Article
Crystal Structures and Inhibition Kinetics Reveal a
Two-Stage Catalytic Mechanism with Drug Design
Implications for Rhomboid Proteolysis
Graphical Abstract
Authors
Sangwoo Cho, Seth W. Dickey,
a Urban
Sinis
Correspondence
surban@jhmi.edu
In Brief
Rhomboid proteases cut proteins in
membranes. Cho et al. provide insights
into the mechanism. They show that that
proteolysis proceeds through two states
and observe interactions that stabilize the
substrate intermediate. They also find
interactions that increase inhibitor
potency.
Highlights
d
Inhibition kinetics reveal two substrate-bound forms exist
during rhomboid catalysis
d
Structural analyses capture an enzyme-stabilized tetrahedral
substrate intermediate
d
A network of interactions at nonessential substrate sites
enhances inhibition 10-fold
d
Reversible not irreversible substrate-based inhibitors
potently block rhomboid action
Cho et al., 2016, Molecular Cell 61, 329–340
February 4, 2016 ª2016 Elsevier Inc.
http://dx.doi.org/10.1016/j.molcel.2015.12.022
Accession Numbers
5F5B
5F5D
5F5G
5F5J
5F5K
Molecular Cell
Article
Crystal Structures and Inhibition Kinetics Reveal
a Two-Stage Catalytic Mechanism with Drug Design
Implications for Rhomboid Proteolysis
a Urban1,*
Sangwoo Cho,1,2 Seth W. Dickey,1,2 and Sinis
1Department of Molecular Biology & Genetics, Howard Hughes Medical Institute, Johns Hopkins University School of Medicine, 725 North
Wolfe Street, Room 507 PCTB, Baltimore, MD 21205, USA
2Co-first author
*Correspondence: surban@jhmi.edu
http://dx.doi.org/10.1016/j.molcel.2015.12.022
SUMMARY
Intramembrane proteases signal by releasing proteins
from the membrane, but despite their importance,
their enzymatic mechanisms remain obscure. We
probed rhomboid proteases with reversible, mechanism-based inhibitors that allow precise kinetic analysis and faithfully mimic the transition state structurally. Unexpectedly, inhibition by peptide aldehydes
is non-competitive, revealing that in the Michaelis
complex, substrate does not contact the catalytic
center. Structural analysis in a membrane revealed
that all extracellular loops of rhomboid make stabilizing interactions with substrate, but mainly through
backbone interactions, explaining rhomboid’s broad
sequence selectivity. At the catalytic site, the tetrahedral intermediate lies covalently attached to the catalytic serine alone, with the oxyanion stabilized by unusual tripartite interactions with the side chains of
H150, N154, and the backbone of S201. We also visualized unexpected substrate-enzyme interactions at
the non-essential P2/P3 residues. These ‘‘extra’’ interactions foster potent rhomboid inhibition in living
cells, thereby opening avenues for rational design of
selective rhomboid inhibitors.
INTRODUCTION
Proteolysis inside the cell membrane lies at the regulatory core of
many pathways that are paramount to the health of a cell (Brown
et al., 2000; Chan and McQuibban, 2013; De Strooper and Annaert, 2010; Wolfe, 2009). Each of the four known families of intramembrane proteases continues to be implicated in diverse
pathologies, including Alzheimer’s disease, Parkinson’s disease,
cancer, malaria infection, hepatitis C virus maturation, tuberculosis virulence, and diabetes (Chan and McQuibban, 2013; De
Strooper and Annaert, 2010; Manolaridis et al., 2013; Urban,
2009). In contrast to soluble proteases, which are arguably the
best understood enzymes and among the most effective therapeutic targets (Drag and Salvesen, 2010), the catalytic mecha-
nisms of these membrane-immersed enzymes are incompletely
understood and have proved difficult to target effectively for
therapeutic benefit (Golde et al., 2013).
Inhibitors that chemically mimic intermediates in the reaction
pathway offer a powerful means to dissect the enzymatic mechanism of a reaction (Hedstrom, 2002). Kinetic analysis of inhibition can reveal how the reaction is ordered and/or functionally
organized, while structural analysis can identify the specific
atomic contacts that the enzyme forges to guide substrates
through the catalytic steps. However, this powerful strategy
has eluded the study of intramembrane proteolysis (Nguyen
et al., 2015); kinetic analysis of catalysis inside the membrane
has not been possible until only recently (Dickey et al., 2013;
Kamp et al., 2015). Moreover, inhibitor co-structures have
been achieved thus far only with rhomboid proteases, and
despite more than a dozen such high-resolution rhomboid-inhibitor structures in the Protein Data Bank (PDB) (Brooks and
Lemieux, 2013), all structural information is limited to irreversible
inhibitors. These agents form adducts that distort the active site
and thus offer limited insights into the reaction mechanism and
do not permit kinetic analysis. In fact, no reversible inhibitors of
any kind have yet been developed for studying rhomboid proteolysis (Nguyen et al., 2015).
For serine proteases such as rhomboid, the ‘‘committed step’’
for proteolysis under physiological conditions is formation of the
covalent acyl intermediate following nucleophilic attack by the
catalytic serine; once a substrate reaches this step, it is destined
to complete the cleavage reaction (Hedstrom, 2002). Peptides
with a C-terminal aldehyde moiety bind as substrates and, after
attack on the terminal aldehyde carbon by the protease, arrest
the reaction. The resulting catalytic complex faithfully mimics
the key high-energy tetrahedral transition state that must be
stabilized by the enzyme for catalysis to proceed (Hedstrom,
2002).
Peptide aldehydes offer several notable advantages for interrogating the enzymatic mechanism of rhomboid proteases. First,
because peptide aldehydes resemble substrates precisely and
inhibit the protease reversibly, they allow detailed kinetic analysis. Structurally, because the only reaction is between the
serine nucleophile and a true carbonyl, peptide aldehydes avoid
the unnatural alkylation of the catalytic histidine base that besets
isocoumarins or chloromethylketones, which ultimately distort
the active site by crosslinking the catalytic serine and histidine
Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc. 329
Figure 1. Inhibition Kinetics of Peptide Aldehydes on Intramembrane Proteolysis by GlpG
(A) Mechanisms of GlpG inhibition (inhibitor warheads are in red). Nucleophilic attack by the serine oxygen of rhomboid (blue) results in a covalent tetrahedral
intermediate that differs only at one substituent between aldehydes (hydrogen, red) and natural substrates (black). Both produce an identical oxyanion (pink) that
rhomboid must stabilize through electrophilic catalysis. Inhibitor adducts that deviate from natural proteolytic intermediates are shaded with red ovals (see text).
(B) Summary of the 11 substrate peptides used in this study (Ac denotes acetylation, CHO is an aldehyde, NH2 is an amide, and CMK is a chloromethylketone).
(C) Comparison of GlpG inhibition by various peptides. GlpG was assayed in real time while reconstituted in liposomes formed from E. coli lipids, and peptide
aldehydes were added to proteoliposomes when the proteolytic reaction was initiated (without pre-incubation). vo and vi denote initial rates in the absence and
presence of inhibitors, respectively. Only Ac-RKVRMA-CHO inhibition was modeled by including an inaccessibility parameter (see Figure S1A).
