Coupling protein engineering with probe design to inhibit and image matrix metalloproteinases with controlled specificity

Article abstract of DOI:10.1021/ja403523p

Matrix metalloproteinases (MMPs) are zinc endopeptidases that play roles in numerous pathophysiological processes and therefore are promising drug targets. However, the large size of this family and a lack of highly selective compounds that can be used for imaging or inhibition of specific MMPs members has limited efforts to better define their biological function. Here we describe a protein engineering strategy coupled with small-molecule probe design to selectively target individual members of the MMP family. Specifically, we introduce a cysteine residue near the active-site of a selected protease that does not alter its overall activity or function but allows direct covalent modification by a small-molecule probe containing a reactive electrophile. This specific engineered interaction between the probe and the target protease provides a means to both image and inhibit the modified protease with absolute specificity. Here we demonstrate the feasibility of the approach for two distinct MMP proteases, MMP-12 and MT1-MMP (or MMP-14).

Full text of DOI:10.1021/ja403523p

Article
pubs.acs.org/JACS
Coupling Protein Engineering with Probe Design To Inhibit and
Image Matrix Metalloproteinases with Controlled Specificity
Montse Morell,† Thinh Nguyen Duc,† Amanda L. Willis,‡ Salahuddin Syed,† Jiyoun Lee,†,⊥ Edgar Deu,†,#
Yang Deng,§ Junpeng Xiao,† Benjamin E. Turk,§ Jason R. Jessen,∥ Stephen J. Weiss,‡
and Matthew Bogyo*,†
†Department of Pathology, Stanford University School of Medicine, Stanford, California 94305, United States
‡Division of Molecular Medicine and Genetics, Department of Internal Medicine, Life Sciences Institute, University of Michigan, Ann
Arbor, Michigan 48109, United States
§Department of Pharmacology, Yale University School of Medicine, New Haven, Connecticut 06520, United States
∥Department of Medicine/Division of Genetic Medicine, Vanderbilt University Medical Center, Nashville, Tennessee 37232, United
States
S
* Supporting Information
ABSTRACT: Matrix metalloproteinases (MMPs) are zinc
endopeptidases that play roles in numerous pathophysiological
processes and therefore are promising drug targets. However,
the large size of this family and a lack of highly selective
compounds that can be used for imaging or inhibition of
specific MMPs members has limited efforts to better define
their biological function. Here we describe a protein
engineering strategy coupled with small-molecule probe design
to selectively target individual members of the MMP family.
Specifically, we introduce a cysteine residue near the active-site
of a selected protease that does not alter its overall activity or function but allows direct covalent modification by a small-
molecule probe containing a reactive electrophile. This specific engineered interaction between the probe and the target protease
provides a means to both image and inhibit the modified protease with absolute specificity. Here we demonstrate the feasibility of
the approach for two distinct MMP proteases, MMP-12 and MT1-MMP (or MMP-14).
probes (ABPs).8 These reagents are based on covalent
irreversible inhibitors that can be appropriately tagged and
designed to target specific proteases. However, the develop-
ment of suitable probes for the MMPs has proven difficult
INTRODUCTION
■
The matrix metalloproteinases (MMPs) are a large family of
enzymes that have been found to regulate many aspects of both
normal cellular biology and pathogenesis of human diseases,
because these proteases use a water nucleophile for initial attack
most notably cancer.1,2 However, attempts to develop novel
of the substrate,9 preventing the design of inhibitors that
anti-cancer agents that target MMPs have proven to be largely
directly modify a primary catalytic residue. To overcome this
unsuccessful. These failures have been attributed, in part, to a
limitation, ABPs for metalloproteases have been designed based
generally poor understanding of the physiological functions of
individual members of the family.3 MMPs, like most proteases,
on a zinc-chelating hydroxamate and a benzophenone photo-
cross-linking group. These probes can be used to covalently
are regulated by multiple post-translational mechanisms,
label the active-sites of many MMP proteases.10−13 However,
including synthesis as inactive zymogens and inhibition by
while useful for proteomic, biochemical, or cell biological
endogenously expressed protein inhibitors such as the tissue
inhibitors of metalloproteinases (TIMPs).4 These post-transla-
studies of multiple MMP proteases, the overall lack of
selectivity of the probes as well as the need for photo-cross-
tional events hinder their functional analysis by conventional
linking to covalently link probes into the active-site limits their
genomic and proteomic methods. In addition, the large size of
the family coupled with the relatively well-conserved active-site
utility for studies of specific MMPs in complex biological
region has prevented the design of highly selective inhibitors.5,6
systems.
Here we present a strategy for selective targeting of a single
MMP protease using a combination of protein engineering
methods and small-molecule probe design strategies. This
Thus, new tools and reagents that enable direct monitoring of
specific MMP activity as well as specific inhibition of these
proteases would enable studies of the specific roles of particular
MMPs in disease as well as in basic biological processes.7
One method for functional studies of proteases that has
Received: April 9, 2013
proven valuable is the use of small-molecule activity-based
© XXXX American Chemical Society
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method is based on the design of a mutant protease that
contains an engineered cysteine residue that does not alter
expression, folding, or substrate recognition but that is capable
of acting as a latent nucleophile that reacts covalently with a
suitably designed probe containing a reactive electrophile
(Figure 1a). We chose this general strategy because this type of
structures and substrate cleavage profiles. Using two matrix
metalloproteinases, MMP12 and MT1-MMP (MMP14), we
demonstrate that it is possible to find locations near the active-
site that can be mutated to cysteine without altering functional
properties. We also demonstrate targeting of the engineered
cysteine with a chemical probe that contains a hydroxamate
group for binding to the active-site zinc coupled with an
appropriately placed electrophile. This modification results in
permanent covalent labeling and inhibition, thus enabling
specific inhibition and imaging studies of a single defined
protease target.
MATERIALS AND METHODS
■
Synthesis. For full details on the synthesis of TND124 and
TND126 see Supporting Information.
Cell Culture and Transfection. MCF7 and HEK293T (both
from American Type Culture Collection) were maintained in DMEM
supplemented with 10% fetal bovine serum, 2 mM L-glutamine, and
antibiotics. All cultures were maintained at 37 °C in 5% CO2/95% air.
Cells were transfected with purified plasmid DNA by Lipofectamine
treatment (LifeTechnologies) as recommended by the manufacturer.
