester bonds; Incomplete pull-down;
incomplete blockade of thiol groups
low abundance proteins
Yes
Pull-down of biotin (ABE) or Western blot; Semi-quantitative
free SH groups (Acyl-RAC)
Mass spec
ABE & Acyl-RAC
Other types of lipidation if missing
NH2OH control;
unspecific fluorophore binding
Non-radioactive;
Time-dependent monitoring;
Reliable comparative quantification
Yes
Yes (þþ)
Fluorescence;
Western blot;
Mass spec
Bioorthogonal labeling IP of protein of interest;
(click chemistry analogs) pull down of click handle
Other types of lipidation if
missing NH2OH control
Potential false positives
Advantages
Time-dependent monitoring;
Reliable comparative quantification
No
Yes (þþ)
Fluorography
IP of protein of interest
Radiolabeling
([3H]-palmitic acid)
Suitable
for omics?
Quantitative?
Detection
method
Enrichment method
Method
S-acylation with acyl chains
other than C16:0; slow,
non-dynamic S-acylation
S-acylation with acyl chains
other than C16:0; slow,
non- dynamic S-acylation
M.-E. ZABALLA AND F. G. VAN DER GOOT
Table 1. Methods to study protein S-acylation. See text for more details.
424
acylation candidate datasets obtained using complementary approaches (lipid- and cysteine-centric), which
significantly increases the confidence of the analysis.
SwissPalm, a comprehensive, manually curated database has been developed to compile all available studies on protein S-acylation, both proteomics-based and
single protein studies, and facilitate comparisons (Blanc
et al. 2015). The second version of SwissPalm (www.
swisspalm.org) includes a total of 661 articles reporting
on the potential S-acylation of almost 10,000 proteins
from 55 organisms. Moreover, several user-friendly tools
allow for the direct comparison of an input protein
dataset with one or more of the available S-acylproteomics studies and a deeper exploration of the database.
DHHC protein S-acyltransferases
DHHC protein S-acyltransferases (DHHC PATs) were discovered decades after the first report on protein S-acylation, even after the identification of protein thioesterases,
the enzymes catalyzing the reverse reaction. Genetic studies in yeast led to the identification of the multipass transmembrane proteins Erf2/Erf4 and Akr1 as the enzymes
responsible for S-acylation of Ras2 and Yck2, respectively
(Bartels et al. 1999; Lobo et al. 2002; Roth et al. 2002). The
S-acyltransferase activity of Erf2 and Akr1 was shown to
be dependent on an Asp-His-His-Cys (DHHC) motif present within a 50 residue-long cysteine-rich domain
(CRD), giving the family its name and allowing the identification of S-acyltransferases in other organisms (Huang
et al. 2004; Keller et al. 2004; Fukata et al. 2006; Mitchell
et al. 2006). The DHHC S-acyltransferase family in yeast is
composed of five (Schizosaccharomyces pombe) to seven
(Saccharomyces cerevisiae) members and this number
goes up to 24 in mammals. Other model organisms such
as Caenorhabditis elegans and Drosophila melanogaster
also have about 20 predicted DHHC PATs (Bannan et al.
2008; Edmonds and Morgan 2014; Fukata et al. 2016).
Whether all members of the family are expressed and
functional in each organism remains to be established. It
is, however, intriguing that the number of enzymes is
roughly conserved, suggesting that the diversity originated early in evolution and has been maintained.
In humans, most DHHC S-acyltransferase family members have four transmembrane domains (Figure 1(B)),
except for zDHHC13, zDHHC17, and zDHHC23, which
are predicted to have six, and zDHHC4 and zDHHC24
predicted to have five. These last two cases are curious
as they imply that the N- (zDHHC4) or C-terminus
(zDHHC24) face the luminal/extracellular space, whereas
in all other DHHC PATs, both ends of the protein face
the cytosol. Even more oddly, zDHHC22 is predicted in
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
425
Table 2. Structural specificities of human DHHC S-acyltransferases (DHHC PATs).
DHHC-PAT
zDHHC1
zDHHC2
(REAM)
zDHHC3
(GODZ)
zDHHC4
zDHHC5
zDHHC6
zDHHC7
(Serz-b)
zDHHC8
zDHHC9
(CGI-89)
zDHHC11
zDHHC12
zDHHC13
(HIP14L, HIP3RP)
zDHHC14 (NEW1CP)
zDHHC15
zDHHC16
(Aph2)
zDHHC17
(HIP-14, HIP-3)
zDHHC18
zDHHC19
zDHHC20
zDHHC21
zDHHC22
zDHHC23
(NIDD)
zDHHC24
Evidence
of regulatory
S-acylation
(non-DHHC cysteines)
C4 (mouse) [1]
Predicted
disordered
regions
by IUPREDb
17–41 þ 324–358 þ 408–447
304–340
Uniprot AC
Q8WTX9
Q9UIJ5
N of
TMD
4
4
Protein
interaction
domainsa
–
–
Q9NYG2
4
PDZ-binding
–
Q9NPG8
Q9C0B5
Q9H6R6
Q9NXF8
5
4
4
4
–
PDZ-binding
SH3_2
–
–
Ser-rich : 359–479
Leu-rich : 193–214
–
C236/C237/C245 [1, 2]
C329/C330/C343 [1, 2, 3]
–
279–715
–
–
Q9ULC8
Q9Y397
4
4
PDZ-binding
–
Pro-rich: 656–744
–
C236/C237/C245 [1, 2]
C161 (mouse) [1]
293–765
299–364
Q9H8X9
Q96GR4
Q8IUH4
4
4
6
–
–
7 ankyrin repeats
–
–
Phe-rich : 328–391
374–412
–
–
Q8IZN3
Q96MV8
Q969W1
4
4
4
–
–
–
Leu-rich : 370–391
–
–
C310 (mouse) [1]
376–482
306–337
–
Q8IUH5
6
7 ankyrin repeats
–
C590 (mouse) [1]
–
Q9NUE0
Q8WVZ1
Q5W0Z9
Q8IVQ6
Q8N966
Q8IYP9
4
4
4
4
2 (4)c
6
–
–
–
–
–
–
Pro-rich : 11–46
–
–
–
–
–
Q6UX98
5
–
Leu-rich : 140–222
Regions with
compositional biasa
Arg-rich : 390–481
–
–
C263 [1, 2, 4]
C18/C19/C20/
C386 (mouse) [1]
9–67
277–309
–
–
–
–
–
“–” indicates that the feature was not predicted. TMD: transmembrane domain.
a
Protein interaction domains and regions with compositional bias were identified using ScanProsite (https://prosite.expasy.org/scanprosite/) (de Castro
et al. 2006).
b
Intrinsically disordered regions were predicted using IUPRED (http://iupred.enzim.hu/) (Dosztanyi et al. 2005). A threshold of disorder tendency >0.5 was
used to define disorder. Only regions with at least 30 consecutive amino acids predicted as disordered are listed. IUPRED is one of the many tools available to predict intrinsically disordered regions and appears to be one of the most conservative ones (B€
urgi et al. 2016).
c
Human zDHHC22 is annotated in Uniprot as having two TMDs but online prediction tools (TMHMM (www.cbs.dtu.dk/services/TMHMM/) and TMPRED
(embnet.vital-it.ch/software/TMPRED_form.html)) identify two additional domains, as explained in the text and shown in Figure 1(C).
[1] Collins et al. (2017).
[2] Yang et al. (2010).
[3] Abrami et al. (2017).
[4] Rana et al. (2018).
Uniprot (www.uniprot.org) to have only two transmembrane domains. Online prediction tools (TMPRED,
TMHMM) however identify two additional transmembrane segments, as shown in Figure 1(C) (embnet.vital-it.
ch/software/TMPRED_form.html, www.cbs.dtu.dk/services/TMHMM/) (Table 2). Importantly, the DHHC-CRD
always localizes to the cytosolic side thus restricting
DHHC PAT-mediated S-acylation to the cytosolic environment. Beyond the CRD there is little sequence homology between different family members, except for
three short conserved motifs: an Asp-Pro-Gly (DPG)
sequence located at the N-end of the cytosolic loop containing the DHHC-CRD; a Thr-Thr-Xxx-Glu motif (TTxE,
where x represents any amino acid) and a 16 residuelong Palmitoyltransferase Conserved C-terminal (PaCCT)
motif, both located after the last transmembrane domain
(Figure 1(B)) (Mitchell et al. 2006; Gonzalez Montoro
et al. 2009). As discussed later, N- and C-terminal
domains might also contain protein-protein interaction
domains/motifs involved in substrate specificity.
DHHC PATs have been found in most compartments
of the endomembrane system. A reliable assessment of
their subcellular distribution has however been hindered by the low protein copy number of these
enzymes (Table 3) and the lack of good antibodies
against the endogenous proteins. Studies using tagged
DHHC PATs indicate that in S. cerevisiae the seven
members of the family distribute between the endoplasmic reticulum (ER) (Erf2, Swf1, and Pfa4), the Golgi
apparatus (Akr1 and Akr2), the vacuole (Pfa3), and the
plasma membrane (Pfa5) (Ohno et al. 2006). In human
HEK293T cells, most ectopic overexpressed tagged
DHHC PATs were found to localize to the ER and the
Golgi apparatus, while a few were found at the plasma
426
M.-E. ZABALLA AND F. G. VAN DER GOOT
Table 3. Protein abundance (in copy number per cell) for enzymes involved in protein S(de)acylation, as reported in quantitative proteomics studies.