(D) Kinetics of inhibition by Gurken peptide aldehydes was non-competitive (also see Figure S1B). Left: initial reaction rates versus substrate concentration in the
presence or absence of peptide aldehydes are plotted. Right graph compares the resulting KM and Vmax parameters under different inhibitor concentrations. Note
that for competitive inhibition, KM would increase in response to inhibitor concentration, while Vmax would be unaltered. For non-competitive inhibition, KM would
be unaffected, but Vmax would decrease in response to inhibitor concentration. p values of parameter differences relative to no inhibitor controls are shown above
each bar.
residues (Figure 1A). The atom being attacked during catalysis
with peptide aldehydes is a carbonyl carbon that generates a
true oxyanion, unlike with phosphonates or sulfonylfluorides
(Powers et al., 2002), in which the oxygen is not negatively
charged and is connected to a non-carbon atom (Figure 1A).
Finally, using peptide aldehydes overcomes the naturally low
affinity of rhomboid for substrates by stabilizing the natural covalent attachment step that follows serine attack, while observing
covalent linkage provides assurance that substrate had adopted
a catalytically competent conformation.
330 Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc.
We recently developed an inducible, real-time assay capable
of quantifying the kinetics of proteolysis directly inside the membrane (Dickey et al., 2013). Therefore, to gain a better understanding of the mechanism underlying intramembrane catalysis,
we sought to adapt this approach to characterize the inhibition
kinetics of substrate-mimicking peptide aldehydes. Next,
because mounting evidence indicates that cell membranes
affect the properties of rhomboid proteases beyond just serving
as their environment (Bondar et al., 2009; Moin and Urban, 2012;
Urban and Moin, 2014; Vinothkumar, 2011), we developed conditions to crystallize a catalytically active rhomboid protease in a
membrane. Soaking rhomboid crystallized from bicelles with the
most potent substrate peptide aldehydes produced high-resolution structures that revealed the characteristics of substrate stabilization during catalysis by a membrane-immersed rhomboid
protease. Finally, we unexpectedly visualized ‘‘extra’’ substrate-enzyme interactions that we subsequently found are
able to drive rhomboid inhibition to completion in living cells,
thereby opening avenues for the rational and selective design
of rhomboid inhibitors.
RESULTS
Kinetic Analysis of Peptide Aldehyde Inhibition
Because the kinetics of inhibition have not been studied within
the membrane, our first goal was to adapt our inducible reconstitution assay to study the inhibition kinetics of peptide aldehydes
directly within the membrane. We therefore modeled a series of
peptide aldehydes (Figure 1B) on three well-studied rhomboid
substrates (Strisovsky et al., 2009; Urban et al., 2002), namely,
TatA, Spitz, and Gurken, and examined their ability to inhibit
GlpG, the rhomboid protease from Escherichia coli, reconstituted within the membrane. A key feature of our approach is
the fact that both substrate and peptide aldehydes engage the
active protease together at the start of the reaction (without
needing to pre-incubate rhomboid with inhibitor as in prior
studies).
Although most peptide aldehydes had Ki values in the millimolar range, Gurken hexapeptide and tetrapeptide aldehydes
proved to be more potent inhibitors of GlpG (Figure 1C). The
Gurken tetrapeptide aldehyde Ac-VRMA-CHO exhibited a Ki of
113 ± 4 mM, and there was a further increase in potency with
the hexapeptide aldehyde Ac-RKVRMA-CHO. As anticipated,
inhibition required the aldehyde moiety, because replacing it
with an amide raised Ki by >20-fold. This is consistent with rhomboid displaying weak affinity for substrates (Dickey et al., 2013)
and forming a reversible covalent product with peptide
aldehydes.
Contrary to expectation, the mode of inhibition proved to
be non-competitive (Figure 1D), with no significant change in
KM (p = 0.16–0.80) but a strong and dose-dependent decrease
in Vmax (p = 2.1 3 109 to 2.0 3 1013). This was also clearly reflected in the residuals when we compared fitting the kinetic data
to a competitive versus a non-competitive model. Moreover, we
observed non-competitive inhibition with tripeptide (Ac-RMACHO), tetrapeptide (Ac-VRMA-CHO), and hexapeptide (AcRKVRMA-CHO) aldehydes (Figure S1) and with Providencia
stuartii AarA (a rhomboid protease with seven transmembrane
segments that shares <15% identity with E. coli GlpG). Taken
together, these observations suggest that non-competitive inhibition is the mechanism by which peptides based on substrate
sequences preceding the cleavage site inhibit rhomboid
proteases.
Crystallization of an Active Rhomboid Protease in a
Membrane
We next sought to reveal the atomic basis of how peptide aldehydes interact with rhomboid as it catalyzes proteolysis in the
membrane environment. Prior work established that an inactive
mutant of E. coli rhomboid GlpG, whose structure has been studied extensively (Brooks and Lemieux, 2013), can be crystallized
from a membrane bicelle (Figure 2A) (Faham and Bowie, 2002;
Vinothkumar, 2011). Although such an inactive mutant prohibits
studying catalysis, this strategy provided a starting point for
crystallizing an active rhomboid in a membrane. We therefore
searched our bank of >200 GlpG mutants for those that maintain
activity and structural stability (Baker and Urban, 2012) and ultimately succeeded in crystallizing an active GlpG variant (Y205F)
in a membrane (Figure 2B). Notably, phenylalanine at position
205 occurs naturally in many rhomboid orthologs (Bondar
et al., 2009), had no effect on the structural stability of GlpG (Figure 2C), and slowed proteolytic rate less than 2-fold (Figure 2D).
This is a very mild effect, because the natural variation in catalytic
rate between rhomboid orthologs ranges 10,000-fold (Dickey
et al., 2013).
The resulting structure revealed a gate-open conformer with
lateral displacement of the entire TM5 gating helix and fully disordered overlying L5 Cap loop, while the rest of the protein was unchanged relative to the closed state (Figure 2B). We next used
this open conformer of rhomboid in a membrane environment
as an excellent starting point for visualizing substrate binding
and catalysis.
Structure of Rhomboid in Catalytic Complex with
Peptide Aldehydes
To form the catalytic complex, we reacted GlpG in the open
conformation crystallized from a membrane bicelle with our
most potent Gurken peptide aldehydes. After iterative optimization, we ultimately succeeded in collecting four data sets that
differed in reaction conditions and ranged from 2.2–2.4 Å in resolution and solved the resulting structures using molecular
replacement (Figure 3A; Table 1).
Globally, only minor changes in the protease accompany substrate binding and catalysis (Figure 3B). The greatest changes
occurred in the extracellular L5 loop to the top of TM6. In the
open conformer, the distal part of the L5 Cap became ordered,
while TM5 maintained its open form (Figure 3B). The small size
of our peptide aldehydes also allowed us to form complexes
with gate-closed GlpG, which defined the minimal movements
necessary for catalysis (Figure 3C). When we used the gateclosed form as a starting point, residues of the extracellular L5
loop and extending to the top of TM6 were displaced outward
by substrate (Figure 3C). Excluding these 13 residues, the
root-mean-square deviation (rmsd) along the remaining 158
alpha carbons of GlpG was only 0.65 Å. Moreover, the catalytic
residues themselves maintained positions that were very similar
Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc. 331
Figure 2. Structure of an Active, Open
Rhomboid Protease in a Membrane
(A) Schematic of E. coli rhomboid GlpG in a DMPC/
CHAPSO membrane bicelle. The short-chain
CHAPSO seals the edges of the DMPC membrane.