Protein Alignment. Protein alignment of the members of the
human MMP family was performed using CLUSTALW2 (http://www.
clustal.org).
Construction of Expression Plasmids. The catalytic domains of
wild-type (WT) or engineered MMPs were recombinantly expressed
in Escherichia coli as fusions of the Vibrio cholerae MARTX toxin
cysteine protease domain (CPD). The pET22b-mMMP12-CPD
construct was generated by PCR amplification of the sequence
encoding the catalytic domain of mouse MMP12 (amino acids 29−
267) using pCMV-SPORT6-mMMP12 (Open Biosystems) as a
template and primers MMP12-5′ and MMP12-3′ (Table S1,
Supporting Information). All the protein mutants were created using
PCR fusion methods (primers MMP12-T184C-5′, MMP12-T184C-3′,
MMP12-T243C-5′, and MMP12-T243C-3′; Table S1, Supporting
Information), and the resulting PCR products were cloned into the
NdeI and SalI sites of pET22b-CPDSalI. The pET22b-mMT1-MMP-
CPD construct was generated by PCR amplification of the sequence
encoding the catalytic domain of mouse MT1-MMP (amino acids 28−
318) using pCDNA3-mMT1-MMP (a kindly gift from Dr. Rozanov)
as a template and primers MT1-MMPcat-5′ and MT1-MMPcat-3′
(Table S1, Supporting Information). All the protein mutants were
created using PCR fusion methods (primers MT1-MMPcat-F198C5′,
MT1-MMPcat-F198C3′, MT1-MMPcat-F260C5′, MT1-MMPcat-
F260C3′, MT1-MMPcat-Q262C5′, and MT1-MMPcat-Q262C3′;
Table S1, Supporting Information), and the resulting PCR products
were cloned into the NdeI and SacI sites of pET22b-CPDSacI. Full-
length HA-tagged mMT1-MMP was created by inserting an HA tag
(YPYDVPDYA) between D515 and E516 as described previously.18
The final insert was inserted into the pCDNA3.1 plasmid between the
HindIII and XbaI cut sites.
Protein Expression and Purification. For purification of His6-
tagged CPD fusion proteins, overnight cultures of the appropriate
strain were diluted 1:500 into 1 L of 2YT media and grown with
shaking at 37 °C. When an OD600 of 0.6 was reached, IPTG was added
to 250 μM, and cultures were grown for 8 h at 16 °C. Cultures were
pelleted, resuspended in 25 mL lysis buffer [500 mM NaCl, 50 mM
Tris-HCl, 1 mM tris(2-carboxyethyl)phosphine (TCEP), pH 7.5, 15
mM imidazole, 10% glycerol] and flash frozen in liquid nitrogen.
Lysates were thawed, then lysed by sonication and cleared by
centrifugation at 15000g for 30 min. His6-tagged CPD fusion proteins
were affinity purified by incubating the lysates in batch with 0.5−1.0
mL Ni-NTA agarose beads (Qiagen) with shaking for 2−4 h at 4 °C.
The binding reaction was pelleted at 1500g, the supernatant was set
aside, and the pelleted Ni2+-NTA agarose beads were washed three
times with lysis buffer.
Figure 1. Protease engineering and probe design. (a) The target
protease is engineered with a non-catalytic cysteine near the substrate
binding pocket. The designed probe contains a reversibly binding
reactive group (hydroxamate), an electrophile (α-chloroacetamide), a
peptide backbone that drives specificity toward MMPs (in orange),
and a fluorescent tag (green). When the hydroxamate binds to the
active-site of engineered protease, the electrophile irreversibly labels
the protein through the engineered cysteine, resulting in a
permanently labeled and inhibited protease. When the probe binds
to WT MMPs the binding is reversible. (b) Sequence alignment of
mouse MMPs. The two non-conserved residues (184, site 1, and 243,
site 2 in MMP12) that were chosen for mutation are indicated by
arrows. (c) Superposition of the crystal structures of MMP12 (1JIZ)
and MT1-MMP (3MA2). The active-site Zn2+ is shown as a red ball.
The candidate residues are drawn using stick representation (left). The
surface representation of the structure of the inhibitor GM6001 bound
to the catalytic domain of MMP12 is also shown (right). The GM6001
is shown as orange sticks. The Zn2+ is shown as a red ball. The two
mutation sites are highlighted in yellow.
proximity-induced labeling of an engineered cysteine has
already been validated for the identification of small-molecule
ligands14 as well as for covalent labeling of a engineered protein
target that has been fused to a small-molecule binding
domain.15 It has also been demonstrated as a way to drive
selectivity of a small-molecule inhibitor toward a specific kinase
target16 and to target a protease with a native cysteine residue
near the active-site.17 Thus, we believed that this approach
could be generally applied as a way to target specific proteases
within large families that have highly similar active-site
To liberate untagged target proteins into the supernatant fractions,
300−500 μL of lysis buffer was added to the Ni2+-NTA beads
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containing CPD-His6 fusion proteins, and the indicated amount of
inositol hexakisphosphate (InsP6, Calbiochem) was added. In general,
on-bead cleavage was performed by incubating the beads in the
presence of 50−100 μM InsP6 for 1−2 h at either room temperature
or 4 °C. The beads were pelleted at 1500g, and the supernatant
fraction was removed. The beads were then washed 3−4 times with
300−500 μL of lysis buffer, and supernatant fractions were retained.
Enzymatic Activity. The recombinant enzyme was diluted to a
final concentration of 100 nM in the reaction buffer (50 mM Tris-HCl,
pH 7.5, 150 mM NaCl, 10 mM CaCl2, 0.02% NaN3), and 10 μL of 100
μM of the substrate (Mca-Pro-Leu-Gly-Leu-Dpa-Ala-Arg-NH2,
Anaspec) was added. The substrate turnover was immediately
measured by fluorescent emission over a 1 h period at 37 °C using
a Spectramax M5 plate reader (Molecular Devices).
The hydrolysis of the fluorogenic substrate (0−10 μM) by WT or
F260C MT1-MMP was measured at 37 °C in a continuous plate
reader-based protocol. Fluorescence associated with the turnover of
the substrate was measured every 30 s for 1 h (λex 325 nm, λem 393
nm) in a Spectramax M5 plate reader (Molecular Devices). The rate of
product formation was determined using linear regression of the
fluorescence−time data with the plate reader software (SOFTmax Pro,
v 4.8; Molecular Devices Inc., Sunnyvale, CA). Linear reaction rates
(R2 > 0.98 for all data) were fitted to the Michaelis−Menten equation
using the nonlinear curve-fitting facility of Kaleida Graph in order to
obtain Vmax and Km values.