Study 1
U2OS cells
Beck
et al. (2011)
< 5 102
–
< 5 102
–
< 5 102
–
–
–
–
–
< 5 102
< 5 102
–
–
–
–
< 5 102
–
–
< 5 102
–
–
–
1.27 105
Study 2
HeLa cells
Nagaraj
et al. (2011)
–
–
1900
–
38,000
1500
–
360
–
–
–
10,000
600
–
–
5600
89
–
11,000
–
–
–
–
550,000
Enzyme
zDHHC1
zDHHC2
zDHHC3
zDHHC4
zDHHC5
zDHHC6
zDHHC7
zDHHC8
zDHHC9
zDHHC11
zDHHC12
zDHHC13
zDHHC14
zDHHC15
zDHHC16
zDHHC17
zDHHC18
zDHHC19
zDHHC20
zDHHC21
zDHHC22
zDHHC23
zDHHC24
LYPLA1 (APT1)
Uniprot AC
Q8WTX9
Q9UIJ5
Q9NYG2
Q9NPG8
Q9C0B5
Q9H6R6
Q9NXF8
Q9ULC8
Q9Y397
Q9H8X9
Q96GR4
Q8IUH4
Q8IZN3
Q96MV8
Q969W1
Q8IUH5
Q9NUE0
Q8WVZ1
Q5W0Z9
Q8IVQ6
Q8N966
Q8IYP9
Q6UX98
O75608
LYPLA2 (APT2)
O95372
6.71 104
470,000
ABHD17A
ABHD17B
ABHD17C
Q96GS6
Q5VST6
Q6PCB6
–
–
–
–
–
–
Study 3
HeLa cells
Hein
et al. (2015)
–
–
–
636
14,026
5189
4370
–
–
–
740
273
–
–
2055
7849
–
–
8125
–
–
–
–
1,251,791
30,161
–
–
–
Study 4
Human heart regions
Doll et al. (2017)
–
min: – ; max: 6764
min: – ; max: 524,496
min: – ; max: 25,724
min: – ; max: 39,351
–
–
–
–
–
–
min: – ; max: 73,797
–
–
–
–
–
–
–
–
–
–
–
min: 106,917
max: 1,432,461
min: 150,400
max: 1,069,056
min: – ; max: 245,813
–
–
“–” indicates no information available.
For study 4, only the minimum (min) and maximum (max) reported copy numbers are shown.
membrane (Ohno et al. 2006). ER retention was shown
to occur through C-terminal KKxx-like motifs for
zDHHC4 and zDHHC6 (Gorleku et al. 2011; Abrami et al.
2017). zDHHC2 and zDHHC5 were found to traffic
through the endocytic system in a manner dependent
on their C-terminus (Greaves et al. 2011; Brigidi et al.
2015). In synaptic spines, these two enzymes were
found in vesicles and at the plasma membrane, where
they regulate the localization and assembly of synaptic
proteins into domains (Fukata et al. 2016). Finally,
zDHHC8 and zDHHC13 were shown to regulate mitochondria architecture and function (Maynard et al.
2008; Shen et al. 2017), but mitochondrial localization
was not established. Also, these DHHC PATs were found
to have roles in other subcellular compartments
(Fernandez-Hernando et al. 2006; Huang et al. 2009;
Singaraja et al. 2009; Lemonidis et al. 2014).
Structure and mode of catalysis of DHHC PATs
DHHC PATs have been proposed to function by following
a two-step mechanism (Jennings and Linder 2012) where
the acyl-CoA provides the acyl chain to the catalytic
DHHC cysteine to form a transient acyl-enzyme
intermediate. This step is often referred to as
“autoacylation”. This nomenclature is however somewhat
confusing because several DHHC PATs undergo regulatory S-acylation on cysteines outside the DHHC motif
(Table 2) (Abrami et al. 2017; Collins et al. 2017). In a
second step, the acyl moiety is transferred from the cysteine of the enzyme to that of the protein substrate. As
discussed later, this two-step model has strong implications on the potential substrate recognition mechanisms.
In a recent seminal paper, the Banerjee group
reported the first X-ray structures of DHHC PATs (human
zDHHC20 and zebrafish zDHHC15), comprising the transmembrane and catalytic domains (Rana et al. 2018). The
two enzymes display remarkably similar structures. The
four transmembrane domains (TMDs) adopt a teepeelike structure (with the top facing the lumenal/extracellular compartments and the bottom facing the cytosol),
positioning the DHHC active site right at the cytosolic
aqueous-membrane interface (Figure 2). This location is
consistent with and might explain the fact that cytosolic
cysteine residues close to TMDs are so far the best Sacylation site candidates (Blanc et al. 2015). Concerning
the zinc ions known to bind to the CRD, the X-ray structures reveal that they play critical structural roles,
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
427
Figure 2. X-ray crystal structures of DHHC S-acyltransferases. (A) Cartoon representation of the X-ray crystal structures of human
zDHHC20 (cyan, PDB 6BMN) and zebrafish zDHHC15 (blue, PDB 6BMS) highlighting their strong overlap. (B) Cartoon representation of human zDHHC20 covalently bound to 2-bromopalmitate (PDB 6BML). The 2-bromopalmitate molecule is shown as red
and yellow spheres and the surface of the protein is depicted to highlight the insertion of the lipid into the hydrophobic cavity
formed by the four transmembrane domains of the protein. (C) Cartoon and sticks representation of the active site and acyl chain
cavity of human zDHHC20 covalently bound to 2-bromopalmitate (PDB 6BML). The four residues forming the DHHC motif are
shown in cyan, the acyl chain of 2-bromopalmitate is shown in yellow and the hydrophobic residues contributing to the acyl
chain binding cavity (as described in Rana et al. 2018) are shown in green. Highlighted in pink is residue Thr241 (from the TTxE
motif) forming a hydrogen-bonding network with the catalytic residues (dashed lines). Zn ions are shown as spheres in all representations. The disposition of the protein relative to the membrane is approximately indicated by light brown lines. Figure created using Pymol v0.99 (DeLano Scientific, San Carlos, CA) (see color version of this figure at www.tandfonline.com/ibmg).
allowing the correct positioning of the DHHC active site
residues and other residues involved in the catalytic site
hydrogen bond network.
The most striking feature of this structure is the
hydrophobic cavity stretching from the active site into
the membrane (Figure 2(B,C)). All four TMDs contribute
to this cavity, which accommodates the acyl chain to be
transferred to the substrate. Through structure-guided
mutagenesis, the authors proposed that the cavity acts
as a “hydrocarbon ruler”, its geometry determining the
acyl chain length preferences of the enzyme (Wyckoff
et al. 1998). This structural analysis explains the experimentally observed acyl chain preferences of zDHHC3
and zDHHC7 and confirms the predictions made by the
Chamberlain group in this respect (Greaves et al. 2017).
Rana, Kumar et al. propose an enzymatic mechanism
based on their structural observations, which is consistent with the two-step model. They show that the first
two residues of the DHHC motif form a hydrogenbonded pair and speculate that this histidine residue
could initially activate the DHHC cysteine for the nucleophilic attack on the carbonyl-thioester of the acyl-CoA
molecule and later on activate the acyl-enzyme thioester
for acylation of the protein substrate.
For some DHHC PATs, such as the yeast Swf1 and
Pfa4 and the human zDHHC17, acyl-enzyme intermediates could not be detected (Gonzalez Montoro et al.
2015; Greaves et al. 2017; Verardi et al. 2017), raising the
possibility of an alternative mechanism in which the first
histidine residue of the DHHC motif would favor the direct nucleophilic attack of the protein substrate to the
acyl-CoA (Gonzalez Montoro et al. 2015). Whether or not
an acyl-enzyme intermediate forms, it seems that the
first histidine residue of the DHHC motif plays a critical
role in catalysis. Interestingly, this residue is replaced by
a glutamine residue in human and murine zDHHC13.
Whereas this replacement is isosteric and it is not
expected to introduce major structural perturbations at
the active site, it is unclear whether this glutamine could
play a role in the nucleophilic attack or an alternative
mechanism needs to be evoked (Mitchell et al. 2010).
Finally, Rana, Kumar et al. speculate that the acyl-CoA
inserts into the hydrophobic cavity directly from within
the membrane (Rana et al. 2018). Whether this is actually the case remains to be addressed. It would require
lateral opening of the teepee-like structure to allow
membrane-anchored acyl-CoA to diffuse into the cavity.
Moreover, considering that DHHC PATs are low abundance proteins (Table 3) and at risk of being outcompeted by more abundant enzymes involved in lipid
biosynthesis and degradation, they might rely on acylCoA binding proteins to deliver acyl-CoA to them.
The observed teepee-like structure is probably common to all members of the DHHC S-acyltransferase family and will be invaluable in guiding the rational design
of inhibitors, a major unmet need in the field.
428
M.-E. ZABALLA AND F. G. VAN DER GOOT
Acyl chain selectivity of DHHC PATs
As alluded to earlier, S-acylation is not restricted to Spalmitoylation. Chains of different lengths and saturation may also be transferred. Indeed, a study in human
platelets revealed that 74% of the pool of S-acylated
proteins was bound to C16:0 while the remaining 26%
was bound to C18:0 or C18:1 acyl chains (Muszbek et al.
1999). While these proportions might partly be due to
the relative abundances of the corresponding acyl-CoA,
DHHC PATs may have an acyl chain preference as first
reported by the Linder group (Jennings and Linder
2012). Using purified DHHC PATs in micelles, zDHHC2
was found to incorporate and transfer a broad range of
acyl-CoA chain lengths, while zDHHC3 showed a clear
preference for C14 (myristoyl-CoA) and C16 (palmitoylCoA and palmitoleoyl-CoA) chains over longer fatty
acids. More recently, the Chamberlain group obtained
comparable results using clickable acyl-CoA analogs in
cell culture (Greaves et al. 2017). zDHHC2 and zDHHC4
did not show any fatty acid preference; zDHHC3, 5, 7,
11, and 15 preferred C14 and C16 fatty acids over C18
chains; zDHHC17 preferred C16 and C18 over C14; and
finally zDHHC23 displayed a marked preference for C18.
The acyl chain preference appears to be determined at
the formation of the acyl-enzyme intermediate and is
thus dictated by the geometry of the hydrophobic cavity revealed by the X-ray structures (Jennings and
Linder 2012; Greaves et al. 2017; Rana et al. 2018). This
is consistent, for instance, with the observation that
coimmunoprecipitation of zDHHC3 with its substrate
NCAM180 was only observed for the wild-type protein
and not for the catalytic-defective Cys157Ser mutant,
suggesting that in this case the formation of the acylenzyme intermediate is required for the interaction
with the protein substrate (Lievens et al. 2016).
Circumstantial evidence, however, indicates that
protein substrates might also dictate acyl length preference. Influenza A virus hemagglutinin has three “Sacylatable” cysteine residues, two of which were found
to be modified by palmitate (C16:0) and one by stearate (C18:0) (Kordyukova et al. 2008). Also, zDHHC6 was
found to catalyze the preferential addition of C16:0 to
the ER chaperone calnexin (Lakkaraju et al. 2012) but
of both C16:0 and C18:0 to transferrin receptor
(Senyilmaz et al. 2015).