(B) Structure of an open E. coli GlpG Y205F (yellow)
in a membrane overlaid with a gate-closed form
(red, 2IC8 crystallized from detergent). Left is topdown view (membrane in plane of the page), middle is a lateral view (dashed lines indicate
approximate location of the membrane), and right
are distance measurements between landmark
TM5 and TM2 residue alpha carbons (beads). In
the membrane environment, TM5 has shifted up
and away from TM2.
(C) Thermostability analysis of GlpG with tyrosine
versus phenylalanine at position 205.
(D) Real-time Michaelis-Menten kinetic analysis of
wild-type versus Y205F GlpG reconstituted in
liposomes formed from E. coli lipids. Inset
compared kinetic parameters of Y205F relative to
wild-type. Kinetic parameters are represented as
mean ± SEM.
to their arrangement in the apoenzyme (Figure 3D). These minor
changes reveal that the apoenzyme monomer observed in most
structures is in the catalytically active form, although it must be
realized that natural substrates are much longer, and greater
motions away from the active site undoubtedly are required to
accommodate intramembrane proteolysis normally.
The Tetrahedral Intermediate
At the active site, the electron density of catalytic S201 was
continuous with the substrate alanine aldehyde residue, indicating that covalent catalysis indeed took place (Figures 3A
and 3E). Interestingly, catalytic H254 moved slightly but still
made a hydrogen bond to S201 (at distances of 2.85 and
3.11 Å in the tripeptide and tetrapeptide structures, respectively). The aldehyde carbon was tetrahedral, thus mimicking
the transition state and directly revealing the enzyme contacts
that stabilize it during catalysis.
Nucleophilic attack of the peptide carbonyl by the serine generates a negatively charged oxygen, the oxyanion (Figure 2A),
that must be stabilized in the transition state for catalysis to proceed (Henderson, 1970). This key electrophilic catalysis feature
of rhomboid proteases has been difficult to define, because prior
studies used diverse non-peptidic inhibitors that suggested a series of possible interactions and at various distances (Vinothkumar et al., 2010, 2013; Xue et al., 2012; Xue and Ha, 2012); initial
structures with isocoumarins suggested backbone interactions
with L200 and S201 and weak (3.4 Å distance) side-chain interactions with H150 and N154, while the most recent b-lactam and
chloromethylketone structures have the oxygen moiety positioned outside the oxyanion hole (Vinothkumar et al., 2013; Zoll
et al., 2014). Our complex structure with a true peptide oxyanion
now resolves these discrepancies (Figure 3E): the oxyanion hole
of rhomboid is ultimately tripartite, being
formed by the side chain of H150 (2.69 Å
distance), amide backbone of catalytic
S201 (2.91 Å distance), and the side chain of N154 (3.04 Å
distance).
Extended Substrate Interactions
Beyond the site of catalysis, the substrate makes a network of
interactions with the enzyme, some of which explain previously
enigmatic observations. We recently engineered and quantified
the effect of 200 mutants on both rhomboid structural stability
and protease activity (Baker and Urban, 2012). Residues in the
third extracellular loop (L3) in particular are generally not
conserved and have little effect on enzyme architecture, yet
dramatically compromise proteolytic activity (Baker and Urban,
2012). Although the basis for this was unclear, the substrateenzyme structure now reveals that L3 makes a collection of close
backbone interactions with the substrate (Figure 4A), the geometry of which is most likely perturbed by even subtle side-chain
alterations in L3.
In fact, most enzyme-substrate interactions are backbone
interactions (Figure 4A), explaining why rhomboid proteases
display rather broad sequence selectivity (Akiyama and
Maegawa, 2007; Moin and Urban, 2012; Strisovsky et al.,
2009). A total of five hydrogen bonds formed between the
substrate backbone and two enzyme loops: three with the inner L3 loop and two with the overlying L5 loop. Moving along
the substrate from the site of catalysis (in the substrate
termed P1 and moving sequentially outward from the aldehyde
to P4), the five sites of contact are P1 NH with CO of G198
(L3), P2 CO with NH of A250 (L5), P2 NH with CO of S248
(L5), P3 CO with NH of G198 (L3), and P3 NH with CO of
W196 (L3).
A number of restrictions have been mapped in substrate side
chains that rhomboid proteases can accommodate, the
332 Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc.
Figure 3. Conformation of Rhomboid Protease GlpG in Catalytic Complex with Substrate Peptide Aldehydes
(A) Electron density map (2Fo–Fc at 2s) of peptide aldehydes (in cyan and red) in the active site of GlpG in the open and closed forms.
(B) Top view of overlaid open GlpG with (yellow) and without (blue) substrate bound reveals changes in conformation. Note that residues 248–252 in the L5 loop
were structured around substrate (arrow).
(C) Top view of overlaid gate-closed GlpG with (yellow) and without (blue) substrate bound reveals minimal global changes in conformation.
(D) Nucleophilic and electrophilic catalysis residues display minor positional changes between apoenzyme (blue) and substrate-bound (yellow) forms.
(E) The substrate oxyanion (red mesh) is stabilized (dashed green lines) by the side-chain nitrogens of H150 (at 2.69 Å) and N154 (at 3.04 Å), as well as the
backbone nitrogen of S201 (at 3.3 Å). Shown in mesh is the electron density map (2Fo–Fc at 2s).
structural basis of which is now revealed in the substrateenzyme complex. In particular, rhomboid enzymes normally
cleave only after small side chains (Akiyama and Maegawa,
2007; Moin and Urban, 2012; Strisovsky et al., 2009). However,
the methyl group of the alanine points into a surprisingly large
cavity (Figure 4B), part of which we previously proposed forms
a water-retention site (Zhou et al., 2012). All three waters were
unperturbed upon substrate binding, and the substrate-enzyme
interaction at this restricted P1 site was surprisingly loose,
again consistent with low affinity for substrates (Dickey
et al., 2013). Yet changing the P1 alanine to valine alone
severely compromised peptide aldehyde inhibition of GlpG
(Figure 4C).
Another side-chain preference has been found at P4 (Strisovsky et al., 2009), where some, but not all (Moin and Urban, 2012;
Parussini et al., 2012), rhomboid enzymes prefer large hydrophobic residues. The substrate P4 valine formed hydrophobic interactions with F146 and M120 on the L1 loop (Figure 4D). However,
this too was a rather shallow interaction, and reacting GlpG with
a tripeptide missing the P4 residue entirely produced a nearly
identical substrate conformation (Figure 4E). This observation
suggests that the P4 residue has limited impact on GlpG catalysis. Indeed, mutation of large substrate residues at both P4
and P20 (I5A+F10A) that dramatically reduce proteolysis by
100-fold with its natural rhomboid protease P. stuartii AarA
(Dickey et al., 2013) reduced E. coli GlpG cleavage by only
2-fold (Figure 4F). Even providing a large phenylalanine at the
P4 residue did not enhance peptide aldehyde inhibitor potency
against GlpG (Figure 4G).
Moving further outward, we extended our structural analysis to
Gurken hexapeptide aldehydes (Figure 4H). However, the N-terminal P5 and P6 residues were disordered in our structure, and
their presence did not alter the position of P1–P4, arguing that
GlpG does not contain specific substrate-interacting subsites
beyond P4.