Peptide Substrate Profiling. The peptide substrate profiling was
performed using a mixture-based oriented peptide library.19 Degen-
erate positions X were prepared using isokinetic mixtures of the 19
naturally occurring L-amino acids excluding cysteine. The exact
proportion of each amino acid at each degenerate position was
determined by Edman sequencing of an unacetylated portion of the
library. Individual peptides were purified by high-performance liquid
chromatography and characterized by mass spectrometry. To
determine the primed-side cleavage specificity, a 1 mM solution of
Ac-X-X-X-X-X-X-X-X-X-X-[D-Lys]-[D-Lys] in MMP reaction buffer
(50 mM HEPES, pH 7.4, 200 mM NaCl, 10 mM CaCl2) was digested
at 37 °C to 5−10% cleavage and heated to 100 °C for 2 min. A 10 μL
aliquot was subjected to Edman sequencing on an Applied Biosystems
(Foster City, CA) Procise 494 automated protein sequencer at the
Tufts University Core Facility (Boston, MA). Amino acid preference
in a given cycle was calculated by dividing the amount of a particular
residue by the average amount per amino acid residue in that cycle.
The data were then corrected for bias present in the library by dividing
each value by the relative amount of that particular amino acid in the
starting mixture.
In Silico Modeling. All modeling was performed using the
Molecular Operating Environment (MOE) software with the default
energy minimization settings. The N238Cys and I179Cys mutations
were modeled into the crystal structure of human MMP8 bound to the
peptidic hydroxamate inhibitor HONH-iBM-Asn-NHBn(m-NH2)
(PDB code: 1A85). For each model, mutated residues and all side
chains within 4.5 Å were energy minimized. ABPs were docked into
the models by sequentially modifying the side chains of the HONH-
iBM-Asn-NHBn(m-NH2) inhibitor into those of potential ABPs
starting from the P1′ position and ending with the addition of the
fluorophore. Energy minimization was performed as follows: each side
chain in the ABP was built manually followed by three energy
minimizations: (i) the modified side chain, (ii) the modified side chain
plus all MMP8 residues within 4.5 Å, and finally (iii) the whole ligand
plus all MMP8 residues within 4.5 Å. This operation was repeated for
each side chain until we obtained models of ABPs bound to MMP8
mutants before covalent modification of the engineered Cys by the α-
chloroacetamide electrophile. Multiple residues at the P2′ position
were tested to identify the optimal side-chain length to achieve close
contacts between the α-chloroacetamide group and the engineered
Cys. The P2′ side chain was build to point toward either Cys179 or
Cys238. After obtaining a model of the ABP non-covalently bound to
MMP8, we modeled the end product of the reaction between the
engineered Cys and the α-chloroacetamide group, followed by energy
minimization of the ABPs and all residues within 4.5 Å. For TND124,
we found that the hydroxamate moiety remained within binding
distance of the active-site Zn2+ (Figure S2, Supporting Information).
Gel Labeling. Recombinant enzyme (100 ng) was spiked in rat
liver lysates (50 μg) and incubated with different concentrations of the
probe for 1 h at 37 °C. The probes were diluted to the desired
concentration from a 1000× stock solution in DMSO. Pre-incubation
with NEM (1 mM), EDTA (10 mM), GM6001 (10 μM), and
TND126 (without Cy5,10 μM) was performed for 30 min at 37 °C
before addition of TND124 (1 μM). These compounds were also
diluted to the final concentration from a 1000× stock solution. After
the incubation time, the samples were resolved by 15% SDS-PAGE gel
and visualized on a Typhoon flatbed fluorescent laser scanner (GE
Healthcare). In the case of the labeling of HEK293T cell lysates
transiently expressing MT1-MMP WT or mutant, cells were lysed
using hypotonic buffer (10 mM PIPES, 10 mM KCl, 5 mM MgCl2,
and 4 mM DTT). The lysates were then incubated with 0.5 μM
TND124 for 1 h at 37 °C. After the incubation time, the samples were
then resolved by 15% SDS-PAGE gel and visualized on a Typhoon
flatbed fluorescent laser scanner (GE Healthcare).
Probe Washout Studies. Recombinant enzyme (1000 ng, WT or
T184C MMP12) was incubated with 3 μM TND124 for 1 h at 37 °C.
Afterward, samples were dialyzed using Slide-A-Lyzer MINI Dialysis
Devices, 10K MWCO (Thermo), for 24 and 48 h at 4 °C. For each
time point (before addition of TND124 and 24 and 48 h after
dialysis), 50 μL of each sample was used to detect the activity of the
enzyme using a fluorogenic substrate following the protocol
mentioned above.
In Vitro TND124 Inhibition Kinetics. Recombinant MMP12 WT
and T184C (10 nM) were incubated with increasing concentrations of
TND124 in activity buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl,
10 mM CaCl2, 0.02% NaN3) at 37 °C in a 96-well plate. After 1 h
incubation, 10 μM of the fluorogenic substrate was added, and the
initial rates (V0) of substrate hydrolysis were determined by
fluorescent reading at 37 °C for 30 min. V0 values were converted
to percentages of residual activity relative to untreated controls. The
IC50 values were determined by KaleidaGraph using sigmoidal fit. The
final presented IC50 values represent the average of three independent
assays.
Inhibition Studies of MMP12 Activity against Natural
Substrates. Gelatin FITC conjugated and collagen IV FAM
conjugated were purchased from Anaspec. Recombinant enzymes
(3000 ng, WT or T184C MMP12) were incubated with TND124 (3
μM) for 1 h at 37 °C. Afterward, the samples were diluted 1/10 with
the activity buffer containing the gelatin or collagen substrate (at a final
concentration of 0.1 or 0.05 mg/mL, respectively) and incubated for
48 h at 37 °C. The samples were centrifuged at 10 000 rpm for 10 min,
and the fluorescence of the supernatant was measured at 515 nm using
a Spectramax M5 plate reader (Molecular Devices).
