Peptide Length and Leaving-Group Sterics Influence Potency of Peptide Phosphonate Protease Inhibitors

Article (PDF Available)inChemistry & biology 18(1):48-57 · January 2011with25 Reads
DOI: 10.1016/j.chembiol.2010.11.007 · Source: PubMed
Abstract
The ability to follow enzyme activity in a cellular context represents a challenging technological frontier that impacts fields ranging from disease pathogenesis to epigenetics. Activity-based probes (ABPs) label the active form of an enzyme via covalent modification of catalytic residues. Here we present an analysis of parameters influencing potency of peptide phosphonate ABPs for trypsin-fold S1A proteases, an abundant and important class of enzymes with similar substrate specificities. We find that peptide length and stability influence potency more than sequence composition and present structural evidence that steric interactions at the prime-side of the substrate-binding cleft affect potency in a protease-dependent manner. We introduce guidelines for the design of peptide phosphonate ABPs and demonstrate their utility in a live-cell labeling application that specifically targets active S1A proteases at the cell surface of cancer cells.
Chemistry & Biology
Article
Peptide Length and Leaving-Group Sterics Influence
Potency of Peptide Phosphonate Protease Inhibitors
Christopher M. Brown,
1,5
Manisha Ray,
1,5
Aura A. Eroy-Reveles,
2
Pascal Egea,
3,6
Cheryl Tajon,
4
and Charles S. Craik
2,
*
1
Graduate Group in Biochemistry and Molecular Biology
2
Department of Pharmaceutical Chemistry
3
Department of Biochemistry and Biophysics
4
Chemistry and Chemical Biology Graduate Program
University of California, San Francisco, CA 94158, USA
5
These authors contributed equally to this work
6
Present address: Department of Biological Chemistry, David Geffen School of Medicine, University of California, Los Angeles, CA 90095
*Correspondence: craik@cgl.ucsf.edu
DOI 10.1016/j.chembiol.2010.11.007
SUMMARY
The ability to follow enzyme activity in a cellular
context represents a challenging technological
frontier that impacts fields ranging from disease
pathogenesis to epigenetics. Activity-based probes
(ABPs) label the active form of an enzyme via cova-
lent modification of catalytic residues. Here we
present an analysis of parameters influencing
potency of peptide phosphonate ABPs for trypsin-
fold S1A proteases, an abundant and important class
of enzymes with similar substrate specificities.
We find that peptide length and stability influence
potency more than sequence composition and
present structural evidence that steric interactions
at the prime-side of the substrate-binding cleft affect
potency in a protease-dependent manner. We intro-
duce guidelines for the design of peptide phospho-
nate ABPs and demonstrate their utility in a live-cell
labeling application that specifically targets active
S1A proteases at the cell surface of cancer cells.
INTRODUCTION
The creation and implementation of high-throughput nucleic acid
analysis techniques have revolutionized medicine and biology.
The developments of affinity capture and protein binding arrays
have allowed similar analyses of protein levels and interactions.
However, such gene and protein profiling data does not always
reflect the dynamic milieu that one finds at the cellular level
in vivo. Furthermore, enzymatic activity adds an additional layer
of complexity in functionality that cannot currently be addressed
using available techniques.
The need to examine enzyme activity is particularly relevant to
the study of proteolytic enzymes in biological systems. Although
synthetic substrates can be used to study individual proteases
in vitro, their efficacy in studying multiple proteases in a complex
mixture is limited. For example, the caspase family of cysteine
proteases have highly overlapping substrate specificities, thus
multiple caspases can often cleave the same synthetic substrate
(McStay et al., 2008). This complicates functional analyses of
individual caspases during apoptosis. However, covalent
labeling of active proteases allows one to monitor individual
proteases in the context of the whole cell. Technologies based
on covalent inhibitors are emerging for activity-based proteo-
mics. Phosphonate inhibitors of serine proteases form the basis
of one such technology (Sienczyk and Oleksyszyn, 2009).
Diisopropylfluorophosphonate (DFP) is one of the earliest
irreversible inhibitors described for serine proteases, and radio-
labeled DFP was one of the first activity-based probes (ABPs)
described (Powers et al., 2002). More recent studies have
modified DFP to include a detection tag for purification and/or
visualization purposes (Liu et al., 1999). Biotin and fluorophores
are the most common tags used, and active enzyme profiles of
several types of human tissues and tumor types have been
created using such molecules (Jessani et al., 2005). Although
these efforts are valuable to the field, these studies identify
a preponderance of serine hydrolases that are not proteases,
and ABPs specific for proteases have been difficult to achieve.