Unexpected Stabilizing Interactions at P2 and P3
Particularly surprising were interactions made by the P2
and P3 side chains, because these positions were thought to
be completely unrestricted in substrates (Akiyama and Maegawa, 2007; Moin and Urban, 2012; Strisovsky et al., 2009).
Indeed, both side chains pointed upward and out from the
active site (Figure 5A), explaining the apparent lack of restriction at these substrate positions. Nevertheless, the P3 arginine
of Gurken in particular made ‘‘extra’’ contacts (Figure 5B); its
guanidinium group made a series of interactions with the top
surface of the L5 loop, including directly with the CO of
the M247 backbone, as well as the side chains of S248,
N251, and S193 (on L3), potentially through bridging water
molecules.
Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc. 333
Table 1. Crystallography Statistics
GlpG + VRMA
GlpG-Y205F
Bicelle
GlpG-Y205F
Bicelle +RMA
GlpG-Y205F
Bicelle +VRMA
GlpG-Y205F
Bicelle +RKVRMA
R32
C222(1)
C222(1)
C222(1)
C222(1)
110.6
71.2
70.8
70.6
70.3
110.6
98.1
97.5
98.0
96.1
126.7
63.0
62.8
62.6
62.7
44.80–2.00
(2.05–2.00)
42.56–2.20
(2.24–2.20)
57.51–2.20
(2.27–2.20)
62.55–2.30
(2.39–2.30)
56.75–2.30
(2.38–2.30)
Data Collection
Space Group
Unit cell (Å)
a, b, c
Resolution (Å)a
Observations
159,585
89,344
55,382
50,063
48,096
Unique reflectionsa
19,181
10,370
10,422
9,921
9,474
Multiplicity
7.9 (8.0)
6.8 (6.1)
5.3 (5.2)
5.0 (5.1)
5.1 (5.2)
I/s(I)
8.2 (1.9)
6.9 (1.9)
6.7 (1.8)
5.7 (1.9)
5.8 (2.4)
Completeness (%)a
99.3 (100.0)
98.4 (74.1)
91.3 (94.0)
99.8 (99.7)
97.6 (97.2)
Rmergea
0.113 (0.443)
0.135 (0.437)
0.138 (0.739)
0.190 (0.571)
0.169 (0.522)
22.40–2.30
(2.36–2.30)
42.56–2.50
(2.56–2.50)
50.00–2.30
(2.36–2.30)
50.01–2.40
(2.46–2.40)
50.01–2.4
(2.46–2.40)
Refinement
Resolution (Å)a
No. of reflections
12,636
7,905
8,921
8,287
7,947
Rwork/Rfree
0.215/0.263
(0.289/0.332)
0.222/0.287
(0.244/0.397)
0.209/0.250
(0.265/0.332)
0.235/0.263
(0.318/0.343)
0.237 /0.286
(0.357/0.408)
No. of atoms
1,526
1,520
1,487
1,504
1,499
Protein
1,482
1,467
1,464
1,475
1,449
Water
44
53
23
29
50
Average B factors (Å2)
54.8
50.6
45.8
39.4
50.6
Protein
54.8
50.3
44.1
37.8
48.4
Water
52.6
55.4
47.5
41.0
52.6
Bonds (Å)
0.018
0.012
0.008
0.012
0.011
Angles ( )
1.89
1.37
1.10
1.38
1.32
Rmsd
Ramachandran plot
Outliers (%)
0.6
0.6
1.1
0.52
0.7
Favored (%)
95.5
95.0
95.5
93.1
90.7
a
Values in parentheses correspond to highest resolution shell. Rfree was calculated over reflections in a test set not included in the atomic refinement:
GlpG-VRMA, 4.9%; Y205F, 5.2%; Y205F-RMA, 4.8%; Y205F-VRMA, 5.1%; and Y205F-RKVRMA, 4.8%.
To evaluate the contribution of these side-chain interactions
to affinity, we mutated the corresponding residues individually
in GlpG to alanine and quantified the ability of the Gurken
tetrapeptide aldehyde to inhibit the GlpG variants (Figure 5C).
The N251A mutant alone proved to be 3-fold less sensitive to
peptide aldehyde inhibition (p = 6.4 3 1011), while S248A
conferred a mild (1.5-fold) but significant (p = 0.00015)
reduction in inhibition. Even mutating M247, which makes a
backbone interaction with the arginine side chain, increased
Ki by 2-fold (p = 5.4 3 107), perhaps by altering conformation of the L5 loop. In contrast to these L5 residues,
mutating the L3 residue S193 that may also make a distant
interaction with the P3 arginine did not affect inhibition
(p = 0.51).
Consistent with a network of interactions between GlpG and
the P3 residue, combining these GlpG loop mutants proved to
be synergistic (Figure 5D). The S248A+N251A double mutant
became 6-fold less sensitive to inhibition, while further incorporating S193A into a triple mutant decreased inhibition by
7.6-fold. As such, residues within the L5 loop provide multifaceted stabilizing interactions with the P3 side chain, with additional minor contributions offered by other nearby residues,
including S193.
To evaluate the functional consequences of these interactions
in substrate processing in living cells, we made the corresponding R243A mutant in full-length Gurken and found that even this
single mutant reduced processing of Gurken in living E. coli cells
by 2-fold (Figure 5E). These unanticipated ‘‘extra’’ interactions
may thus explain ‘‘sequence contexts’’ that lead to some
substrates being cleaved more efficiently than others (Akiyama
and Maegawa, 2007; Moin and Urban, 2012; Strisovsky et al.,
2009).
334 Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc.
Figure 4. An Interaction Network Stabilizes the Substrate in the Rhomboid Active Site
(A) Backbone interactions between the extended peptide substrate (cyan) and the L5 (left) and L3 (right) loops of GlpG stabilize the catalytic complex.
(B) Magnified view of the P1 substrate alanine side chain in the S1 and water-retention site (shaded), showing the three bound water molecules (red spheres).
(C) Inhibition kinetics of Gurken tetrapeptide aldehydes with P1 alanine versus valine.
(D) The P4 side chain forms hydrophobic interactions with M120 and F146 of the L1 loop as an S4 pocket (shaded).
(E) Overlay of substrate conformation in the GlpG active site with (tetrapeptide in cyan) and without (tripeptide in gray) the P4 valine.
(F) Kinetics of P. stuartii AarA versus E. coli GlpG on TatA harboring alanines at both P4 and P20 (I5A+F10A) in reconstituted liposomes formed from E. coli lipids.
(G) Inhibition kinetics of the indicated P4 Gurken tetrapeptide aldehydes.
(H) Overlay of the tetrapeptide (gray) and hexapeptide (orange) substrate conformation in the GlpG active site. Shown in mesh is the experimental electron density
map (2Fo–Fc at 2s), revealing that the P5 and P6 residues (hypothetically drawn in ball and stick) were disordered and thus not making specific contacts
with GlpG.
Finally, because the rhomboid:substrate interactions that we
visualized at P3 had an effect even on processing of a fulllength substrate, we examined whether these interactions
play a particularly important role in achieving inhibition of
rhomboid by small molecules. In agreement with this postulate, we found that a Gurken tetrapeptide aldehyde with just
its P3 residue changed to alanine, as occurs naturally in the
TatA substrate, was 10-fold less potent at inhibiting GlpG (Figure 5F). As such, our structural and kinetic observations reveal
how contacts at ‘‘unrestricted’’ positions can make ‘‘extra’’
stabilizing interactions that dramatically increase inhibitor
potency.