Casein Zymography. Casein zymography was performed based
on a previously described technique with slight modifications.20
Recombinant protein (500 ng, WT or T184C MMP12) was incubated
with 3 μM TND124 for 1 h at 37 °C. The samples were diluted 1/10
with the activity buffer and analyzed by electrophoresis. Electro-
phoresis was carried out on a sodium dodecyl sulfate (SDS)−10%
polyacrylamide gel containing α-casein (Sigma) at a concentration of 1
mg/mL. Gels were then incubated in 2.5% (v/v) Triton X-100 for 30
min and soaked for 16 h in a buffer containing 10 mM CaCl2 and 100
mM NaCl at 37 °C. The gels were then stained with Coomassie
brillant blue G250. Caseinolytic activity was detected as white lysis
zones against the blue background.
Intact Mass Spectrometry. Recombinant enzymes (50 μg, WT
or T184C MMP12) were incubated with TND124 and then analyzed
using a Thermo LTQ-FT mass spectrometer (Thermo Fisher
Scientific) using FTMS + p NSI full MS scanning mode, mass range
400.00−2000.00, FT resolution 100 000. The MW of eluted peptides
was determined by deconvolution using Isopro 3.0 (MS/MS
software).
Tryptic Digestion of Labeled Protein. Recombinant enzymes
(50 μg, WT or T184C MMP12) were incubated with TND124 and
digested with 1 μg of trypsin at room temperature for 18 h. The tryptic
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reactions were separated on a PLRPS 150 mm × 0.1 mm column
(Varian, 5 μM particle size, 300 Å pore size) at a flow rate of 700 nL/
min in 0.1% trifluoroacetic acid/water (A):0.1% trifluoroacetic acid/
acetonitrile (B). The column was run on a gradient of 10% B to 60% B
for 25 min, 60% B to 90% B for 2 min, held at 90% B for 2 min, rapidly
decreased to 10% B over 0.1 min, and then run for 11 min at 10% B.
Eluted samples were analyzed using a Thermo LTQ-FT mass
spectrometer (Thermo Fisher Scientific) using FTMS + p NSI full
MS scanning mode, mass range 400.00−2000.00, FT resolution 100
000. The MW of eluted peptides was determined by deconvolution
using Isopro 3.0 (MS/MS software). The measured masses were
compared to the predicted MW of possible peptide cleavage products
determined from the primary sequence of recombinant polypeptides
derived from MMP12 (modified or unmodified by TND124) using
PROTPARAM at http://ca.expasy.org/tools/protparam.html.
Collagen/Gelatin Degradation and Invasion Assays. Acid-
extracted rat tail type I collagen was prepared as previously described
and stored in acetic acid (0.035 M).21 Collagen solutions were
neutralized using equimolar concentrations of NaOH in a 1× MEM +
HEPES (25 mM) buffered solution and allowed to gel at 37 °C for 1 h
in a 95% air/5% CO2 atmosphere.21 For degradation assays, collagen
gels (2 mg/mL) were prepared as a thin coating in four-well
chambered coverglass slides. The film was then fluorescently labeled
using an AlexaFluor-(488 or 594) protein labeling kit (Invitrogen).
Cells were plated atop the collagen film at a density of 2 × 103 cells/
well in MEM supplemented with 10% FBS. In the experiments using
the inhibitor TND124, prior to plating atop the collagen film, the cells
were incubated in serum-free medium with 3 μM TND124 for 1.5 h at
37 °C. Afterward, cells were washed with media three times for 10 min
periods and plated atop the collagen film as described above, except
serum-free medium was used. Where indicated, gelatin degradation
was assessed using heat-denatured type I collagen (60 °C for 1 h) in
place of the native substrate.21 All assays were allowed to proceed for
72 h prior to imaging. For invasion assays, collagen gels (2 mg/mL)
were prepared at a thickness of 0.8−1.0 mm in six-well transwell
inserts (3 μm pore size, polystyrene, Corning). Cells were plated atop
the upper surface of the collagen gels at a density of 5 × 104 cells/well.
Medium containing 10% FBS was placed in the lower chamber, and
serum-free medium was added to the upper chamber. Invasion was
allowed to proceed for 5 days and quantified in randomly selected
fields at 20× on a phase-contrast microscope as described previously.21
Zymography for proMMP2 Activation. MT1-MMP activity was
confirmed by zymographic analysis of MMP-2 activation when the
transfected cells were incubated with human proMMP-2 expressed
from Timp2 deficient cells22 as has been described before.23
generated by PCR mutagenesis by overlap extension. Capped synthetic
mRNA was generated by Sp6 RNA polymerase in vitro transcription
(Ambion). For rescue experiments, one-cell-stage embryos were
injected with 8 ng of mmp14a MO, followed immediately by injection
with 200 pg of mRNA encoding either WT or mutant mt1-mmpa.
Embyros were incubated at 28.5 °C until 5 days post-fertilization and
fixed using 4% paraformaldehyde in PBS, and craniofacial cartilage
elements were visualized using Alcian blue as described previously.25
Imaging of Embryos. Using a pressure-based microinjection
apparatus (WPI), one-cell stage embryos were co-injected with 2 nL
of 50 μM TND124 and 300 pg of either WT or modified synthetic
mmp14a mRNA. At either the 8-somite or 18-somite stage, embryos
were fixed in 4% PFA/PBS overnight at 4 °C, followed by PBS
washing and manual dechorionation. Embryos were washed three
times for 5 min each in PBS/0.1% Triton X-100. DIC and Cy5 images
of the developing posterior trunk regions were taken at 200×
magnification using a Leica DMI6000 B inverted microscope equipped
with a Hamamatsu ORCA ER digital camera and Simple PCI software.
RESULTS
■
Engineering the Protease. We first set out to identify
residues that would be sufficiently proximal to an active-site-
bound probe to allow modification but that would not cause
changes in the overall substrate selectivity of the protease when
mutated to cysteine. Therefore, we looked for non-conserved
residues that are unlikely to contribute to folding or substrate
binding but that are near the substrate-binding pocket of the
enzyme. The MMPs have a high degree of structural
conservation in the active-site region, and high-resolution X-
ray structures of several members of this family are available. A
preliminary search of the primary sequences of the human
MMPs revealed two possible candidates (corresponding to
residues 198 and 260 in the protein alignment, Figure 1b).