The specificity of DFP can be improved in two ways. Replacing
the highly electronegative fluorine atoms with two phenoxy
groups reduces reactivity and increases stability (Powers et al.,
2002). Further modifying these functional groups modulates
the electrophilicity and shape of the phosphonate reactive
center, which can add selectivity to the compound. For example,
incorporating different electron withdrawing or donating groups
onto the phenyl rings of diphenyl phosphonates (DPPs) results in
differential specificity between the urokinase-type plasminogen
activator (uPA) and trypsin (Sienczyk and Oleksyszyn, 2006).
A second way to reduce DFP promiscuity is to include a short
peptide, which helps the inhibitor bind to the active site of the
targeted protease. By utilizing substrate cleavage data and
combinatorial peptide libraries, the peptide sequence can direct
the inhibitor toward a target protease or group of proteases (Lim
and Craik, 2009). This methodology was used to design specific
ABPs for Granzymes A and B (Mahrus and Craik, 2005).
Despite the aforementioned tunable properties, the emer-
gence of peptide phosphonates as the premier ABPs for specific
serine proteases has yet to occur. One can imagine a platform
where peptide phosphonates are incorporated into existing
microarray technology to create high-throughput assays to
48 Chemistry & Biology 18, 48–57, January 28, 2011 ª2011 Elsevier Ltd All rights reserved
monitor active proteases at the bench or in the clinic. However,
such a technology has yet to be developed, and the utility of
peptide phosphonates has come into question.
Here we describe the synthesis, evaluation, and application of
peptide phosphonate inhibitors designed to target S1A family
proteases. This subfamily of Clan PA proteases contains the
trypsin-like enzymes, and it is the largest family of proteases in
higher eukaryotes (Rawlings et al., 2010). These proteases play
many important roles in biology and disease progression, and
thus are of interest to many fields. This work examines the effects
of peptide sequence and length on inhibition of two similar S1A
proteases, thrombin and MT-SP1/matriptase, via enzymatic,
structural, and imaging methods. We find that peptide length
and leaving group sterics are large determinants of potency,
whereas sequence composition contributes to a lesser degree.
Our findings suggest general guidelines for the design of phos-
phonate ABPs that are optimized for S1A proteases. By applying
these guidelines, we demonstrate that peptide phosphonates
can quantitatively label and follow proteases on the surface of
cancer cells, a novel use of ABPs that can be applied to many
systems.
RESULTS
Improvement to the Synthesis of Peptide Phosphonates
Several modifications to published protocols have improved the
synthesis of the diphenyl phosphonate ester of 4-amidino-
phenyl-glycine (H-(AmPhg)
P
(OPh)
2
, for brevity referred to herein
as H-Bz-DPP, 12) and the final biotinylated peptide phosphonate
probes (Oleksyszyn et al., 1994). The synthesis of 12 (Figure 1)
has been reported previously, though isolation of pure
compound has been difficult. Briefly, compounds 911 were
synthesized as described in (Mahrus and Craik, 2005; Oleksys-
zyn et al., 1994). Published methods describe isolation of 11
by diethyl ether precipitation, however, under the basic reaction
conditions to produce 11, the phosphonate center is not stable.
Hydrolysis of one of the phenyl ester groups can occur and be
replaced with either an ethyl or methyl ester adduct. Reducing
reaction c from 5 to 2 days prevented accumulation of the side
product as monitored by LC-MS. Longer incubations increased
levels of the undesired product. Direct hydrogenation of 11
followed by high-performance liquid chromatography (HPLC)
produced pure 12. Alternatively, 11 and 12 were separable by
silica gel flash chromatography (10:1 CH
2
Cl
2
:MeOH and 5:1
CH
2
Cl
2
:MeOH with ninhydrin staining, respectively) at R
f
z 0.3.
Improvements to the synthesis of the final biotinylated peptide
diphenyl phosphonate probes include (1) separate construction
of the biotinylated peptide moiety; and (2) optimization of the
coupling reaction of the peptide to 12 to increase yield and
reduce reaction time. Instead of building the probe by extending
12 one amino acid at a time as described in Boduszek et al.
(1994) and Jackson et al. (1998), biotinylated peptides were
synthesized on 2-chlorotrityl chloride resin by standard Fmoc
chemistry, cleaved under mild acidic conditions to retain the
protecting groups, and coupled to the phosphonate. Previous
reports indicate this phosphonate coupling reaction to be time
consuming (12–48 hr) and low yielding (5%–30%)(Oleksyszyn
et al., 1994). Optimization of this reaction proceeded with
a TFE:CH
2
Cl
2
(2:5) solvent mixture to reduce the reaction time
from overnight to 6 hr, where N,N-dimethylformamide (DMF)
had been used previously. Treatment with the coupling agent
EDAC resulted in the greatest yield of product (Table 1)as
compared to PyBOP or DCC (!20%). EDAC protects the
base-sensitive phosphonate from hydrolysis because it is
typically utilized in the pH range of 4–6. After the coupling reac-
tion, deprotection of the amino acid side chains followed. HPLC
purification was applied to both the protected and deprotected
biotinylated peptide phosphonates. As a result, a combination
of TFE:CH
2
Cl
2
and EDAC resulted in a fast, efficient coupling
that protected the integrity of the phosphonate center and
improved product yield (Table 1).