Protein substrate recognition by DHHC PATs
One of the remaining enigmas in the field is: what
makes a cytosolic cysteine an S-acylation site? Is it protein structure? Is it sequence? Is it its proximity to
membranes? What controls the enzyme-protein substrate specificity?
A first layer of specificity is provided by the subcellular localization, since the presence of the enzymes in a
specific membrane-bound compartment confines their
action to transmembrane protein substrates that pass
by or reside in this compartment. For soluble proteins,
it is generally believed that for S-acylation to occur, the
protein substrate needs to weakly bind to membranes
through either another lipidation event (e.g. myristoylation or prenylation) or some intrinsic weak membrane
affinity (electrostatic or hydrophobic) (Shahinian and
Silvius 1995; Greaves et al. 2008, 2009; Weber et al.
2017). While some DHHC PATs have been shown to
preferentially acylate soluble, transmembrane, or lipidated protein substrates (Roth et al. 2006; Gonzalez
Montoro et al. 2011; Ohno et al. 2012), a firm conclusion
on this classification awaits a comprehensive view of
enzyme-substrate pairs.
Protein substrate recognition can also be mediated
by protein-protein interaction motifs/domains in the Nor C-termini of DHHC PATs. zDHHC3, 5, and 8 contain
potential PDZ-binding motifs; zDHHC13 and 17 contain
several N-terminal ankyrin repeats; and zDHHC6 contains a C-terminal SH3_2 domain (Singaraja et al. 2002;
Tsutsumi et al. 2008; Gottlieb and Linder 2017). The
ankyrin repeats of zDHHC13 and 17 mediate their interaction with proteins such as huntingtin (HTT), SNAP23,
SNAP25, and cysteine string protein (CSP). Moreover, an
ankyrin repeat-binding short linear motif was identified
in these and other protein substrates (Huang et al.
2009; Lemonidis et al. 2014, 2015). The ankyrin repeat
domain of zDHHC17 (ANK17) was co-crystalized with
the SNAP25b recognition motif revealing key residues
involved in binding that were also present in another
substrate, HTT (Gao et al. 2009; Verardi et al. 2017).
When an ankyrin repeat domain was added to zDHHC3,
the enzyme acquired the ability to S-acylate HTT
(Huang et al. 2009). Similarly, the PDZ-binding domains
of zDHHC5 and 8 are required to recruit their respective
substrates Grip1 and Pick1 (Thomas et al. 2012, 2013).
The presence of a classical protein-protein interaction
domain does not exclude binding to some protein substrates through other domains. zDHHC5 was indeed
found to bind phospholemman through an intrinsically
disordered 120 residue-long region after the fourth
transmembrane domain (TMD) (Howie et al. 2014).
Interaction with their protein substrates through disordered regions could be a common theme amongst
DHHC PATs (Table 2) (Howie et al. 2014; Gottlieb and
Linder 2017). It is important to note that interaction of
a protein that can undergo S-acylation with a member
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
of the DHHC S-acyltransferase family does not imply
that it is a substrate of that enzyme (see section Nonenzymatic functions of DHHC PATs). For instance,
zDHHC13 strongly interacts with SNAP25 and CSP, but
does not modify them (Lemonidis et al. 2014).
On the protein substrate side, specific sequences
or regions involved in interaction with their
S-acyltransferases have also been reported. For instance,
the C-terminal membrane-binding domain of ClipR-59
was shown to mediate the interaction of the protein
with its S-acyltransferase zDHHC17 (Ren et al. 2013b). In
turn, an amphipathic a-helix in the large intracellular
loop of the electrogenic sodium/calcium exchanger
NCX1 was recently found to critically control the S-acylation of one specific cysteine residue (Plain et al. 2017).
A potential interplay between an amphipathic helix and
the S-acylation of nearby cysteine residues has also
been suggested for the interferon-induced transmembrane protein IFITM3 (Chesarino et al. 2017).
Strikingly, some residues or regions were shown to
dictate the interaction of a protein substrate with only
one of its several DHHC PATs. The interaction between
SNAP25b and one of its S-acyltransferases, zDHHC17,
was shown to be regulated by a proline residue
(Pro117) near the S-acylation sites (Greaves et al. 2009),
which it is proposed to constrain SNAP25b in order to
fit into the concave side of the ANK17 domain (Verardi
et al. 2017). This residue however does not seem to be
important for the interaction of SNAP25b with zDHHC3
or 7, also responsible of its S-acylation but not having
ankyrin repeat domains (Greaves et al. 2009). Similarly,
the closely related SNAP25 and SNAP23 SNARE proteins
are not S-acylated by the exact same set of DHHC PATs
as zDHHC15 is active against SNAP25b but not SNAP23.
However, a single Cys-to-Phe mutation in the cysteinerich domain of SNAP23 renders it susceptible to
zDHHC15-mediated S-acylation (Greaves et al. 2010). In
yeast, it was shown that the vacuolar protein Vac8 is Sacylated exclusively by Pfa3, but the isolated SH4
domain of Vac8 (containing the cysteine residues target
of S-acylation) could get S-acylated by all five DHHC
PATs tested in the study (Nadolski and Linder 2009).
The proximity of cysteine residues to TMDs makes
them strong candidates for S-acylation, despite the fact
that this is not picked up by the available prediction
programs (Blanc et al. 2015). In a recent study based on
native MS, Rodenburg et al. proposed that S-acylation
occurs stochastically whenever a cysteine localizes at
the appropriate distance from a membrane (Rodenburg
et al. 2017). In support of their hypothesis, they
observed that prokaryotic proteins (normally not S-acylated (Mitchell et al. 2006)) undergo S-acylation when
429
ectopically expressed in human cells. Mutagenesis
experiments on the mammalian hepatic asialoglycoprotein receptor (ASGP-R) similarly indicated that the distance of the cysteine residue relative to the TMD is the
major determinant for its S-acylation, more so than the
identity of surrounding amino acids (Yik and Weigel
2002). In contrast with these reports, early studies on
viral glycoproteins showed that the spacing between
the cysteine residue and the TMD was not sufficient but
rather S-acylation was promoted by conformational signals contained in the sequence of the transmembrane
and cytoplasmic domains (Ponimaskin and Schmidt
1998). Likewise, MS-based thermodynamics studies
indicated that a proline residue at the -1 position of
the target cysteine increases flexibility and lowers the
enthalpic barrier for the S-acylation reaction (Khanal
et al. 2015). The presence of prolines was similarly
found to influence other post-translational modifications, such as phosphorylation and N-glycosylation
(Bause 1983; Zhu et al. 2005; Alexander et al. 2011).
When analyzing reported S-acylation sites, proline residues at the -1 position are however relatively rare. In
contrast, positively charged residues are frequently
found after S-acylation sites (Blanc et al. 2015) and it
has been suggested that this could favor binding of the
negatively charged acyl-CoA molecule and/or increase
the ionization of the target thiol (Bizzozero et al. 2001).
Thus, while juxtamembrane cysteines are frequent Sacylation sites, not all proteins having cysteine residues
at this location get S-acylated. Moreover, it is unclear
that their S-acylation is stochastic since this would indicate that any DHHC PAT that localizes to the same
membranes could modify target proteins, which conflicts with reported enzyme–protein substrates specificities. Also, stochasticity is a priori incompatible with our
observations that the two contiguous “acylatable” cysteine residues in the ER chaperone calnexin get S-acylated in a preferred sequential manner (Dallavilla et al.
2016). It thus remains to be investigated whether the
reported stochastic S-acylation corresponds to an
enzymatic or non-enzymatic process.
Finally, whether it is important for substrate recognition or not, some DHHC PATs require cofactors to be
active. In S. cerevisiae, Erf2 was shown to depend on
Erf4 to acylate Ras (Lobo et al. 2002). Interestingly, this
requirement seems to be evolutionarily conserved as
the Erf2 orthologs Sp-Erf2 in S. pombe and zDHHC9 in
human rely on Sp-Erf4 and GCP16, respectively, to modify Ras (Swarthout et al. 2005; Young et al. 2014).
Mammalian genomes contain several GCP16 related
proteins suggesting that other DHHC PATs might also
employ cofactors (Tsutsumi et al. 2008). Human
430
M.-E. ZABALLA AND F. G. VAN DER GOOT
zDHHC6 was reported to interact with selenoprotein K
(also known as SelK or SelenoK) via its SH3_2 domain
and this interaction to be required for activity
(Fredericks et al. 2014). SelK was first found as necessary for the palmitoylation of CD36 in macrophages
(Meiler et al. 2013) and later on proposed to be a
zDHHC6 cofactor for the S-acylation of the inositol
1,4,5-triphosphate (IP3) receptor type 1 (IP3R1) and of
ASAP2 (Fredericks et al. 2014; Fredericks and Hoffmann
2015; Norton et al. 2017).
out non-canonical functions, since both the wild-type
and DHHA variants could regulate the actin cytoskeleton and polarized secretion by interacting with the
Rho GTPase Cdc42 (Dighe and Kozminski 2008). DHHA
Swf1 might, however, have partial S-acyltransferase
activity since it was shown that a DHHR Swf1 variant as
well as DHHA and DHHR variants of Pfa4 retain partial
activity (Gonzalez Montoro et al. 2015). Whether canonical and non-canonical functions of these enzymes are
coordinated or whether they influence one another is
currently unknown.
Non-enzymatic functions of DHHC PATs
It is important to keep in mind that some DHHC PATs
might exert functions that are independent of their Sacyltransferase activity. zDHHC17 was found to contribute to acute ischemic brain injury through interaction
with c-Jun N terminus kinase (JNK) and recruitment of
its mitogen-activated protein kinase kinase MKK7, independently of its S-acyltransferase activity (Harada et al.
2003; Yang and Cynader 2011). Both zDHHC13 and 17
were reported to mediate Mg2þ transport and proposed
to be “chanzymes”, i.e. proteins mediating ion transport
and bearing an enzymatic activity (Goytain et al. 2008).
zDHHC1 was found to be a positive regulator of STING
(stimulator of interferon genes), involved in DNA virustriggered innate immunity (Zhou et al. 2014).