These sites mapped to regions near the S2′ substrate binding
pocket and were in close proximity to a bound inhibitor (Figure
1c). We initially chose MMP12 as a target because this protease
has been expressed in recombinant form and very little is
known about its primary biological function other than it is
mainly secreted by macrophages26 and is involved in several
pathologies, such as emphysema.27
In order to assess the possible sites for introduction of the
cysteine residue, we first had to express the mutants in
recombinant form. We initially expressed the catalytic domains
of three mutant forms of MMP12 (T184C, T243C, and the
double mutant T184C/T243C) in E. coli using a protein
expression platform recently developed in our laboratory. This
method makes use of the cysteine protease domain (CPD) of
the bacterial MARTX toxin that autocatalytically processes itself
upon addition of inistol hexakis phosphate (IP6).28 One of the
main benefits of this approach is that it avoided refolding of the
protein from inclusion bodies as had been published for the
purification of active MMP12.29 We found that while all three
of the cysteine mutants were proteolytically active toward a
fluorogenic substrate, both T243C and T184C/T243C mutant
enzymes had a reduced activity (Figure 2a).
Therefore, we chose to focus our efforts on the T184C
variant that had activity essentially identical to that of the WT
enzyme.
Because mutation of residues near the active-site of the
protease could result in a change in overall substrate binding,
we next decided to assess the specificity of the T184C mutant
protease using a peptide library method.19 This method
involves incubating the protease with a diverse pool of peptides
and then measuring the most optimal substrates by sequencing
Live Cell Labeling and Immunofluorescence. Cells were
transfected with lipofectamine and seeded in eight-well chamber slides
(Lab-tek Nunc) at a density of 5 × 104 cells/well. After 24 h, they were
incubated with 1.5 μM TND124 or TND126 for 1 h at 37 °C,
followed by multiple washes with media. The monolayer was washed
with PBS and fixed with 4% paraformaldehyde for 20 min at room
temperature. The cells were then permeabilized with PBS, 0.1% Triton
X-100, and 1% heat-inactivated goat serum for 1 h. Next the cells were
washed with PBS and incubated with anti-HA tag (1:100; Covance
clone 16B12) primary antibody in blocking buffer (PBS with 1% heat-
inactivated goat serum) for 1 h at room temperature. The cells were
then washed with PBS and incubated with AlexaFluor 488 conjugated
secondary antibody (1:1000, Invitrogen) for 30 min. The cells were
treated and mounted using Vectashield with DAPI (Vector
laboratories) for imaging using a Zeiss LSM700 confocal microscope.
In the case of pre-incubation with a general MMP inhibitor, cells were
treated with 10 μM GM6001 for 30 min, washed, and incubated with
the probe TND124 (1.5 μM) as described.
Zebrafish. Fish Strain and Maintenance. Wild-type zebrafish
(Danio rerio) were maintained under standard laboratory conditions.
Embryos were collected from natural matings, reared at 28.5 °C, and
staged according to age and morphology as described previously.24
Morpholino Injections, Rescue, and Alcian Blue Staining. The
mt1-mmpa MO and WT mt1-mmpa pCS2 clone were previously
described.25 The F260C mutant mt1-mmpa pCS2 construct was
D
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structure of a hydroxamate-based inhibitor bound to the active-
site of MMP8 (Figure S2, Supporting Information), we
designed a probe, TND124, that contains the hydroxamic
acid group and primary P1′ recognition group of GM6001 to
drive tight binding in the active-site. Furthermore, we
incorporated an α-chloroacetamide electrophile positioned at
the P2′ position and a Cy5 fluorescent tag in the P3′ position
(Figure 2b). Based on docking of this probe into the active-site
of MMP8, the electrophile was predicted to lie within 1.8 Å of
the engineered cysteine (Figure S2, Supporting Information).
The electrophile was selected due to its previously reported
restricted reactivity profile toward cysteines.30 We designed a
synthetic route that comprised standard peptide chemistry to
build the main scaffold and click chemistry to introduce the
electrophile at the end of the synthesis (Supporting
Information).
To determine if the probe was able to covalently label the
engineered MMP12, we performed mass spectrometry analysis
of the WT and T184C MMP12 after treatment with the probe.
Analysis of intact mass confirmed that TND124 formed a single
covalent modification of the engineered cysteine (Figure S3,
Supporting Information). Tryptic digestion of the labeled
protein further confirmed that the peptide containing the
engineered cysteine shifted by a mass consistent with the
addition of a single TND124 molecule (an increase of 1193
amu), thus demonstrating specific modification of the
engineered cysteine. We next needed to determine if this
confirmed modification was specific for the target protein by
performing a labeling study in complex protein extracts.
Therefore, we added either mutant or WT MMP12 to crude
tissue extracts, incubated with TND124, and monitored probe
labeled proteins by SDS-PAGE followed by fluorescence
scanning (Figure 2c). Under these conditions, TND124
showed highly selective labeling of the cysteine-engineered
catalytic domain with negligible background labeling when used
at low probe concentrations (<1 μM). Importantly, we did not
observe labeling of the WT target, and pretreatment of the
samples with the general MMP inhibitor, GM6001, completely
blocked labeling, indicating that probe binding to the
engineered protease was dependent on enzymatic activity as
well as the cysteine residue.
Finally, we sought to determine if the covalent labeling of the
engineered protease resulted in irreversible inhibition of
enzyme activity. We therefore performed a kinetic inhibition
study using a fluorogenic substrate (Figure S4a, Supporting
Information). These data confirmed that TND124 inhibited the
T184C mutant with increased potency relative to the WT
enzyme (IC50 = 0.12 0.01 μM for T184C versus IC50 = 0.75
0.05 μM for WT). More importantly, the T184C protease,
unlike the WT form, was unable to regain activity after removal
of the probe by dialysis (Figure 2d). We also obtained similar
results for natural protein substrate processing, where only the
activity of the engineered enzyme against collagen or gelatin
was inhibited (Figure S4b,c, Supporting Information). Finally
irreversible inhibition of only the engineered enzyme was
confirmed using casein zymography (Figure 2e). Overall these
results demonstrate that, while the probe is a weak, reversible
inhibitor of the WT enzyme, it acts as a potent irreversible
inhibitor of the engineered cysteine variant of the enzyme.