Peptide Aldehydes Are Potent and Specific Rhomboid
Inhibitors in Living Cells
Our serendipitous discovery potentially offers a strategy for the
rational design of small-molecule rhomboid inhibitors that are
effective in living cells. Prior groundbreaking high-throughput
screens of >58,000 compounds using detergent-solubilized
rhomboid enzymes identified a series of b-lactams that proved
to be potent inhibitors in vitro but failed to block endogenous
GlpG activity completely in E. coli cells even when pre-incubated
with cells prior to inducing substrate expression (Pierrat et al.,
2011). We therefore examined whether our peptide aldehydes
were effective in vivo by examining their activity against endogenous GlpG in the same E. coli strain.
Adding Ac-VRMA-CHO to growing E. coli cells resulted in a
complete block to TatA-Flag processing by GlpG, and this block
did not require pre-incubating cells with inhibitor prior to
inducing substrate expression (Figure 6A). As expected, inhibition relied on the aldehyde moiety, because converting it to an
amide in Ac-VRMA-NH2 abolished inhibition completely. Even
the apparent half maximal inhibitory concentration (IC50) of AcVRMA-CHO (98 ± 11 mM) was indistinguishable from the
Ki (113 ± 4 mM) that we measured using our proteoliposome system (Figure 6B), indicating that proteoliposomes are an effective
mimic of GlpG in the natural setting of the E. coli membrane.
We next examined whether the ‘‘extra’’ elements we found to
be important for inhibition in vitro are required for blocking GlpG
activity in living cells (Figure 6C). Importantly, substituting the P3
arginine with alanine essentially abolished inhibitor efficacy,
again indicating that, in addition to the much studied P4 and
P1 residues, ‘‘extra’’ interaction at neglected residues can be
very effective at driving inhibitor potency.
Contrary to expectation, however, we discovered that converting the aldehyde moiety to a chloromethylketone warhead
in our tetrapeptides actually compromised inhibitor potency by
nearly an order of magnitude (Figure 6C). To date, all rhomboid
Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc. 335
Figure 5. Unanticipated P3 Residue Interactions and Their Implications on Substrate Cleavage and Rhomboid Inhibition
(A) The P2 (methionine) and P3 (arginine) residue side chains point out from the active site. Shown (and in B) is Ac-VRMA-CHO bound to GlpG, with dashed lines
denoting hydrogen bonds.
(B) The guanidinium group of the P3 substrate arginine side chain makes a series of ‘‘extra’’ stabilizing contacts, including hydrogen bonding (dashed line) directly
with the backbone CO of M247 at a distance of 3.06 Å, and potentially with the hydroxyl side chains of S193, S248, and/or N251 (perhaps via water-mediated
interactions).
(C) Inhibition kinetics of Gurken tetrapeptide aldehyde Ac-VRMA-CHO quantified against four single GlpG loop mutants.
(D) Bar graph comparing the Ki of Ac-VRMA-CHO with eight different GlpG loop variants (wild-type, single, double, and triple mutants are shown, with p values
relative to wild-type above each bar).
(E) Mutating the P3 arginine in Gurken (R243A) alone reduced cleavage 2-fold in E. coli cells.
(F) Inhibition kinetics of Gurken tetrapeptide aldehydes with arginine versus alanine at the P3 position.
inhibitors are irreversible and form covalent adducts with the
protease (Nguyen et al., 2015), which is thought to increase inhibitor potency. Instead, our analysis indicates that the key
element for achieving effective inhibition of rhomboid in vivo is
a reversible warhead.
Finally, although we did not set out to develop a potent inhibitor, even our limited series of peptide aldehydes already yielded
an inhibitor that was effective at blocking endogenous GlpG
activity in living E. coli cells with an IC50 in the low micromolar
range: Ac-RKVRMA-CHO displayed an IC50 of 2.9 ± 0.3 mM (Figure 6D). These early observations provide proof of principle for
using reversible warheads and cultivating ‘‘extra’’ interactions
at nonessential substrate positions as a promising strategy for
developing rhomboid inhibitors.
DISCUSSION
In conclusion, we integrated kinetic analysis of rhomboid inhibition by substrate-mimicking peptide aldehydes with highresolution X-ray crystallography to interrogate the enzymatic
mechanism of rhomboid proteases. An important advance is
our ability to study rhomboid catalysis with reversible inhibitors
and entirely inside the membrane (both kinetically and crystallographically), which is known to affect the catalytic properties of
these membrane-immersed enzymes (Bondar et al., 2009;
Moin and Urban, 2012; Urban and Moin, 2014; Vinothkumar,
2011). Ultimately, we identified an additional, early step in the kinetic scheme for rhomboid proteolysis, visualized several key
features of rhomboid catalysis at atomic resolution, and, unexpectedly, discovered a promising alternative avenue for targeting these enzymes.
Inhibition Kinetics Identify Two Distinct SubstrateBound Complexes
Particularly unusual but very informative is the non-competitive
mode of inhibition by substrate-mimicking peptide aldehydes,
because it reveals that rhomboid proteases exist in two distinct
substrate-bound forms (Figure 7). To result in non-competitive
inhibition, peptide aldehydes must, by definition, be able to
bind to the substrate-enzyme Michaelis complex. Importantly,
in addition to structural analysis, we further validated the catalytic center as the binding site that leads to this non-competitive
336 Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc.
Figure 6. Peptide Aldehydes Inhibit Endogenous GlpG in Living E. coli Cells
(A) Western analysis (anti-Flag) of TatA-Flag cleavage by endogenous GlpG in cells cultured in the presence or absence of 1 mM tetrapeptides Ac-VRMA-NH2
(NH2) or Ac-VRMA-CHO (CHO). In all panels, D indicates a control strain in which the endogenous GlpG gene was deleted, while ± indicates whether expression
of the TatA-Flag substrate was induced or not induced. The first (and in C last) lane of each gel image shows cells in which substrate expression was not induced.
(B) Quantification of inhibition by Ac-VRMA-CHO of endogenous GlpG in living cells (blue line) or pure GlpG reconstituted in proteoliposomes (black line).
(C) Western analysis (anti-Flag) of TatA-Flag cleavage by endogenous GlpG in cells cultured in the presence or absence of the indicated tetrapeptides (CMK
denotes chloromethylketone).
(D) Western analysis (top, anti-Flag) and quantification (lower graph) of inhibition by Ac-RKVRMA-CHO of endogenous GlpG in living cells.
inhibition by engineering ‘‘resistant’’ mutants in GlpG that
compromised peptide aldehyde inhibition. Therefore, in the Michaelis complex, substrate does not contact the catalytic center.
Non-competitive inhibition exhibited by substrate peptide
aldehydes reveals what is likely to be the organizing feature underlying rhomboid proteolysis. We previously speculated that,
by analogy to DNA glycosylases (Friedman and Stivers,
2010), rhomboid proteases could exist in two forms: the interrogation complex and the scission complex (Dickey et al.,
2013). Initial binding of a substrate transmembrane segment
laterally in the membrane to a gate-open rhomboid would
form the interrogation complex, which in kinetic terms is the Michaelis complex. At this point, a ‘‘pause’’ would give those
transmembrane segments that are predisposed to unwind an
opportunity to extend toward the inner site of catalysis, while
those that do not unwind could not be cleaved and are ultimately ejected (Dickey et al., 2013; Moin and Urban, 2012).