Intriguingly, STING does undergo S-acylation and this is
necessary for its ability to signal (see below) (Mukai et al.
2016), yet S-acylation is not mediated by zDHHC1.
zDHHC1 appears to act upstream of STING, independently of its DHHC motif, promoting STING multimerization and recruitment of the downstream signaling
components TBK1 and IRF3 (Zhou et al. 2014). A role for
zDHHC1 in STING-dependent innate immune signaling
was also reported in teleosts but whether this required
acyltransferase activity was not addressed (Xu et al.
2017). The S-acyltransferase zDHHC16 was first named
Abl-philin 2 (Aph2) given it was identified as a c-Ablinteracting protein (Li et al. 2002). Aph2 was proposed
to regulate the levels of c-Abl and jointly affect apoptosis irrespective of the presence of its DHHC motif.
zDHHC16 has subsequently been linked to DNA damage
repair. In the latter case, however, its involvement
required the S-acyltransferase activity (Cao et al. 2016).
In yeast, Akr1 was shown to be responsible for the
suppression of Gbc-mediated mating signaling pathway
in the absence of mating pheromone independently of
its S-acyltransferase activity (Hemsley and Grierson
2011). Gbc requires S-acylation for plasma membrane
targeting but this modification is mediated by Erf2 and
Swf1 (Roth et al. 2006). In turn, Swf1 might also carry
Acyl protein thioesterases
The S-deacylation reaction is catalyzed by a set of
enzymes named acyl protein thioesterases (APTs) or
deacylases/depalmitoylases, belonging to the metabolic
serine hydrolase superfamily (mSH) (Long and Cravatt
2011). Members of the mSH family adopt an a/b hydrolase fold, consisting of a central b-sheet surrounded by
a-helices, and rely on an active site serine for catalysis.
This serine is part of a catalytic dyad or triad, which
increases its nucleophilicity and allows for the attack of
the substrate (Long and Cravatt 2011). The first deacylase identified was palmitoyl-protein thioesterase 1
(PPT1) (Camp and Hofmann 1993) but it was later
shown to reside inside lysosomes and thus not responsible for deacylation of cytosolic substrates (Verkruyse
and Hofmann 1996). Its homolog PPT2 appears to be
devoid of catalytic activity (Soyombo and Hofmann
1997; Calero et al. 2003). Two other members of the
family, APT1 and APT2 (also called LYPLA1 and LYPLA2,
respectively), were first identified as lysophospholipases
(Sugimoto et al. 1996; Toyoda et al. 1999) but later
shown to have enhanced activity as protein thioesterases (Duncan and Gilman 1998; Rocks et al. 2005; Rusch
et al. 2011). The crystal structures of human APT1 and
APT2 confirm they have a classical catalytic triad with
the nucleophilic residue in a strained conformation
between a b-strand and an a-helix, although they adopt
a rather atypical a/b hydrolase fold with a sevenstranded b-sheet (Figure 3) (Devedjiev et al. 2000; Won
et al. 2016). A detailed review on protein depalmitoylases has been recently published (Won et al. 2018).
Few targets have so far been reported for these acyl
protein thioesterases but some substrate specificity is
already apparent. APT2 but not APT1 hydrolyzes prostaglandin glycerol esters (Manna et al. 2014) and regulates the dynamic S-acylation of human zDHHC6
(Abrami et al. 2017), GAP-43 (Tomatis et al. 2010), and
Scrib (Hernandez et al. 2017). In contrast, BK channel
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
Figure 3. Tridimensional structure of acyl protein thioesterases. (A) Cartoon representation of the X-ray crystal structure
of human APT1 (PDB 1FJ2) showing its seven-stranded a/b
hydrolase fold and highlighting the position of the catalytic
triad (Ser119, Asp174, and His209, shown as sticks). (B)
Cartoon and sticks representation of the active site of human
APT2 co-crystallized with its selective inhibitor ML349 (PDB
5SYN). The inhibitor is shown in green and the catalytic triad
Ser122-Asp176-His220 is highlighted in cyan. Figure created
using Pymol v0.99 (DeLano Scientific, San Carlos, CA) (see
color version of this figure at www.tandfonline.com/ibmg).
deacylation was reported to be catalyzed by APT1 but
not by APT2 (Tian et al. 2012).
APT1 and APT2 were reported, and widely accepted,
to display a cytosolic localization with preferred membrane association to the Golgi apparatus and the
plasma membrane based on overexpression of fusion
proteins using myc-FLAG (Kong et al. 2013) or mCitrine
(Vartak et al. 2014) tags. We have, however, recently
found that endogenous APT1 or APT1 overexpressed as
a fusion to small tags localize primarily (although not
exclusively) to mitochondria, whereas overexpression of
APT1 using a bulky tag such as mCitrine prevents it
from accumulating in mitochondria (Kathayat et al.
2018). While Kong et al. (2013) concluded that there
was a clear plasma membrane staining, they did not
discuss the membrane-associated intracellular staining
observed in their microscopy images, which, to our
opinion, is not incompatible with a mitochondrial localization of their construct. APT2 does not display this
sensitivity to tag size showing a broad distribution
including the cytosol, the Golgi apparatus and the
plasma membrane (Kong et al. 2013; Vartak et al. 2014;
Kathayat et al. 2018). Whereas APT1 has been included
in the Mitocarta compendium as a potential mitochondrial protein (Pagliarini et al. 2008; Calvo et al. 2016), no
clear mitochondrion-targeting sequence has been
reported. Besides, given the high sequence identity
between APT1 and APT2, it is not known which features
determine their differential subcellular localization.
Mitochondria-targeting sequences are predicted to
form amphipathic a-helices and to be enriched in
431
positively charged residues (Bolender et al. 2008). A
close examination of the N-termini of APT1 and APT2
reveals the presence of three negatively charged residues in APT2, absent in APT1, which we speculate could
interfere with a potential weak mitochondrial targeting.
So far, no human diseases have been linked to mutations in the genes encoding APT1 or APT2. Still, these
proteins are remarkably conserved across mammals
and beyond, suggesting that their functions (which
may not be restricted to protein S-deacylation) are
under strong evolutionary pressure.
An homolog of APT1, LYPLAL1, was reported to
mediate deacylation of BK channels (Tian et al. 2012).
However, the X-ray crystal structure of LYPLAL1 reveals
that this enzyme has a shallow, solvent-accessible
active site with some charged residues preventing the
formation of a hydrophobic cavity of the required size
to fit long acyl chains as those bound to S-acylated pro€rger et al. 2012). Interestingly, the main resiteins (Bu
dues affecting the shape of the active site are highly
conserved in the LYPLAL but less conserved in the
LYPLA subfamily. Furthermore, the LYPLAL subfamily
€rger et al.
seems to be present only in metazoans (Bu
2012). These observations suggest that the LYPLAL subfamily might play an important role hydrolyzing yet
unidentified small substrates.
In the search for additional deacylases, the
a/b-hydrolase domain-containing protein 17 members A,
B, and C (ABHD17A-C) were found to deacylate N-Ras
and the neuronal protein PSD-95 (Lin and Conibear
2015; Yokoi et al. 2016; Lin et al. 2017). Also, ABHD17A-C
were reported to mediate dynamic S-acylation of microtubule-associated protein 6 (MAP6), controlling its shuttling between membranes and microtubules and its
retention in axons (Tortosa et al. 2017). ABHD12 and
ABHD13 have also been reported to reduce PSD-95 Sacylation upon overexpression (Yokoi et al. 2016). Most
likely other acyl protein thioesterases remain to be identified. Protein thioesterases (notably APT1 and APT2) are
much more abundant than S-acyltransferases (copy
numbers per cell up to several orders of magnitude
higher, Table 3). A similar quantitative balance (i.e. more
genes codifying for the direct enzymes and less but
more abundant proteins catalyzing the reverse reaction)
has been reported for kinases and phosphatases as well
as regulators of histone acetylation and protein ubiquitination, suggesting a general feature of reversible posttranslational modifications (Smoly et al. 2017).
Interestingly, APT1, APT2, and ABHD17A-C all have
been reported to undergo S-acylation near their N-terminus and that this acylation controls their localization
(Yang et al. 2010; Kong et al. 2013; Vartak et al. 2014;
432
M.-E. ZABALLA AND F. G. VAN DER GOOT
Won et al. 2018). An additional S-acylation site has
been identified by MS for APT1 but not yet characterized (Yang et al. 2010). APT1 was reported to catalyze
its own deacylation and that of APT2, and knockdown
of APT1 was shown to promote APT2 plasma membrane association, but not vice versa (Kong et al. 2013).
Whether S-acylation (by which DHHC PATs?) regulates
APT function remains to be addressed.
Deacylase substrate specificity
Very little is known about substrate recognition by
acyl protein thioesterases. While deacylases other
than APT1/2 and ABHD17 are likely to exist, the lower
number of different deacylating enzymes compared to
S-acyltransferases suggests that the former are more
promiscuous. Several reports, however, indicate that
enzymatic deacylation also relies on protein substrate
recognition mechanisms (Tomatis et al. 2010; Lin and
Conibear 2015; Abrami et al. 2017; Hernandez et al.
2017; Tortosa et al. 2017). Besides, as for DHHC PATs,
differential subcellular localization of deacylases
would already restrict their set of substrates. We have
recently shown that APT1, but not APT2, localizes primarily to mitochondria, which immediately opens a
new spectrum of substrates for APT1 inaccessible to
APT2 (Kathayat et al. 2018).
Interestingly, recognition by APTs can also be controlled by conformational changes in the protein substrate. S-deacylation of H-Ras has indeed been shown
to be dependent on the cis-trans isomerization of a
peptidyl-prolyl bond nearby the S-acylated cysteine residues (Ahearn et al. 2011). The prolyl-isomerase FKBP12
was shown to bind to H-Ras in an acylation-dependent
manner and promote protein S-deacylation. The use of
peptidyl-prolyl isomerization as a molecular timer to
trigger protein S-deacylation could extend beyond this
particular example of H-Ras, as it has been reported
that proline residues are enriched close to S-acylation
sites (Khanal et al. 2015).