Engineering MT1-MMP. After demonstrating the feasi-
bility of our approach using MMP-12, we wanted to see if this
method was generalizable across the MMP family. We therefore
chose to apply our strategy to one of the most intensely studied
Figure 2. Engineering MMP12 and the corresponding selective probe
TND124. (a) Progress curves for processing of a quenched fluorescent
peptide substrate [Mca-PLGL-Dpa-AR-NH2, where MCA (7-methox-
ycoumarin-4-yl)acetate) is the fluorophore and Dnp (dinitrophenyl) is
the quencher] by WT, T184C, T243C, and T184CT243C MMP12
catalytic domains. (b) Structure of the general MMP inhibitor
GM6001, the designed probe TND124, and the control probe
TND126. The electrophile (α-chloroacetamide) is shown in blue, the
electrophile coordinating the zinc (hydroxamate) in TND124 or the
control methyl amide in TND126 is shown in red, and the fluorescent
tag (Cy5) is shown in green. (c) WT or T184C MMP12 catalytic
domains were added to rat liver lysates. The general MMP inhibitor
GM6001 (10 μM) was added prior to labeling of the samples by
addition of TND124 at the indicated concentrations. Samples were
analyzed by SDS-PAGE followed by scanning of the gel for Cy5
fluorescence using a flatbed laser scanner. The location of the labeled
MMP12 catalytic domain is indicated with an arrowhead. (d)
Recombinant WT or T184C MMP12 catalytic domains were
incubated with TND124 for 30 min followed by dialysis for 48 h.
The residual activity of both proteases was measured after 24 and 48 h.
Data are presented as average of activity from an experiment
performed in triplicate (error bars
SD, bottom panel). (e)
Recombinant WT MMP12 and T184C MMP12 catalytic domains
were incubated with DMSO (−) or 3 μM TND124 (+), and activity
was detected by casein zymography. The white signals are produced by
degradation of the substrate by active MMP12. Note the specific
inhibition of this activity only in the T184C sample treated with
TND124.
the cleaved peptides (Figure S1, Supporting Information).
These results indicated that the T184C mutant had substrate
selectivity nearly identical to that of the WT enzyme with only
very minor changes in P2′ position, where we observed a
modest preference for phenylalanine, leucine, methionine, and
tyrosine residues. Since the majority of substrate specificity of
the MMP family is derived from the P1′ position as well as
from binding of the substrate to the carboxy-terminal
hemopexin domain, the T184C mutant is likely to have a
substrate specificity virtually identical to that of the WT
enzyme.
Engineering of the Probe. In order to design a suitable
probe that contains an electrophile in close proximity to the
engineered cysteine residue, we started with the broad
spectrum MMP inhibitor GM6001 that has been used to
make photo-cross-linkable probes10 (Figure 2b). Using the
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members of the MMP family, MT1-MMP (MMP-14). This
protease is a transmembrane protease that has been shown to
be involved in the regulation of cell migration in processes such
as tumor growth and metastasis.31,32 Because of the similarity in
the catalytic domain of all of the MMP family members, we
were able to focus on residues corresponding to the locations of
the mutants tested in MMP12. Therefore, we generated
catalytically active WT MT1-MMP as well as the F198C and
F260C mutants. We also tried a third mutation site, Q262. As
observed for MMP12, both the F198C and F260C mutants
were proteolytically active, with the F260C mutant being closer
in activity level to the WT enzyme (Figure 3a). Interestingly,
compared to the WT enzyme, with only very minor changes in
the P1′ position (Figure S5, Supporting Information).
Finally, we checked the selectivity of the probe for the
engineered mutant by labeling the recombinant protein in
tissue extracts (Figure 3c). As observed for the MMP12
mutants, TND124 showed robust labeling of the F260C
mutant protein with no detectable labeling of the WT enzyme.
In addition, pre-incubation of the samples with a cysteine
alkylating agent (N-ethylmaleimide, NEM) or a chelating agent
(EDTA) confirmed that the probe required the engineered
cysteine as well as the zinc in the active-site for effective
labeling. We also generated a control probe (TND126) lacking
the hydroxamate group but containing the α-chloroacetamide
group (Figure 2b). This probe did not label the F260C MT1-
MMP (Figure S6, Supporting Information), nor did it block
labeling of the enzyme by TND124 (Figure 3c), confirming
that the zinc coordination by the hydroxamate anchors the
probe in the active-site to facilitate irreversible modification of
the enzyme. Consistent with this result, kinetic inhibition
studies using the peptide substrate confirmed that TND124
inhibited the activity of F260C MT1-MMP (IC50 = 1 0.1 μM
for the mutant protease versus IC50 = 2.8 0.8 μM for the WT
enzyme; Figure 3d).
Functional Validation of the Cysteine-Engineered
MT1-MMP in Cells. To determine if the newly identified
cysteine mutant form of MT1-MMP could be synthesized,
folded, and correctly trafficked to the cell surface, we
transfected the full-length WT and F260C mutant MT1-
MMP in MCF-7 cells. These cells do not display proteolytic
activity against collagen substrates and are devoid of tissue-
invasive behavior.33 However, upon expression of MT1-MMP,
the cells have been shown to hydrolyze subjacent substrates
(e.g., native type I collagen as well as heat-denatured type I
collagen/gelatin) and to display collagen-invasive activity.33
Therefore, we performed gelatin degradation and collagen
invasion assays with MCF7 cells expressing either WT or
F260C MT1-MMP. Using either construct, the MCF-7 cells
degraded the underlying substrate while also showing
comparable invasive activity, thereby demonstrating that the
F260C mutant is expressed in an active form that is capable of
degrading native substrates (Figure S7, Supporting Informa-
tion).
To further test the function of the F260C variant, we
measured the ability of cells expressing the mutant to process
the native MT1-MMP substrate, pro-MMP2, to its active form.
To this end, we collected conditioned medium from cells that
secreted pro-MMP2, incubated it with HEK293T cells
expressing either the WT or the F260C variant MT1-MMP,
and then measured the extent of pro-MMP2 processing to the
mature MMP2 using gel zymography (Figure 4b). These results
confirmed that both the WT and the F260C variant cleaved
pro-MMP2, while neither the non-transfected MCF-7 cells nor
MCF-7 cells expressing the proteolytically inactive, catalytic site
mutant, E240A MT1-MMP, were unable to process the
zymogen.