CN
HO
CN
P
H
N
Cbz
O
OPh
OPh
P
H
N
Cbz
O
OPh
OPh
P
H
N
Cbz
O
OPh
H
2
N NH
PH
2
N
O
OPh
OPh
H
2
N NH
H-Bz-DPP
12
A
B
C
D
0198
11
NHEtO
OPh
P
H
N
O
OPh
OPh
H
2
NNH
R
1 R = Biotin-PEG-QRV
2 R = Biotin-PEG-LTP
3 R = Biotin-PEG-GSG
4 R = Biotin-PEG-EPI
5 R = Biotin-PEG
6 R = Biotin-PEG-V
7 R = Biotin-PEG-RV
13 R = Biotin-PEG-FTGSG
E
R-COOH
Figure 1. General Synthesis of 4-Amidinophenyl-Glycine that Incorporates Benzamidine at the P1 Position
Reagents and conditions: (A) P(OPh)
3
, benzyl carbamate, HOAc, 1 hr at room temperature, 1 hr at 85
"
C; (B) dry EtOH, CHCl
3
, HCl in dioxane, 5 days at 4
"
C; (C)
NH
3
in dioxane, dry EtOH/dioxane (1:1), 2 days at room temperature; (D) H
2
/Pd, HCl, EtOH, 6 hr at room temperature; (E) EDAC, HOBt, TFE:DCM, 6 hr at room
temperature. See also Table S1.
Table 1. Yields and Masses of Each Phosphonate Probe
Synthesized
Compound (biotin-PEG-R) Yield (%) m/z Calculated (Found)
R = QRVBz-DPP (1) 72 1308.61 (656.34)
R = LTPB z-DPP (2) 56 1236.57 (1237.12)
R = GSGBz-DPP (3) 65 1126.46 (1126.92)
R = EPIBz-DPP (4) 57 1264.56 (1266.31)
R = Bz-DPP (5) 25 925.38 (926.7)
R-VBz-DPP (6) 20 1024.45 (1025.82)
R = RVBz-DPP (7) 77 1180.55 (1181.64)
R = FTGSGBz-DPP (13) 40 1374.57 (1376.27)
Chemistry & Biology
Potency Determinants of Peptide Phosphonates
Chemistry & Biology 18, 48–57, January 28, 2011 ª2011 Elsevier Ltd All rights reserved 49
Increasing Peptide Length Improves Phosphonate
Inhibition of Serine Proteases
Having optimized synthesis of the phosphonate inhibitors, the
inhibitors were next characterized in vitro. Previous work using
positional scanning, synthetic combinatorial library (PS-SCL)
profiling had identified RKSR as the preferred P4-P1 tetrapep-
tide sequence for MT-SP1 and LTPR as the preferred sequence
for thrombin (Bhatt et al., 2007; Harris et al., 2000). Combining
this data with other validated substrates and structural informa-
tion led to the creation of a series of peptide phosphonates that
were designed to target MT-SP1 and thrombin (Table 1). Two
sequences, QRVBz (1) and LTPBz (2) were rationally designed
to maximize specificity for the two S1A proteases. A peptide
element designed to be moderately effective against both prote-
ases (GSGBz 3) was also synthesized, as was a fourth sequence
(EPIBz 4) containing suboptimal amino acids for both proteases
at each P2-P4 position. Thus, a series of peptides were incorpo-
rated to create ABPs that were predicted to cover a range of
activities against both MT-SP1 and thrombin.
IC
50
values were calculated for each inhibitor against both
MT-SP1 and thrombin, and k
inact
/K
I
values were calculated for
inhibitors of a representative subset against MT-SP1 (Table 2).
For MT-SP1, both the IC
50
and k
inact
/K
I
values for the tetrapep-
tide inhibitors trended as expected, with the optimal sequence
QRVBz-DPP (1) inhibiting best at 0.37 mM, and the nonoptimal
sequence EPIBz-DPP (4) inhibiting worst at 76 mM(Table 2,
sections A and D). These values were in agreement with k
cat
values measured for synthetic fluorogenic substrates with
sequences corresponding to each inhibitor (see Table S1 avail-
able online). This correlation between k
cat
and inhibitory potency
has been previously observed (Drag et al., 2010).