Inhibitors and probes
The S-acylation field has suffered from the lack of good
probes (there are no commercially available antibodies
against acylated cysteines) and specific inhibitors for
Figure 4. Chemical structures of the S-(de)acylation inhibitors and probes mentioned in this review (see color version of this figure at www.tandfonline.com/ibmg).
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
both S-acylation and deacylation. In the past two decades, 2-bromopalmitate (2-BP) (Figure 4) has been extensively used as a general inhibitor of protein S-acylation.
Inhibition of DHHC PATs is based on the formation of
an irreversible covalent bond between the thiolate
group of the catalytic cysteine (from the DHHC motif)
and the second carbon of the 2-BP molecule, with the
bromide acting as an excellent leaving group in the
nucleophilic substitution reaction. In 2013, the Martin
group published an MS-based profiling study where
they identified about 450 protein targets of the clickchemistry analog of 2-BP, 2BPN3; among them several
(but not all) DHHC S-acyltransferases (Davda et al.
2013). Alarmingly, most of the identified 2BPN3 targets
were proteins unrelated to S-acylation including carriers, transporters, and channels, as well as many proteins known to be susceptible of S-acylation themselves
(Davda et al. 2013). Similar results were obtained by
Zheng et al. (2013) using other 2-BP-based probes.
Furthermore, 2-BP was long-ago shown to inhibit the
activity of carnitine palmitoyl transferase (Chase and
Tubbs 1972), thus disrupting cellular lipid metabolism,
and even responsible of the in vitro inhibition of
enzymes that are not related to lipid metabolism
(Coleman et al. 1992). Whereas 2-BP treatment has
been considered a useful tool to study protein S-acylation, the available evidence on its off-target effects
and cellular toxicity prevents firm conclusions to be
drawn (Lanyon-Hogg et al. 2017).
Likewise, the natural antifungal and antibacterial
agent cerulenin (Figure 4) was reported to inhibit protein S-acylation (Lawrence et al. 1999) but it is also
known to bind and inhibit b-keto-acyl-ACP synthase,
HMG-CoA synthetase and other lipid metabolic
enzymes (Resh 2006). The natural compound tunicamycin was also shown to inhibit protein S-acylation
(Patterson and Skene 1994) but its major application in
cell biology relies on its inhibitory effects on protein
glycosylation (Kuo and Lampen 1974). Fortunately, the
effects of these two compounds beyond protein S-acylation are better described and thus their use as exclusive protein S-acylation inhibitors is rare.
A rather similar situation held true, until recently, for
the inhibition of the S-deacylation reaction. The acylb-lactone Palmostatin B (Figure 4) was originally
reported to be an APT1-specific inhibitor (Dekker et al.
2010) and used as such in several studies but later on
found to act on APT2 as well (Rusch et al. 2011).
Moreover, activity-based protein profiling studies indicate that this compound can also inactivate several
other lipid-processing serine hydrolases, including
members of the ABHD17 family, and even enzymes
433
unrelated to protein S-deacylation such as PNPLA6 and
FASN (Lin and Conibear 2015). Other serine hydrolases
inhibitors with varied activity on APTs (including other
b-lactones, piperazine amides, benzodiazepinediones,
boronic acids, triazole ureas, and chloroisocoumarins)
have been reported and are reviewed elsewhere
(Davda and Martin 2014). Especially in the case of
deacylases, where it is probable that other S-deacylating enzymes exist, the use of so-called specific inhibitors demands special caution. As a promising
alternative, Adibekian et al. have reported the design of
two reversible selective inhibitors: ML348 acting on
APT1, and ML349 acting on APT2 (Adibekian et al.
2010a, 2010b). Evidence on their selectivity in cellulo is
starting to become available (Vujic et al. 2016;
Hernandez et al. 2017). These results also bring optimism to the S-acylation field in the quest for DHHC
PAT-selective inhibitors. It has been long thought that
the high sequence conservation around the catalytic
site of DHHC PATs would prevent the design of selective inhibitors. APT1 and APT2 share more than 60%
sequence identity. The crystal structures of the APT1ML348 and APT2-ML349 complexes reveal that despite
the remarkable structural similarity of both polypeptides, each compound adopts a distinct conformation
within the active site leading to selective inhibition
(Figure 3(B)) (Won et al. 2016).
Finally, promising fluorescent in vivo depalmitoylation activity probes, named DPPs, were recently developed by the Dickinson group (Kathayat et al. 2017; Qiu
et al. 2018; Kathayat et al. 2018). DPPs feature a profluorescent molecule tethered through a carbamate
linkage to an S-acylated peptide substrate and have
been tuned to localize to specific compartments (DPP-2
and DPP-3 in cytosol, and mito-DPP-2 and mito-DPP-3
in mitochondria), to react better against APT1 than
APT2 (DPP-3 versus DPP-2 variants), or to have
increased water solubility (DPP-5) (Figure 4).
Non-enzymatic protein S-acylation and
deacylation
While in cells S-acylation is generally thought to be
enzymatic, this reaction can occur spontaneously in
vitro, as was shown for several proteins when mixed
with palmitoyl-CoA in absence of any enzymes
€mmel et al.
(Bizzozero et al. 2001; Yang et al. 2005; Ku
2006; Jenkins et al. 2011; Chan et al. 2016). Besides, due
to the intrinsic lability of the thioester linkage, it is
assumed that steady state non-enzymatic hydrolysis
may occur leading to deacylation (Davda and Martin
2014). Still, to what extent spontaneous S-(de)acylation
434
M.-E. ZABALLA AND F. G. VAN DER GOOT
occurs in vivo and what its physiological relevance is
remains unknown. In vivo, acyl-CoA binding proteins
(ACBPs) bind medium and long-chain acyl-CoA with
high affinity and act as intracellular carriers thus regulating acyl-CoA availability (Knudsen et al. 2000; Burton
et al. 2005). Early work has shown that addition of
ACBP at physiological molar ratios compared with longchain acyl-CoA strongly inhibited spontaneous protein
S-acylation in vitro (Leventis et al. 1997). In turn, the
addition of ACBP was shown to have only a minor
inhibitory effect on in vitro PAT-mediated S-acylation
(Dunphy et al. 2000). Calculations based on reported
cytosolic concentrations of acyl-CoA and ACBP in mammalian cells suggest that physiological non-enzymatic
protein S-acylation would be strongly suppressed and
thus enzyme-mediated S-acylation is likely the predominant mechanism in vivo (Leventis et al. 1997; Dunphy
et al. 2000; Knudsen et al. 2000). It remains unknown
whether DHHC PATs use ACBP-acyl-CoA complexes as
substrates or they are able to efficiently function at the
very low concentrations of free acyl-CoA available in
the
cytosolic
environment
(Faergeman
and
Knudsen 1997).
Interestingly, the transport protein particle component Bet3 and the TEAD transcription factors were
reported to undergo non-enzymatic “autoacylation”,
which was shown to be non-dynamic and to play a role
€mmel et al.
in protein structure and/or stability (Ku
2006; Chan et al. 2016; Noland et al. 2016). Proteins
having high affinity acyl-binding pockets might represent a special case in this respect.
A different situation could take place at subcellular
compartments with different physiological concentrations
of acyl-CoA and other players, such as cytosolic sites of
acyl-CoA generation or inside mitochondria (Faergeman
and Knudsen 1997). Whether local high concentrations of
acyl-CoA in the cytosol are linked to non-enzymatic or
compartmentalized DHHC PAT-mediated protein S-acylation remains to be explored. A variety of proteins in the
mitochondrial matrix, including >methylmalonate semialdehyde dehydrogenase, aldehyde dehydrogenase, glutamate
dehydrogenase,
nicotinamide
nucleotide
transhydrogenase, adenine nucleotide transferase, carbamoyl-phosphate synthetase 1, and several mitochondrial
carriers have been found to be S-acylated. In some cases,
S-acylation was shown to occur at the active site and
resulted in enzyme inhibition (Stucki et al. 1989;
Berthiaume et al. 1994; Corvi et al. 2001; Kostiuk et al.
2008). So far, no DHHC PATs have been reported in mitochondria, even though, as discussed above, protein Sdeacylation was found to occur in mitochondria (Kathayat
et al. 2018). Together these observations raise the
possibility that enzymatic S-deacylation could play a critical role in mitochondria homeostasis by “repairing”
uncontrolled, spontaneous S-acylation of mitochondrial
proteins, as proposed for sirtuin-3 in reverting toxic lysine
acetylation (Wagner and Hirschey 2014; Weinert
et al. 2015).
Physiological relevance of S-acylation
While it has been known for many years that diverse
physiological processes critically depend on protein Sacylation, the true importance of this modification is
only starting to be revealed. Recent reviews have summarized the roles of S-acylation in synaptic plasticity,
immunological synapses, host–pathogens interactions,
and membrane trafficking (Fukata and Fukata 2010;
Aicart-Ramos et al. 2011; Blanc et al. 2013; Chamberlain
and Shipston 2015; Fukata et al. 2016; Globa and Bamji
2017), therefore, these subjects will not be covered
here. Additionally, recent studies highlighted the role of
protein S-acylation in modulating membrane curvature
in Influenza A virus assembly (Chlanda et al. 2017); regulating the shuttling of SCP1 between the nucleus and
other subcellular compartments (Liao et al. 2017);
among others. This section provides selected recent
examples illustrating important regulatory roles of protein S-acylation in several physiological processes.
Interplay with ubiquitination and protein
homeostasis
Work from several groups, including ours, has highlighted the interplay between protein S-acylation and
ubiquitination and the regulation of protein homeostasis. S-acylation of the yeast SNARE protein Tlg1 was
shown to prevent its ubiquitination and subsequent
degradation in the vacuole by reducing the accessibility
of the lysine residues nearby the S-acylation site to the
E3 ubiquitin ligase (Valdez-Taubas and Pelham 2005).
Likewise, S-acylation of the anthrax toxin receptors
TEM8 and CMG2 prevent their association with lipid
rafts and, thus, their premature ubiquitination and lysosomal degradation (Abrami et al. 2006). In the biosynthetic pathway, S-acylation of the yeast chitin synthase
Chs3 and the Wnt signaling co-receptor LRP6 was
shown to be required for their ER export (Lam et al.