Figure 3. Engineering MT1-MMP. (a) Progress curves for processing
of a quenched fluorescent peptide substrate (Mca-PLGL-Dpa-AR-
NH2) by WT, F198C, F260C, and Q262C MT1-MMP catalytic
domains. (b) Kinetic analysis of the enzyme activity of WT and F260C
MT1-MMP using the fluorogenic substrate in (a). Data are presented
as the average of three independent experiments (error bars
standard deviation [SD]). (c) Recombinant WT or F260C MT1-
MMP catalytic domains were added to rat liver lysate followed by pre-
incubation with DMSO, GM6001 (10 μM), EDTA (10 mM), or NEM
(1 mM). Samples were then labeled with TND124 (1 μM) and
analyzed by SDS-PAGE followed by in-gel fluorescent scanning for
Cy5. The location of the MT1-MMP catalytic domain is indicated. (d)
Recombinant WT or F260C MT1-MMP catalytic domains were
incubated with TND124 over a range of concentrations, and the
activity of the enzyme against the MCA-PLGL-Dpa-AR-NH2 substrate
was measured. Data are plotted as percent activity relative to enzyme
incubated with DMSO (IC50 = 1 0.1 μM for the mutant protease
versus IC50 = 2.8 0.8 μM for the WT enzyme). Data are presented
as average of three independent experiments (error bars SD).
the Q262C mutant showed a complete loss of protease activity,
highlighting the fact that some residues near the active-site
cannot be mutated without affecting activity. Detailed enzyme
kinetic analyses revealed that F260C had a 2-fold higher Km
than the WT protease but a similar Vmax (Figure 3b), indicating
that the F260C mutant can reach the same catalytic rate as the
WT enzyme. The specificity of F260C MT1-MMP was further
tested using the peptide substrate library, and we found that,
like MMP12, it had nearly identical primary substrate selectivity
Finally, to determine if the engineered cysteine mutant MT1-
MMP could be specifically inhibited, we treated cells expressing
the different MT1-MMP variants with the probe and, after
various washouts, plated the cells atop a gelatin-coated matrix.
Under these conditions, the probe only inhibited gelatin
degradation in cells expressing the mutant enzyme, thus
demonstrating that TND124 is selective for the engineered
form of MT1-MMP (Figure 4c).
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Figure 4. The engineered MT1-MMP is expressed and is functional in
cells. (a) MCF7 cells were transfected with full-length WT or F260C
mutant MT1-MMP and then plated on a type-I collagen matrix. A
typical image is shown to indicate the foci that are formed when the
cells invade through the matrix (marked by an arrowhead). These foci
were counted, and the average number of invasive foci per field was
plotted for WT and F260C expressing cells (error bars SD). (b)
Zymography showing the activation of proMMP2 to MMP2 by media
from cells expressing WT or F260C MT1-MMP. (c) Cells that were
transfected with WT or F260C MT1-MMP were incubated with
TND124 at the indicated concentrations, washed, and plated onto a
layer of fluorescently labeled gelatin. Typical sites of degradation
(highlighted by circles) are observed as dark patches in the surface.
Specific Imaging of Cysteine-Engineered MT1-MMP
Activity in Cells. To further validate our protein engineering
strategy, we tested the ability of the probe to specifically label
the active pools of the F260C variant MT1-MMP in an intact
cell system. Biochemical analysis confirmed labeling of the
mature, active form of MT1-MMP (i.e., the 55 kDa mature
form of the protease generated after prodomain processing1)
only in lysates from cells expressing the F260C variant (Figure
5a). Labeling was also specifically competed away by pre-
incubation with GM6001, suggesting that the labeling was
specific for the active cysteine-engineered variant.
To assess the selectivity of labeling, we treated HEK293T
cells that were transiently transfected with HA-tagged WT or
F260C MT1-MMP with TND124 for 1 h. Following extensive
washing, the cells were fixed, stained with anti-HA tag antibody,
and visualized by microscopy. Importantly, the probe showed
highly specific labeling of cells that were positive for expression
of the F260C variant of MT1-MMP (Figure 5b), with no signal
detected in untransfected cells or cells expressing WT or the
proteolytically inactive E240A mutant. In addition, pre-
incubating the cells with GM6001 (Figure S8, Supporting
Information) or using a negative control probe, TND126
(Figure S9, Supporting Information), confirmed that the
labeling was specific to active pools of the engineered MT1-
MMP variant. The probe signal was mainly localized to
intracellular, endosomal-like compartments, as well as to
cellular protrusions. This could be the result of internalization
of the MT1-MMP protein after surface labeling, similar to what
has been reported when the WT protease is expressed in intact
cells.34 In support of this, when we lowered the incubation
temperature to 4 °C to slow endocytosis, we found that MT1-
MMP labeling was confined primarily to the cell surface (Figure
5b).
Figure 5. Specific labeling and imaging of the engineered MT1-MMP
in cells. (a) Cell lysates of untransfected HEK293T cells (control) or
cells expressing the full-length WT or F260C MT1-MMP were pre-
treated with GM6001 (+) or DMSO (−), followed by labeling with
TND124 (0.5 μM). Samples were analyzed by SDS-PAGE followed by
in-gel fluorescence scanning for Cy5. The location of the mature 55
kDa MT1-MMP is indicated. (b) HEK293T cells were transfected
with full-length WT, F260C, or catalytically inactive mutant E240A
MT1-MMP (all tagged with HA tag) and incubated with TND124
(red, 1.5 μM) at 37 °C or at 4 °C to block re-uptake of the expressed
MT1-MMP (bottom row). Cells were washed, fixed, and stained with
an anti-HA antibody (green) for MT1-MMP and DAPI nuclear stain
(blue). Scale bars are 20 μm.
Knockdown of MT1-MMPa/b using anti-sense morpholino
oligonucleotide (MO)25 was shown to induce a craniofacial
morphogenesis phenotype. Importantly, this phenotype could
be rescued by co-injection of the MO with synthetic mt1-mmp
mRNA. Therefore, we could determine if the F260C variant
MT1-MMP was functional in the context of a whole organism
by assessing its ability to rescue the knockdown phenotype.
When synthetic F260C mt1-mmpa mRNA was co-injected with
a MO directed against the zebrafish MT1-MMP, we observed a
dramatic reduction in the number of fish with the morphant
phenotype (Figure 6a). Importantly, the F260C mutant gene
rescued the phenotype to the same extent as the wt mt1-mmpa,
suggesting that the cysteine-engineered variant is functionally
competent even when expressed in a whole organism (Figure
6b).
Functional Expression and Imaging of Cysteine-
Engineered MT1-MMP in Zebrafish. As a final confirmation
of the MT1-MMP mutant to retain normal functional activity in
vivo, we expressed the mutant gene in zebrafish. The zebrafish
genome encodes two maternally expressed isoforms homolo-
gous to human MT1-MMP: MT1-MMPa and MT1-MMPb.25
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in embryos that had been injected with the F260C variant
MT1-MMP. These data suggest that the probe is sufficiently
selective and stable in vivo for imaging studies of MT1-MMP
activity in whole fish.