This has functional implications, because it gives rhomboid a
separate step for discriminating substrates from non-substrates, but this step would render the substrate-bound Michaelis complex nevertheless sensitive to peptide aldehyde
attack. The non-competitive mode of inhibition now provides
direct experimental evidence for two such functionally separate
states of the protease.
The kinetic scheme for rhomboid proteolysis has, therefore, a
step in which a conformation change converts the Michaelis
complex to a catalytic complex. This is an important distinction,
because it corrects a recent study with peptide chloromethylketones bound at the catalytic center that was assumed to be the
Michaelis complex (Zoll et al., 2014). This was an entirely reasonable assumption at the time, because the kinetic mode of inhibition could not be determined in that study because rhomboid
had to be pre-incubated with the irreversible inhibitors for 3 hr
prior to adding substrate.
We next used X-ray crystallography to visualize the catalytic
features of the scission complex itself, namely, covalent linkage
between substrate and nucleophilic serine, the tetrahedral intermediate formed by substrate, electrophilic stabilization of the
oxyanion by rhomboid, and the network of interactions that
secure substrate for catalysis.
Electrophilic Catalysis and Oxyanion Stabilization
The decisive step in serine protease catalysis is formation of the
high-energy transition state tetrahedral intermediate. In fact,
although most attention is usually paid to the nucleophilic attack
by the serine, progressing through this step absolutely relies on
enzymatic stabilization of the oxyanion. Our work here finally reveals the true nature of this electrophilic form of catalysis by
rhomboid proteases, because it visualizes a peptide with a true
tetrahedral carbon and oxyanion. Prior structures observed a
variety of oxyanion interactions and distances, which have
been confounding to extrapolate to natural proteolysis because
the inhibitors that were studied distort the catalytic center by
covalently attaching to both the catalytic serine and histidine,
and/or were not peptidic, and/or did not produce a carbon-oxyanion (Vinothkumar et al., 2010, 2013; Vosyka et al., 2013; Xue
et al., 2012; Xue and Ha, 2012; Zoll et al., 2014).
Interestingly, the mechanism of oxyanion stabilization in rhomboid also proved to be unexpected, because it is tripartite, which
has not, to our knowledge, been observed previously for any protease. This multifaceted oxyanion stabilizing network might also
explain the long-standing puzzle of why mutating the asparagine
residue compromises protease activity to different extents in
different rhomboid enzymes (Urban et al., 2001); local geometric
Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc. 337
Figure 7. Kinetic Scheme of Rhomboid Intramembrane Proteolysis and Its Inhibition by Substrate-Mimicking Peptide Aldehydes
A rhomboid protease is drawn in blue, with the substrate transmembrane helix in green, and the aldehyde moiety of the substrate-mimicking peptide aldehyde
inhibitor depicted as a red star. The white S (scission site) in rhomboid marks the inner site of catalysis, while the I (interrogation site) indicates the peripheral gateopen site to which transmembrane segments first dock. We previously determined experimentally that KM is equal to Kd for rhomboid proteolysis (Dickey
et al., 2013).
differences in some rhomboid enzymes might allow them to
compensate more easily than others for the absence of asparagine during oxyanion stabilization.
Distal Rhomboid:Substrate Interactions
Beyond the site of catalysis, we also visualized the network of
distal rhomboid:substrate interactions. Rhomboid proteases
generally display broad sequence selectivity, and indeed the
predominant substrate:enzyme interactions were backbonebackbone interactions with the extended substrate chain sandwiched between the L3 and L5 loops. Nevertheless various
substrate mutagenesis studies have converged on the importance of small P1 residues for substrate cleavage (Akiyama
and Maegawa, 2007; Moin and Urban, 2012; Strisovsky et al.,
2009). Contrary to expectation, the P1 methyl side chain points
into a surprisingly large cavity that could potentially accommodate valine, yet valine in this position blocks cleavage completely
(Dickey et al., 2013; Strisovsky et al., 2009). This large cavity contains three water molecules that we previously identified and
postulated form a water-retention site to aid proteolysis (Zhou
et al., 2012). The stark requirement for small residues at P1
may thus not be steric but rather may result from a disruptive
effect larger side chains may have on the water-retention site.
It must be emphasized that further analysis is required to test
this hypothesis.
Some rhomboid enzymes (Strisovsky et al., 2009), but not
others (Moin and Urban, 2012; Parussini et al., 2012), also
display a preference for hydrophobic P4 residues for efficient
substrate proteolysis. With GlpG, the P4 side chain forms a
shallow but specific interaction with the top of the L1 loop. However, because GlpG has only a mild preference for large residues
at this position, visualizing a P4-dependent rhomboid like AarA
would provide more information. More distant residues appear
to have limited effect on proteolysis; both P5 and P6 residues
were disordered in our structure, arguing that GlpG does not
contain specific substrate-binding subsites beyond P4. The P1
and P4 interactions we observed are consistent with a beautiful
study that used peptide chloromethylketones modeled on the
TatA substrate (Zoll et al., 2014); despite major differences in
warhead chemistry and peptide sequence, the overall distal contacts are similar, providing confidence that these approaches
are revealing true characteristics of extended substrate:rhomboid interactions.
Unexpected Substrate Interactions Drive Rhomboid
Inhibition
More surprising and potentially useful are the unexpected P2
and P3 interactions we discovered, because these positions
were thought to be unrestricted in rhomboid substrates, and
the side chains capable of making these interactions were not
studied in other structures (Strisovsky et al., 2009; Zoll et al.,
2014). Indeed, although the side chains face up and out of the
active site, they make fortuitous yet highly stabilizing interactions. The series of ‘‘extra’’ P3 arginine contacts with the L5
loop alone increase peptide aldehyde potency by 10-fold, yield
inhibitors that are effective without requiring any pre-incubation
with rhomboid, and are able to block proteolysis inside the membrane (rather than in detergent, which renders the active site
more accessible). In fact, these peptide aldehydes were effective
at inhibiting endogenous rhomboid activity to completion in living
E. coli cells.
An even more surprising feature that proved to be critical for
effective inhibition in vivo was the reversible aldehyde warhead
itself. It is widely (and reasonably) assumed that irreversible
inhibitors are more potent, because they trap proteases covalently in dead-end complexes. This, however, turned out to be
338 Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc.
incorrect with rhomboid, because identical peptides harboring
the well-studied irreversible chloromethylketone warhead in
place of the aldehyde were 10-fold less effective at inhibiting
rhomboid in living cells. Whether this results from the unusual
catalytic properties of rhomboid proteases (Dickey et al., 2013)
and/or reflects chemical differences imparted by the membrane
environment (Moin and Urban, 2012) remains to be determined.
Nevertheless, these observations should motivate a re-evaluation of current strategies for rhomboid inhibition, which are
centered entirely on irreversible warheads (Nguyen et al.,
2015). In simple terms, starting with reversible warheads such
as aldehydes and systematically cultivating interactions at neglected ‘‘nonessential’’ positions in substrates may prove to be
an untapped but valuable strategy for the rational design of selective rhomboid inhibitors.