2006; Abrami et al. 2008). In the latter case, we proposed the existence of a ubiquitin-dependent folding
system controlling the proteostasis of poor folders, in
which S-acylation would prevent targeting to ER-associated degradation (ERAD) allowing proteins more time
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
to fold and thus increase the flux of proteins exiting the
ER (Abrami et al. 2008; Perrody et al. 2016).
The interferon-induced transmembrane protein 3
(IFITM3, see below) was also shown to be regulated by
the opposing effects of S-acylation and ubiquitination
(Yount et al. 2012). Along the same line, S-acylation of
the related protein IFITM1 and of the chemokine receptor CCR5 regulates protein turnover by preventing their
proteasomal or lysosomal degradation, respectively
(Percherancier et al. 2001; Hach et al. 2013). Interplay
between S-acylation and ubiquitination was also
reported for sortilin, a sorting receptor involved in targeting proteins to the secretory or endocytic compartments. S-acylation enables sortilin to be recycled back
from endosomes to the Golgi apparatus whereas nonS-acylated sortilin gets mono-ubiquitinated and trafficked to lysosomes for degradation (McCormick et al.
2008; Dumaresq-Doiron et al. 2013). Notably, dimerization of sortilin through a disulfide bond involving the
same cysteine residue identified as S-acylation site
leads to trafficking of sortilin to extracellular vesicles
(Itoh et al. 2018). Thus, the amount and localization of
sortilin appears to depend on the coordinated regulation of S-acylation, ubiquitination and disulfide
bond formation.
The counteracting effects of S-acylation and ubiquitination are likely to be widespread. Systems analysis,
aided by the S-palmitoylation database SwissPalm, has
indeed revealed that whereas ubiquitination sites are
annotated for about one third of the total human
proteome, this number almost doubles when considering the human candidate S-acylproteome (Blanc et al.
2015). Individual knock-out of zDHHC2, 5, or 6 in HAP1
cells was associated with a general increase in protein
ubiquitination (Blanc et al. 2015).
Moreover, several proteins involved in protein
quality control are S-acylation substrates, including
the ER chaperone calnexin, the transitional ER
ATPase (VCP), several components of the CCT micro
complex (TRiC), and the E3 ligase gp78 (Fairbank
et al. 2012; Lakkaraju et al. 2012; Lynes et al. 2012;
Blanc et al. 2015; Fang et al. 2016). Thus, the impact
of S-acylation on protein turnover is not limited to
its direct effect on a particular protein but it can
include indirect effects such as regulation of members of the cellular protein homeostasis machinery.
Additional indirect effects on turnover have been
observed, such as S-acylation-mediated membrane
targeting of inhibitory Smads (I-Smads), which negatively regulate TGF-b/BMP (bone morphogenetic protein) signaling by targeting type I receptors for
ubiquitination and turnover (Li et al. 2017).
435
Cell death
S-Acylation was found to regulate components of both
the extrinsic and the intrinsic apoptosis pathways. The
cell death receptor Fas (CD95) and its ligand FasL were
shown to get S-acylated (Chakrabandhu et al. 2007; Feig
et al. 2007; Guardiola-Serrano et al. 2010). zDHHC7-mediated S-acylation of Fas facilitates a series of events
required for cell death signaling including (i) formation
of supramolecular receptor aggregates, (ii) targeting to
specific membrane nanodomains, and (iii) recruitment of
ezrin and the actin cytoskeleton leading to FasL-induced
Fas internalization. Additionally, Fas S-acylation has been
reported to prevent its degradation through the lysosomal pathway thus guaranteeing a proper expression
level (Rossin et al. 2015). Moreover, Fas-FasL binding
leads to rapid and transient S-acylation of the tyrosine
kinase Lck and activation of downstream proteins in Tcells (Akimzhanov and Boehning 2015). Inhibition of Lck
S-acylation by downregulation of the corresponding
enzyme, zDHHC21, prevents Lck activation and Fas signaling and renders cells resistant to Fas-mediated apoptosis (Akimzhanov and Boehning 2015).
The intrinsic apoptosis pathway relies on the proapoptotic BCL-2-associated X protein (BAX), which
resides primarily in the cytosol of non-apoptotic cells
but stably associate with mitochondria to trigger the
permeabilization of the outer mitochondrial membrane
(OMM) and the release of cytochrome c into the cytosol
(Youle and Strasser 2008). A constitutive shuttling
between cytosol and mitochondria (Schellenberg et al.
2013) and a targeting to mitochondria modulated by Nterminal acetylation (Alves et al. 2018) have been
postulated for BAX. Notably, it has been shown that
association of BAX with the OMM and its subsequent
proapoptotic action is regulated by S-acylation at a single cysteine residue, and that a lack of BAX S-acylation
reduces BAX mitochondrial translocation and oligomerization as well as caspase activation and apoptosis
€hlich et al. 2014). Significant reduction of BAX S(Fro
acylation levels was observed in malignant B cells from
patients with Hodgkin disease, suggesting this could be
a mechanism by which tumor cells resist apoptosis
€hlich et al. 2014).
(Fro
Caspase-6 (CASP6), another important player in apoptosis, has also been shown to be S-acylated, in this case
by zDHHC17 (Skotte et al. 2017). Reduced CASP6 S-acylation resulted in increased CASP6 activity, and vice versa
(Skotte et al. 2017). Importantly, CASP6 was shown to
play a critical role in neurodegenerative diseases such as
Alzheimer disease (AD) and Huntington disease (HD)
(Graham et al. 2011) and zDHHC17 is known to also play
436
M.-E. ZABALLA AND F. G. VAN DER GOOT
a direct role in HD by mediating the S-acylation of huntingtin (Singaraja et al. 2002; Huang et al. 2004).
Cell polarity and migration
The apical-basolateral polarity is a key property of
epithelia to fulfill their barrier and communication
roles. The scaffolding protein Scribble (Scrib), which
localizes to cell-cell junctions, is involved in establishing and maintaining this polarity (Dow et al. 2003),
and mislocalizes to the cytosol in many human cancers (Vaira et al. 2011). Its plasma membrane localization and tumor suppression capacity depend on
the S-acylation of two conserved cysteine residues,
mostly mediated by zDHHC7 (Chen et al. 2016).
Scrib S-acylation appears to be highly dynamic
(Martin et al. 2012). Expression of the epithelial-tomesenchymal transcription factor Snail in benign epithelial cells resulted in a mislocalization of Scrib to
the cytosol and also led to repression of select
DHHC PATs and increased expression of APT2.
Inhibition of APT2 by the selective inhibitor ML349
efficiently drove membrane re-localization of Scrib in
this system (Hernandez et al. 2017).
In addition to modifying Scrib, zDHHC7 was found to
S-acylate junctional adhesion molecule C (JAM-C), promoting its localization to tight junctions and thereby
inhibiting cell migration (Aramsangtienchai et al. 2017).
Cell migration in turn was shown to depend on the Sacylation of the adhesion protein CD44 (Babina et al.
2014). Interestingly, non-canonical Wnt signaling was
reported to induce APT1-dependent S-deacylation of
CD44 and melanoma cell adhesion molecule (MCAM) in
melanoma cells thus promoting polarized protein localization and cell invasion (Wang et al. 2015).
Finally, Rho GTPases, key modulators of cell polarity,
adhesion and migration, have also been reported to
undergo S-acylation. Prenylated Rac1 is targeted to actin
cytoskeleton-linked ordered membrane regions through Sacylation and when this fails to occur it leads to defects in
cell spreading and migration (Navarro-Lerida et al. 2012).
S-Acylation of prenylated bCdc42 similarly leads to stable
membrane association (Nishimura and Linder 2013).
Interestingly, S-acylation of the atypical non-prenylated
Rho GTPases, RhoU and RhoV, leads to transient and
dynamic membrane association (Berzat et al. 2005;
Chenette et al. 2005). Finally, S-acylation of isoprenylated
RhoB leads to its targeting to multivesicular bodies and
degradation, and thereby to its inactivation (Perez-Sala
et al. 2009).
Innate immunity
The innate immune system represents the first line of
defense against infections by activating a variety of
mechanisms, including the production of interferons
(IFN). In this pathway, the ER receptor protein STING
(stimulator of interferon genes) is essential to initiate
effective IFN production and trigger the expression of
IFN-stimulated genes (Ishikawa et al. 2009). S-Acylation
of STING is required for its activation and for the subsequent type I IFN response. STING S-acylation appears to
occur in the late Golgi, primarily on two nearby cysteine
residues (Mukai et al. 2016). Remarkably, mutation of
these two cysteine residues prevented downstream signaling not only for wild-type STING but also for the
STING variants found in SAVI (STING-associated vasculopathy with onset in infancy) patients, where a gain-offunction mutation allows them to trigger the production of interferon independently of a stimulus thus
causing an autoinflammatory disease (Mukai et al.
2016). Thus, targeting S-acylation of STING represents
an interesting therapeutic strategy to treat autoimmune or inflammatory diseases. The Ablasser group
recently performed a small-molecule searching for
inhibitors of STING and identified a molecule that
indeed binds covalently to a transmembrane cysteine
residue and blocks its S-acylation maintaining the protein in a signaling-incompetent state. These STING
antagonists are able to attenuate pathological features
of autoinflammatory disease in mice highlighting the
potential of this therapy (Haag et al. 2018).
Among the interferon-induced proteins, IFITM proteins play critical roles in blocking the entry of a variety
of viruses, notably influenza A, dengue and hepatitis C
viruses (Brass et al. 2009; Bailey et al. 2014; Narayana
et al. 2015). IFITM3 inhibits viral infection by preventing
virus fusion with endolysosomal membranes, presumably by disrupting intracellular cholesterol homeostasis
and affecting membrane curvature (Amini-Bavil-Olyaee
et al. 2013; Chesarino et al. 2014b, 2017). S-acylation of
IFITM3 at three conserved cysteine residues promotes
antiviral activity, most probably by mediating efficient
membrane anchoring (Chesarino et al. 2014b, 2017).
Whereas multiple DHHC PATs were shown to be able to
S-acylate IFITM3 in human cells, zDHHC3, 7 and 20 had
the most striking effect (McMichael et al. 2017). SAcylation of IFITM proteins represents an interesting
case as it was found to be conserved in all tested IFITM
proteins but shown to have different effects on different
members of the family. As mentioned earlier, murine
IFITM1 S-acylation also regulates protein stability by preventing proteasomal degradation (Hach et al. 2013); and
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
S-acylation of IFITM5 was reported to promote its function as a bone formation factor (Tsukamoto et al. 2013).