DISCUSSION
■
We present here a general method that can be used to
specifically target a protease by combining the use of protein
engineering and small-molecule design. One of the major
limitations in collective efforts taken to date to understand
protease function is the difficulty in developing small molecules
that can be used to specifically image or inhibit a single protease
target in the context of a whole cell or organism. In the case of
the matrix metalloprotienases, there are a number of hurdles
including the lack of a reactive nucleophilic residue in the
active-site and the high degree of similarity in the substrate and
inhibitor binding sites within the catalytic domains of this
multigene family. Therefore, methods that permit the use of a
small molecule to inhibit or image a single MMP target with
absolute specificity should prove transformative in terms of our
ability to characterize the functions of these important
proteases.
Here we show that it is possible to identify a site on MMPs
that can be mutated to cysteine without dramatically altering
enzyme activity. Importantly, these cysteine variants can be
expressed in cells where they function with efficiency similar to
that of WT enzymes, supporting the contention that these
mutations do not interfere with protein folding or cellular
trafficking. In the case of MT1-MMP, the specificity of the
mutant protease (as characterized by substrate library
hydrolysis) was slightly modified in the P1′ position. However,
we find that the mutant protease could process natural
substrates and replace the WT protease in a complex organism
like zebrafish. This result is likely explained by the fact that
MT1-MMP is a multidomain protease and its substrate
preference and binding are dependent not only on the
interactions of the substrate within the shallow active-site
pocket but also within exosites located in other domains, such
as the hemopexin domain.35
We have designed a first-generation probe containing a
reactive electrophile that specifically labels the cysteine mutant
protease. Furthermore, the probe acts as a potent, irreversible
inhibitor of the engineered protease but only a weak reversible
inhibitor of the WT enzyme. Because the probe directly labels
only the active form of the cysteine-engineered protease, it can
be used to both specifically block activity and also image
localization of the active protease. Therefore, this new method
will allow studies of the relationship between MT1-MMP
cellular trafficking and the regulation of its proteolytic activity,
something that has been studied only using indirect methods.
This strategy could be particularly valuable for studies of the
role of MT1-MMP in signaling pathways (i.e., Wnt signaling
pathway) or in remodeling of the extracellular matrix (ECM)
during embryogenesis.
Figure 6. Expression and imaging of MT1-MMP in zebrafish. (a)
Wild-type, day 5 zebrafish embryos in which expression of the mt1-
mmpa gene was knocked down with specific antisense morpholinos
(mt1-mmpa MO) and day 5 embryos in which the mmp14a MO was
injected along with mRNA expressing the WT (mt1-mmpa MO + WT
mRNA) or F260C mutant mt1-mmpa (mt1-mmpa MO + F260C
mRNA) were stained with alcian blue to visualize cartilage elements.
(b) Total number of fish observed with WT, mild mutant, and severe
mutant phenotypes were determined and plotted for fish rescued with
the WT or F260C mutant mt1-mmpa mRNA. (c) Zebrafish embryo at
the single-cell stage were either not injected (no probe), injected with
the TND124 probe (2 nL of 50 μM DMSO stock) alone (no mRNA),
or injected with probe and 300 pg of WT (WT MT1-MMPa) or
F260C mutant (F260C MT1-MMPa) mmp14a mRNA. Embryos were
fixed, washed, and imaged for Cy5 fluorescence at either the 8- or 18-
somite stage. Images are from representative embryos selected from a
total of ∼20 embryos that gave similar results. Scale bars are 100 μM
for full images and 25 μM for inset images.
As a final test of the method, we wanted to determine if the
probe could be used to label active pools of the engineered
cysteine MT1-MMP in the whole zebrafish embryo. Therefore,
we injected embryos at the single-cell stage with the mRNA for
either the WT or F260C variant of MT1-MMP and then fixed,
washed, and imaged the embryos at the 8-somite stage (Figure
6c). While there was some level of background fluorescence
both in embryos that had not been injected with mRNA and in
embryos injected with WT MT1-MMP mRNA, we were
consistently able to observe specific accumulation and staining
While the TND124 probe has proven to be valuable for
imaging and inhibition studies of MMP-12 and MT1-MMP, we
believe that it will be possible to further improve the utility of
next-generation probes by modulating overall reactivity of the
electrophile as well as potency of the main probe scaffold for
specific targets. In all of our lysate labeling studies, we observed
a background signal at higher probe concentrations that is likely
due to the reactivity of the α-chloroacetamide. Regardless, we
are encouraged by the fact that the probe produces very
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minimal background signals when used to label intact cells that
either do not express the engineered protease or express the
catalytically inactive protease. This discrepancy in background
signal in lysates relative to intact cells is likely a result of
reduced probe access to intracellular protein targets and the
distinct conditions of the labeling reaction when used in cell
culture compared to lysates. Therefore, labeling specificity
should be assessed in both systems in order to more clearly
assess probe selectivity.
National Institutes of Health grants R01-EB005011 and R21-
EB012311 (to M.B.). M.M. was supported by AGAUR through
the Beatriu de Pinos program. Y.D. was supported by a James
Hudson Brown-Alexander Brown Coxe postdoctoral fellowship
from Yale University School of Medicine.
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ASSOCIATED CONTENT
* Supporting Information
Supplementary methods, synthetic schemes, Table S1, and
Figures S1−S9. This material is available free of charge via the
Internet at http://pubs.acs.org.
■
S
AUTHOR INFORMATION
Corresponding Author
mbogyo@stanford.edu
Present Addresses
■
⊥J.L.: Department of Global Medical Science, Sungshin
Women’s University, Seoul 142-732, Korea
#E.D.: MRC National Institute for Medical Research Division
of Parasitology, The Ridgeway, Mill Hill, London NW7 1AA,
UK
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
We thank the Bogyo lab for useful discussions and specifically
Aimee Shen for help in cloning, PCR-based mutagenesis, and
protein expression and Andrew Guzzetta for help with the mass
spectrometry analysis of full-length MMPs. We thank Chris
Overall and Charlotte Morrison for the TIMP2-deficient cells
and the zymography protocols and Dimitri Rozanov for the
full-length MT1-MMP plasmid. This work was supported by
I
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