Finally, although this marks a promising start both to understanding catalysis in the membrane and discovering opportunities for its selective inhibition, additional high-resolution
structures are essential to achieving a full understanding of
rhomboid proteolysis. We now have structures of the catalytic
scission complex but an obvious lingering gap is the Michaelis
complex, which remains a mystery. Nevertheless, our current efforts establish a platform for leveraging the unusual features of
rhomboid proteases to achieving these goals for this fascinating
but incompletely understood class of membrane-immersed
enzymes.
EXPERIMENTAL PROCEDURES
Protein Purification, Crystallization, Data Collection, and Refinement
E. coli GlpG (DN-GlpG: residues 87–276) was expressed and purified as
described (Dickey et al., 2013). Thermostability analysis was performed using
a StarGazer-384 differential static light-scattering system (Harbinger Biotech)
as described in detail previously (Baker and Urban, 2012). For structural analysis, all rhomboid enzymes were expressed and purified under identical conditions, indicating that differences in conformation result mainly from their final
environment (detergent versus bicelle) during crystallization. The active
DN-GlpG Y205F variant was crystallized from a bicelle composed of DMPC/
CHAPSO (2.8:1) that was prepared using a previously developed method as
described (Faham and Bowie, 2002). Reservoir buffer contained 0.1 M NaOAc
(pH 5.5), 3M NaCl, and 5% ethylene glycol. Crystals of DN-GlpG in the gateclosed form were obtained in hanging drops at room temperature over a reservoir containing 0.1 M Tris (pH 8.5), 3 M NaNO3, and 15% glycerol. Peptide
aldehydes were custom synthesized commercially using solid-phase chemistry, purified to >90% purity by reverse-phase high-performance liquid chromatography, and verified by mass spectrometry. Crystals were incubated with
2.5 mM peptide aldehydes in reservoir buffer for 7–24 hr at room temperature
to generate the substrate complex, and flash-frozen in a nitrogen stream. X-ray
diffraction data were collected at the Cornell High Energy Synchrotron Source,
processed with iMosflm 1.0.7, and scaled with Aimless in the CCP4 program
suite. Structures were solved by molecular replacement using Molrep. Electron density maps calculated from the solution of molecular replacement
clearly revealed density directly above of connected with catalytic S201, and
was modeled as the substrate aldehyde. Structures were further refined by
refmac5 and PHENIX with iterative manual model building using COOT.
In Vitro Rhomboid Proteolysis Assay
Real-time kinetic assays were performed as described (Dickey et al., 2013):
2–80 pmoles of rhomboid were mixed with 30 mg of liposomes formed from
an E. coli polar lipid extract (Avanti Polar Lipids) and 25–1,600 pmoles of fluorescein isothiocyanate (FITC)-TatA substrate in 50 mM Na-acetate (pH 4.0)
and 150 mM NaCl. Proteoliposomes were formed by rapid dilution and
collected by ultracentrifugation at 600,000 3 g for 30 min, and proteolysis
was initiated by neutralizing proteoliposomes with 50 mM Tris (pH 7.4) and
150 mM NaCl (and simultaneously adding peptides without pre-incubation
for inhibition experiments). Because membranes quench FITC-TatA, rhomboid-mediated proteolytic release was monitored at 37 C in real time by quantifying FITC fluorescence in a Synergy H4 Hybrid plate reader (BioTek). For
inhibition experiments, initial rates were extracted from progress curves monitored in real time, and reduction in rate was plotted as a function of inhibitor
concentration and modeled using the R environment with the equation
v0;I
Ki
;
=
v0 Ki + ½I
in which v0,I and v0 are initial rates in the presence and absence of inhibitor,
respectively.
Analysis of Rhomboid Proteolysis and Inhibition in E. coli Cells
The GlpG+ E. coli K12 strain NR698 (kind gift from Tom Silhavy, Princeton University) that allows more accurate IC50 determination by rendering the outer
membrane more permeable to small molecules (Ruiz et al., 2005) was used
to generate a DGlpG strain in a two-step process. First, we introduced
GlpG::Kan through P1-mediated transduction and then excised the Kan
gene using FRT-mediated recombination. The wild-type and DGlpG strains
were transformed with pBAD-PsTatA-Flag (Dickey et al., 2013) and grown in
LB + ampicillin to an optical density at 600 nm of 0.4 by shaking cultures
at 250 rpm at 37 C. Expression of the PsTatA-Flag substrate was then induced
by adding arabinose to 25 mM or 1 mM, the indicated peptides were added to
the induced cultures, and the cultures were grown for 2 hr, shaking at 250 rpm
at 37 C. Whole-cell lysates were prepared by pelleting 0.8 ml of each culture
and resuspending the cell pellets in reducing Tricine-SDS sample buffer (Life
Technologies) and lysed by sonication. Proteins were resolved on 16% Tricine-SDS-PAGE gels (Life Technologies), transferred to nitrocellulose using
the Trans-Blot Turbo system (Bio-Rad), and probed with the indicated antibodies. Immune complexes were visualized and quantified by infrared laser
scanning on an Odyssey imager (Li-Cor Biosciences).
ACCESSION NUMBERS
Structure coordinates have been deposited into the PDB (http://www.rcsb.
org/pdb/home/home.do) under the accession codes PDB: 5F5B (GlpG+
VRMA), PDB: 5F5D (GlpG-Y205F_bicelle), PDB: 5F5G (GlpG-Y205F_bicelle+
RMA), PDB: 5F5J (GlpG-Y205F_bicelle+VRMA), and PDB: 5F5K (GlpGY205F_bicelle+RKVRMA).
SUPPLEMENTAL INFORMATION
Supplemental Information includes one figure and can be found with this
article online at http://dx.doi.org/10.1016/j.molcel.2015.12.022.
AUTHOR CONTRIBUTIONS
S.U. designed the study and oversaw its execution. S.C. solved the structures
of GlpG with substrate peptides. S.U. and S.W.D. generated DNA constructs,
S.W.D. performed enzyme activity and inhibition analyses, and S.U. conducted the in vivo inhibition experiments. S.U. wrote the paper, and all authors
approved the final manuscript.
ACKNOWLEDGMENTS
We are grateful to members of the Urban lab for helpful discussions, particularly to Rosanna Baker, who purified the mutants analyzed in Figures 5C and
5D. This work was supported by National Institutes of Health (NIH) grants
2R01AI066025 and R01AI110925, the Howard Hughes Medical Institute, and
the David and Lucile Packard Foundation. X-ray diffraction data were collected
using instruments at the Cornell High Energy Synchrotron Source (beamline
supported by National Science Foundation grant DMR-0936384 and NIH grant
GM-103485).
Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc. 339
Received: July 16, 2015
Revised: November 20, 2015
Accepted: December 15, 2015
Published: January 21, 2016
Nguyen, M.T., Kersavond, T.V., and Verhelst, S.H. (2015). Chemical Tools for
the Study of Intramembrane Proteases. ACS Chem. Biol. 10, 2423–2434.
REFERENCES
Parussini, F., Tang, Q., Moin, S.M., Mital, J., Urban, S., and Ward, G.E. (2012).
Intramembrane proteolysis of Toxoplasma apical membrane antigen 1 facilitates host-cell invasion but is dispensable for replication. Proc. Natl. Acad.
Sci. U S A 109, 7463–7468.