Metabolism
Numerous recent articles support a link between protein S-acylation and cellular metabolism. In 2013, an
S-acylproteomics study on murine adipocytes revealed
more than 600 S-acylation candidates (Ren et al. 2013a).
In a follow-up study, the authors reported that
S-acylation of glucose transporter 4 (Glut4), mainly
mediated by zDHHC7, is critical for its insulin-dependent translocation to the plasma membrane (Ren et al.
2015; Du et al. 2017). Moreover, zDHHC7 knock-out
mice developed hyperglycemia and glucose intolerance. Strikingly, the authors observed that zDHHC7
activity (estimated from autoacylation levels) was
increased upon insulin treatment, illustrating that activity of DHHC S-acyltransferases can be regulated by
extracellular signaling (Du et al. 2017). The plasma
membrane-associated protein ClipR-59, involved in the
regulation of Glut4 translocation, was also reported to
depend on S-acylation, in this case primarily mediated
by zDHHC17 (Ren et al. 2013b). Importantly, zDHHC17
has been found as a candidate disease gene in Type 1
diabetes and shown to regulate glutamate-stimulated
insulin secretion and beta-cells survival (Berchtold et al.
2011). How the reported indirect and direct effects of
protein S-acylation on the localization and function of
the Glut4 transporter are coordinated remains to
be explored.
zDHHC13 has also been linked to metabolism and
mitochondrial function. In one study, zDHHC13-deficient mice were found to exhibit hypermetabolism and
defective lipid metabolism (Shen et al. 2017). Another
study focused on the zDHHC13-dependent S-acylation
of dynamin-related protein 1 (Drp1) and reported that
loss of zDHHC13 in cortex and cerebellum resulted in
altered mitochondrial dynamics accompanied by
increased glycolysis, glutaminolysis, and lactic acidosis
(Napoli et al. 2017).
A variety of studies revealed roles for protein S-acylation in lipid metabolism. The long-chain fatty acid
receptor CD36 plays important roles in lipid homeostasis and, not surprisingly, its expression is tightly regulated at the transcriptional, translational, and posttranslational levels, amongst which by S-acylation (Tao
et al. 1996). S-Acylation of CD36 was shown to affect its
half-life and regulate its trafficking through the secretory pathway (Thorne et al. 2010). S-Acylation of
ELMOD2 is required for its targeting to lipid droplets
and subsequent regulation of adipocyte triglyceride
437
lipase (ATGL) recruitment and triglycerides biosynthesis
(Suzuki et al. 2015). S-acylation of the ATP-binding cassette transporter (ABC) A1 (mostly mediated by
zDHHC8) is essential for its trafficking and function as
mutation of the target cysteine residues resulted in
reduced plasma membrane localization and reduced
ABCA1-mediated lipid efflux (Singaraja et al. 2009). A
general connection between S-acylation and metabolism has also been suggested after curation and combined analysis of published S-acylproteomics studies
(Blanc et al. 2015; Sanders et al. 2015).
Finally, Berthiaume et al. have proposed that S-acylation of mitochondrial enzymes, which, as mentioned,
might be non-enzymatic, could play a role in metabolic
regulation and energy homeostasis. In particular, they
observed enzyme inhibition by acylation of a cysteine
residue at the active site and proposed a possible metabolic crosstalk between fatty acids metabolism and
other pathways, such as amino acids catabolism or
ketogenesis (Stucki et al. 1989; Berthiaume et al. 1994;
Corvi et al. 2001; Kostiuk et al. 2008). In line with aforementioned observations, Kathayat et al. (2018) reported
that mitochondrial protein S-deacylation (as monitored
by the fluorescent probe mito-DPP2) is sensitive to the
levels of acyl-CoA thioesterase 11 (ACOT11). This suggests a connection between mitochondrial S-deacylase
activity (primarily mediated by APT1) and the metabolic
state of the cell, although no direct link with the S-acylation status of any mitochondrial enzyme has been so
far established (Kathayat et al. 2018). As mentioned
before, it is still unknown whether APT1 in mitochondria serves as part of a regulated enzymatic S-acylation/
deacylation cycle or it has a rather “repairing” function
removing non-enzymatically attached acyl chains, for
example at cysteine residues in the active site of matrix
mitochondrial enzymes. In the case of a metabolic
crosstalk, the regulation of APT1 activity in a coordinate
manner would be expected, although there is currently
no evidence to support this.
Cancer
Over the last decade, the expression and activity of various DHHC PATs have been reported to be altered in
numerous cancers and the connection between protein
S-acylation and cancer has been extensively reviewed
(Ducker et al. 2004; Greaves and Chamberlain 2014; YesteVelasco et al. 2014; Tian et al. 2015; Sharma et al. 2017;
Chen et al. 2017b; De and Sadhukhan 2018). The most
classic example of the link between S-acylation and cancer is the small GTPases Ras, and more specifically H-Ras
and N-Ras. Ras gain-of-function mutations have been
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M.-E. ZABALLA AND F. G. VAN DER GOOT
found in about 25% of all human cancers (Hobbs et al.
2016). The activity of H- and N-Ras requires S-acylation,
and deacylation is essential to downregulate signaling
(Swarthout et al. 2005; Misaki et al. 2010; Eisenberg et al.
2013; Lin et al. 2017; Pedro et al. 2017). Here we have
selected two additional, recently reported examples highlighting the potential therapeutic applications of targeting
protein S-acylation in cancer treatment.
The epidermal growth factor receptor (EGFR), an
oncogenic receptor tyrosine kinase (Normanno et al.
2006), was also shown to get S-acylated. Recently,
Bollu et al. (2015) and Ali et al. (2018) showed that
FASN-mediated S-acylation positively regulates EGFR
activity. Runkle et al. (2016), in contrast, reported
that zDHHC20-mediated S-acylation suppresses EGFR
signaling. Different consequences of S-acylation on
EGFR subcellular localization have been reported by
the three groups (Bollu et al. 2015; Runkle et al.
2016; Ali et al. 2018), possibly due to the different
mutational status of EGFR in each study (Ali et al.
2018). Interfering with EGFR S-acylation has been
suggested as a clinical approach to treat cancers
that are resistant to EGFR-targeted therapy (Runkle
et al. 2016; Rana et al. 2018). Independently of its
potential activity on EGFR, zDHHC20 has been
reported to play a role in cellular transformation and
its expression was shown to be significantly elevated
in several types of human tumors including colon,
prostate, ovarian, breast and kidney (Draper and
Smith 2010).
S-Acylation was recently found to protect against
skin cancer (Chen et al. 2017a). In a preliminary small
molecule screen searching for modulators of melanocortin-1 receptor (MC1R) red hair color variants, the
authors identified palmitic acid as an interesting candidate. MC1R is a G-protein-coupled receptor expressed
in melanocytes, where it mediates the cutaneous
responses to solar ultraviolet radiation, including melanin production and DNA repair. Following a series of
elegant experiments, Chen et al. found that MC1R
undergoes acylation at a C-terminal cysteine residue in
a reaction primarily mediated by zDHHC13, and that
this modification is essential for activating MC1R signaling. The S-acylation-deficient mutant of MC1R failed to
upregulate downstream genes and mice expressing this
variant had a similar coat color and skin pheomelanin
content to the MC1R loss-of-function mice (Chen et al.
2017a). Finally, enhancing MC1R S-acylation by overexpression of zDHHC13 or treatment with Palmostatin B
(a deacylases inhibitor) resulted in enhanced DNA
Figure 5. Mathematical modeling of S-acylation data. For a protein with two S-acylation sites, an open system with five possible
species (U: unfolded; C00: folded, non-acylated; C10 and C01: S-acylated in one or the other site; C11: doubly-acylated) is modeled
and the transitions between species can be represented using a set of ordinary differential equations. The model is fed with
quantitative experimental data, such as [35S]-cysteine/methionine pulse-chase or [3H]-palmitate incorporation, and iterative cycles
of calibration, prediction, and validation are followed until an adequate model is obtained. A model capturing the system allows
to deconvolute experimental data into the contribution of each species and thus to study the effect of each S-acylation event on
protein stability and function. OE: overexpression, KD: knock-down. See text for more details (see color version of this figure at
www.tandfonline.com/ibmg).
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
439
damage repair, tumor size reduction and/or extended
mice survival after prolonged UVB irradiation. These
approaches could thus provide a potential clinical prevention for melanoma in patients with MC1R red hair
color variants (Chen et al. 2017a).
Regulation and dynamics of S-acylation
An essential feature of protein S-acylation, which distinguishes it from most other types of lipidation, is its
reversibility. Thus, S-acylation is not simply a structural
static protein modification; it can be a dynamic, tightly
regulated post-translational modification (PTM) involved
in signaling and protein activation as is phosphorylation.
Despite decades of research in the field, our understanding of protein S-acylation regulation and dynamics is still
rudimentary.
In-depth understanding of S-acylation dynamics
requires considering that proteins in the cell can exist
as a mixed population of S-acylated and non-acylated
species. Furthermore, many proteins can get S-acylated
at two or more cysteine residues (Blanc et al. 2015)
and the modification of each of these residues can
have different consequences. Thus, a protein with two
S-acylation sites might exist in four different S-acylation
states or species, three S-acylation sites give rise to
eight possible species, and so on. The properties and
characteristics of the individual species cannot be
accessed experimentally. This is partly due to the fact
that there are so far no antibodies able to recognize
the S-acylation of a specific cysteine. Single cysteine
mutants may also be useful but the remaining cysteines
will still have two possible states, modified or nonmodified. Access to features of the individual species
can however be obtained by data-driven mathematical
models of the system. Let us assume a protein such as
calnexin having two S-acylation sites. The system is
modeled as an open system, where the protein is first
synthesized and folded, it can then be S-acylated on
one or the other site, and it can undergo deacylation at
one or the other site. The folded form can thus adopt
four S-acylation states. All species can undergo degradation. Each enzymatic reaction has its specific rate constants and the dynamic transitions between states/
species of the network can be represented by a general
model consisting of ordinary differential equations
(ODE) (Figure 5). The model is then calibrated using
data from a subset of experiments, which can include
[35S]-cysteine/methionine pulse-chase experiments and/
or kinetics of [3H]-palmitate incorporation or loss for
the wild-type protein and cysteine mutants, as well as
any other quantitative experimental data. Using
Figure 6. Schematic representation of zDHHC6 S-acylation cascade. zDHHC6 requires S-acylation for activation and subsequent
S-acylation of its different substrates, including transferrin receptor (TfnR), calnexin (CNX), and inositol 1,4,5-triphosphate (IP3)
receptor type 1 (IP3R1). zDHHC6 S-acylation is mediated by
zDHHC16 and S-deacylation is catalyzed by APT2. The latter is
susceptible to S-acylation and deacylation by a yet-to-be-identified zDHHC PAT and APT1, respectively, which, in turn, can
potentially require S-acylation for function (see color version of
this figure at www.tandfonline.com/ibmg).
optimization algorithms, families of parameters that
simultaneously fit the entire calibration dataset are
defined. These are subsequently used to predict a
second set of experiments, which are then compared
with the experimental results from these experiments
(which were not part of the calibration dataset). If predictions match the experimental results, the model is
considered to capture the system. If there is a mismatch, the model needs to be modified with additional
knowledge, which could for instance be that the protein can form dimers, or exist in different cellular compartments, etc. Iterative cycles between modeling and
experiments lead to an adequate model that can then
be interrogated. The modeling allows to deconvolute
each experimental kinetic curve into the contribution
to this curve of each of the species. This in turn allows
to determine for example: the turnover rate of the nonS-acylated versus fully S-acylated species, the rate of Sdeacylation of partially S-acylated species versus fully
S-acylated species, the relative distribution of species
under different cellular conditions, if relevant the activity of the different S-acylation species, etc. (Figure 5).
By combining experimental determination of protein S-acylation and its dynamics with mathematical
modeling by the Hatzimanikatis group, we have
increased our understanding of the dynamics of this
PTM and the complexity of its regulation (Dallavilla
et al. 2016; Abrami et al. 2017). Notably, it was found
that the presence of two contiguous S-acylation sites,
frequently seen in S-acylated transmembrane proteins
(Collins et al. 2017; Blanc et al. 2015), could act as an
440
M.-E. ZABALLA AND F. G. VAN DER GOOT
important regulator of S-acylation kinetics and protein
stability (Dallavilla et al. 2016; Abrami et al. 2017). More
specifically, we found that the presence of two neighboring juxtamembranous cysteines, when both sites are
modified, tends to render S-acylation irreversible, probably by triggering membrane imbedding of the cysteines leading to their inaccessibility to thioesterases.
We also found that S-acyltransferase activity can be
regulated by S-acylation, i.e. S-acylation can occur in
cascades, as phosphorylation or ubiquitination. We
found that zDHHC16 catalyzes the S-acylation of
zDHHC6 at three cysteine residues in its SH3_2 domain
and this S-acylation is required for the correct functioning of the enzyme and subsequent S-acylation of key
substrates in the endoplasmic reticulum, such as calnexin and transferrin receptor (Lakkaraju et al. 2012;
Senyilmaz et al. 2015; Abrami et al. 2017). While S-palmitoylation of one of the cysteines confers high activity
to the enzyme, it also targets it to the ER-associated
degradation pathway. Therefore, zDHHC6 must rapidly
be depalmitoylated by APT2 (Abrami et al. 2017). APT2
itself was found to undergo S-acylation at a conserved
cysteine residue in its N-terminus to be active (Kong
et al. 2013). Thus, the fact that DHHC S-acyltransferases
can be regulated by cycles of S-acylation/deacylation,
through enzymes that undergo similar cycles, leads to
rapid increase in the complexity of the regulatory network (Figure 6). Many members of the human DHHC Sacyltransferase family appear to undergo regulatory Sacylation (Table 2). Thus, it is important to consider
their possible regulation through S-acylation cascades.
Beyond S-acylation cascades, DHHC S-acyltransferases and acyl protein thioesterases can be subjected to
other complex regulatory schemes. In neurons, expression of human zDHHC9 and the subsequent membrane
trafficking of its substrate H-Ras have been reported to
be regulated by the microRNA-134 (in somatostatinpositive interneurons) (Chai et al. 2013), and microRNA138-dependent expression of the depalmitoylase APT1
has been linked to dendritic spine morphogenesis
through, at least partially, membrane localization of its
substrate G-protein a subunit a13 (Siegel et al. 2009).
Interestingly, H-Ras has also been reported to be a substrate of APT1 (Kong et al. 2013), and microRNA-134
and microRNA-138 appear to be differentially regulated
by neuronal activity (Siegel et al. 2009; Chai et al. 2013).
Yet, whether zDHHC9 and APT1 are regulated in a coordinated fashion remains unexplored. More recently,
APT1 and APT2 were shown to be under the control of
microRNA-138 and -424, respectively, and regulate cell
death through S-deacylation of Fas/CD95 in primary
chronic lymphocytic leukemia cells (Berg et al. 2015).
Dynamic S-acylation was also shown to be stimulated
by extracellular signaling. As mentioned before, zDHHC7
was reported to be activated upon insulin treatment in
HEK293 cells (Du et al. 2017). The abundance of zDHHC3
and S-acylation of its substrate d-catenin were increased
upon exposure of cultured dorsal root ganglion neurons
to the inflammatory cytokine TNF-a (Zhang et al. 2018).
Additionally, zDHHC3 enzymatic activity was reported to
be controlled by tyrosine phosphorylation mediated by
fibroblast growth factor receptor 1 (FGFR1) and Src kinases in neuroblastoma N2a cells (Lievens et al. 2016).
Notably, it was found that Src-mediated phosphorylation
of Tyr295 and Tyr297 has an inhibitory effect on zDHHC3
enzymatic activity while FGFR1-mediated phosphorylation of Tyr18 can overcome such inhibition (Lievens
et al. 2016). This study provides the first evidence of
DHHC PAT activity modulated by phosphorylation and
highlights, once more, that S-acylation enzymes can be
part of complex and dynamic signaling networks. Finally,
an additional regulatory role has been proposed for
dynamic changes in DHHC PAT localization (Greaves
et al. 2011; Lai and Linder 2013; Chamberlain and
Shipston 2015; Abrami et al. 2017).
Figure 7. Regulation and physiological relevance of protein
S-acylation. S-acylation can be regulated by affecting the
activity of the S-(de)acylation enzymes or by direct regulation
of the select S-acylation event of a given protein substrate or
cysteine residue. In turn, the molecular consequences of this
S-acylation can be varied and impact a broad spectrum of
physiological processes (see color version of this figure at
www.tandfonline.com/ibmg).
CRITICAL REVIEWS IN BIOCHEMISTRY AND MOLECULAR BIOLOGY
S-Acylation of a given protein can be modulated
independently of the regulation of S-(de)acylation
enzymes. Besides the previously described interplay
between S-acylation and ubiquitination (see above),
phosphorylation of several proteins can positively or
negatively regulate their S-acylation. While phosphorylation of phosphodiesterase 10A (PDE10A) and calnexin
was shown to act as a negative modulator of their Sacylation (Charych et al. 2010; Dallavilla et al. 2016), Sacylation of tissue factor (TF) was reported to negatively
regulate its phosphorylation (Dorfleutner and Ruf 2003).
In turn, the cross-talk between S-acylation and phosphorylation was shown to regulate BK potassium channels activity (Tian et al. 2008) and GAP43 plasma
membrane targeting (Gauthier-Kemper et al. 2014).
Other PTMs targeting cysteine residues, and thus acting
as potential competitors, have been reported to affect
protein S-acylation. S-Nitrosylation of PSD-95 was
shown to decrease its S-acylation (Ho et al. 2011), and
S-glutathionylation of H-Ras was reported to prevent its
S-acylation under specific metabolic stress conditions
(Burgoyne et al. 2012).
Thus, as schematized in Figure 7, the S-acylation of a
given protein can be modulated at multiple levels and
have diverse physiological effects. The activity of S(de)acylation enzymes can be regulated at the translational and post-translational level as well as by extracellular signaling; and the S-acylation of the target protein
might respond to the interplay with other PTMs. In
turn, S-acylation can impact protein structure, stability,
localization, and/or function, and have varied (patho)physiological consequences.
Concluding remarks
The last few years have witnessed major advances in
the field of protein S-acylation: technologies to study
this modification have drastically improved, many proteins for which S-acylation is key for their function have
been identified, structures of the enzymes involved in
acyl chain attachment and removal are becoming available, and promising probes and inhibitors are being
developed. These advances will allow the field to reach
the next level of understanding. Particularly exciting
will be the study of this modification in the context of
signaling cascades and considering its possible crosstalk with other regulatory events. Obtaining a more systems level of comprehension will allow the identification of feedback loops. Modeling approaches will be
particularly useful and the fact that DHHC PATs are
present in far lower numbers than kinases or ubiquitinating enzymes might provide a unique opportunity to
441
model the full system and obtain an understanding of
the full network of a dynamic post-translational modification. The coming years are also likely to see the
development of new drugs to control S-acyltransferases
and thioesterases, which will not only expand our basic
knowledge of this complex modification but could also
be of therapeutic use for a number of diseases with an
identified S-acylation component.
Acknowledgements
The authors thank Laurence Abrami, Oksana Sergeeva, and
Luciano Abriata for helpful discussions and critical reading of
this manuscript.
Disclosure statement
The authors declare no potential conflicts of interest.
Funding
This work was supported by the EPFL, the Swiss National
Science Foundation, the Swiss National Centre of
Competence in Research (NCCR) Chemical Biology, and the
European Research Council under the European Union's
Seventh Framework Programme (FP/2007–2013)/ERC Grant
Agreement No. 340260—PalmERa to F.G.v.d.G. M.E.Z. was a
recipient of a long-term postdoctoral fellowship from the
Human Frontier Science Program (LT000152/2014-L) and a
transition postdoctoral fellowship from the Swiss
SystemsX.ch initiative (TPdF 2013/143), evaluated by the
Swiss National Science Foundation.
ORCID
Marıa-Eugenia Zaballa
0613-9519
F. Gisou van der Goot
8522-274X
http://orcid.org/0000-0002http://orcid.org/0000-0002-
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