Akiyama, Y., and Maegawa, S. (2007). Sequence features of substrates
required for cleavage by GlpG, an Escherichia coli rhomboid protease. Mol.
Microbiol. 64, 1028–1037.
Pierrat, O.A., Strisovsky, K., Christova, Y., Large, J., Ansell, K., Bouloc, N.,
Smiljanic, E., and Freeman, M. (2011). Monocyclic b-lactams are selective,
mechanism-based inhibitors of rhomboid intramembrane proteases. ACS
Chem. Biol. 6, 325–335.
Baker, R.P., and Urban, S. (2012). Architectural and thermodynamic principles
underlying intramembrane protease function. Nat. Chem. Biol. 8, 759–768.
Bondar, A.N., del Val, C., and White, S.H. (2009). Rhomboid protease dynamics and lipid interactions. Structure 17, 395–405.
Brooks, C.L., and Lemieux, M.J. (2013). Untangling structure-function relationships in the rhomboid family of intramembrane proteases. Biochim. Biophys.
Acta 1828, 2862–2872.
Brown, M.S., Ye, J., Rawson, R.B., and Goldstein, J.L. (2000). Regulated intramembrane proteolysis: a control mechanism conserved from bacteria to humans. Cell 100, 391–398.
Powers, J.C., Asgian, J.L., Ekici, O.D., and James, K.E. (2002). Irreversible inhibitors of serine, cysteine, and threonine proteases. Chem. Rev. 102, 4639–
4750.
Ruiz, N., Falcone, B., Kahne, D., and Silhavy, T.J. (2005). Chemical conditionality: a genetic strategy to probe organelle assembly. Cell 121, 307–317.
Strisovsky, K., Sharpe, H.J., and Freeman, M. (2009). Sequence-specific intramembrane proteolysis: identification of a recognition motif in rhomboid substrates. Mol. Cell 36, 1048–1059.
Urban, S. (2009). Making the cut: central roles of intramembrane proteolysis in
pathogenic microorganisms. Nat. Rev. Microbiol. 7, 411–423.
Chan, E.Y., and McQuibban, G.A. (2013). The mitochondrial rhomboid protease: its rise from obscurity to the pinnacle of disease-relevant genes.
Biochim. Biophys. Acta 1828, 2916–2925.
Urban, S., and Moin, S.M. (2014). A subset of membrane-altering agents and
g-secretase modulators provoke nonsubstrate cleavage by rhomboid proteases. Cell Rep. 8, 1241–1247.
De Strooper, B., and Annaert, W. (2010). Novel research horizons for presenilins and g-secretases in cell biology and disease. Annu. Rev. Cell Dev. Biol. 26,
235–260.
Urban, S., Lee, J.R., and Freeman, M. (2001). Drosophila rhomboid-1 defines a
family of putative intramembrane serine proteases. Cell 107, 173–182.
Dickey, S.W., Baker, R.P., Cho, S., and Urban, S. (2013). Proteolysis inside the
membrane is a rate-governed reaction not driven by substrate affinity. Cell
155, 1270–1281.
Drag, M., and Salvesen, G.S. (2010). Emerging principles in protease-based
drug discovery. Nat. Rev. 9, 690–701.
Faham, S., and Bowie, J.U. (2002). Bicelle crystallization: a new method for
crystallizing membrane proteins yields a monomeric bacteriorhodopsin structure. J. Mol. Biol. 316, 1–6.
Friedman, J.I., and Stivers, J.T. (2010). Detection of damaged DNA bases by
DNA glycosylase enzymes. Biochemistry 49, 4957–4967.
Golde, T.E., Koo, E.H., Felsenstein, K.M., Osborne, B.A., and Miele, L. (2013).
g-Secretase inhibitors and modulators. Biochim. Biophys. Acta 1828, 2898–
2907.
Hedstrom, L. (2002). Serine protease mechanism and specificity. Chem. Rev.
102, 4501–4524.
Urban, S., Schlieper, D., and Freeman, M. (2002). Conservation of intramembrane proteolytic activity and substrate specificity in prokaryotic and eukaryotic rhomboids. Curr. Biol. 12, 1507–1512.
Vinothkumar, K.R. (2011). Structure of rhomboid protease in a lipid environment. J. Mol. Biol. 407, 232–247.
Vinothkumar, K.R., Strisovsky, K., Andreeva, A., Christova, Y., Verhelst, S.,
and Freeman, M. (2010). The structural basis for catalysis and substrate specificity of a rhomboid protease. EMBO J. 29, 3797–3809.
Vinothkumar, K.R., Pierrat, O.A., Large, J.M., and Freeman, M. (2013).
Structure of rhomboid protease in complex with b-lactam inhibitors defines
the S20 cavity. Structure 21, 1051–1058.
Vosyka, O., Vinothkumar, K.R., Wolf, E.V., Brouwer, A.J., Liskamp, R.M., and
Verhelst, S.H. (2013). Activity-based probes for rhomboid proteases discovered in a mass spectrometry-based assay. Proc. Natl. Acad. Sci. U S A 110,
2472–2477.
Wolfe, M.S. (2009). Intramembrane proteolysis. Chem. Rev. 109, 1599–1612.
Henderson, R. (1970). Structure of crystalline alpha-chymotrypsin. IV. The
structure of indoleacryloyl-alpha-chyotrypsin and its relevance to the hydrolytic mechanism of the enzyme. J. Mol. Biol. 54, 341–354.
Xue, Y., and Ha, Y. (2012). Catalytic mechanism of rhomboid protease GlpG
probed by 3,4-dichloroisocoumarin and diisopropyl fluorophosphonate.
J. Biol. Chem. 287, 3099–3107.
Kamp, F., Winkler, E., Trambauer, J., Ebke, A., Fluhrer, R., and Steiner, H.
(2015). Intramembrane proteolysis of b-amyloid precursor protein by g-secretase is an unusually slow process. Biophys. J. 108, 1229–1237.
Xue, Y., Chowdhury, S., Liu, X., Akiyama, Y., Ellman, J., and Ha, Y. (2012).
Conformational change in rhomboid protease GlpG induced by inhibitor binding to its S0 subsites. Biochemistry 51, 3723–3731.
Manolaridis, I., Kulkarni, K., Dodd, R.B., Ogasawara, S., Zhang, Z., Bineva, G.,
O’Reilly, N., Hanrahan, S.J., Thompson, A.J., Cronin, N., et al. (2013).
Mechanism of farnesylated CAAX protein processing by the intramembrane
protease Rce1. Nature 504, 301–305.
Zhou, Y., Moin, S.M., Urban, S., and Zhang, Y. (2012). An internal water-retention site in the rhomboid intramembrane protease GlpG ensures catalytic efficiency. Structure 20, 1255–1263.
Moin, S.M., and Urban, S. (2012). Membrane immersion allows rhomboid proteases to achieve specificity by reading transmembrane segment dynamics.
eLife 1, e00173.
Zoll, S., Stanchev, S., Began, J., Skerle, J., Lepsı́k, M., Peclinovská, L., Majer,
P., and Strisovsky, K. (2014). Substrate binding and specificity of rhomboid intramembrane protease revealed by substrate-peptide complex structures.
EMBO J. 33, 2408–2421.
340 Molecular Cell 61, 329–340, February 4, 2016 ª2016 Elsevier Inc.