Each of the peptide phosphonates tested proved to be slow
inhibitors of MT-SP1. The fastest inhibitor, QRVBz-DPP (1),
was only 200 M
#1
s
#1
. Previous studies have reported values
approaching 37,000 M
#1
s
#1
for phosphonate inhibition of serine
proteases (Powers et al., 2002). The data shown here indicates
that diphenyl phosphonate ABPs did not inhibit MT-SP1 rapidly,
and either long incubation times or high phosphonate concentra-
tions were needed to reach complete inhibition. Although
optimizing the amino acid composition of the peptide improved
inhibition, we were unable to gain a large degree of selectivity.
The best inhibitor for MT-SP1, 1, was better than optimal inhib-
itors designed against thrombin or uPA, another S1A protease
(data not shown), further evidence suggesting that sequence
composition plays a minor role in inhibitory potency for certain
proteases.
The minor contribution due to sequence is reiterated by the
ability of the peptide phosphonates to label proteases in vitro.
The inhibitors were incubated with MT-SP1 and thrombin at
increasing concentrations for 16 hr, and biotin-labeled proteases
were detected via a western blot using avidin-HRP. Figure S1
shows that the ability of the ABPs to label both proteases was
also not strongly dependent on sequence composition.
GSGBz-DPP (3), designed to confer intermediate potency for
MT-SP1 and thrombin, labeled more efficiently than the ABPs
with optimal sequences for each protease. This observation is
partly explained by the greater stability of 3 over time in aqueous
solution (Table S1), indicating that sequence stability was an
additional parameter for labeling efficiency. Additionally, QRVBz,
the best kinetic inhibitor of MT-SP1, contains a second cleavage
site after the P3 Arg. This cleavage removed the biotin from the
ABP, rendering it undetectable by avidin-HRP and complicating
analysis. This secondary cleavage event is evidenced by the
appearance of an 846Da fragment on postincubation LC-MS
analysis.
In addition to the ABPs described above that were designed to
test the effects of amino acid sequence on inhibition, three addi-
tional ABPs were synthesized to test the effects of peptide length
on potency. Previous reports have noted a dependence of
peptide length on the mechanistic rate constant k
2
, which
suggest a possible effect of length on peptidyl DPP potency
(Case and Stein, 2003). A mono-, di-, and tripeptide phospho-
nate was synthesized for the QRVBz-DPP probe (1), as this
sequence proved to inhibit both proteases well (Table 2). A clear
correlation was observed between increasing peptide length and
improvement in inhibition, with a 10-fold increase between the
IC
50
of the longest inhibitor (QRVBz-DPP, 1) and the shortest
inhibitor (Bz-DPP, 5), from 0.37 mM to 3.5 mM. A similar trend
was seen in the k
inact
/K
I
data across the same parameters.
Table 2. Inhibition Kinetics of Phosphonate Probes against MT-
SP1 and Thrombin
Enzyme Compound Inhibitor IC
50
(mM)
A MTSP1 5 Bz 3.5 ± 0.6
6 VBz 8 ± 0.1
7 RVBz 0.56 ± 0.09
1 QRVBz 0.37 ± 0.2
3 GSGBz 14 ± 1
2 LTPBz 1.1 ± 0.02
4 EPIBz 76 ± 1
13 FTGSGBz 3.5 ± 0.5
B MTSP1 F99A 5 Bz 1.5 ± 0.04
6 VBz 1.3 ± 0.06
7 RVBz 0.23 ± 0.01
1 QRVBz 0.22 ± 0.01
3 GSGBz 9.2 ± 0.4
C Thrombin 5 Bz 0.097 ± 0.01
6 VBz 0.28 ± 0.05
7 RVBz 0.068 ± 0.04
1 QRVBz 0.13 ± .002
3 GSGBz 0.027 ± 0.01
2 LTPBz 1.8 ± 0.5
4 EPIBz 0.76 ± 0.2
D MTSP1 Compound Inhibitor k
inact
/K
I
(M
–1
s
–1
)
5Bz50±2
6 VBz 60 ± 10
7 RVBz 490 ± 40
1 QRVBz 200 ± 60
3 GSGBz 26 ± 0.5
4 EPIBz 11.7 ± 3
MT-SP1 and thrombin were incubated with varying concentrations of
inhibitor and activity was monitored on addition of substrate. See also
Table S2 and Figure S1.
Chemistry & Biology
Potency Determinants of Peptide Phosphonates
50 Chemistry & Biology 18, 48–57, January 28, 2011 ª2011 Elsevier Ltd All rights reserved
It was clear that not all positions affected MT-SP1 inhibition
equally. The addition of a P3 residue increased potency by an
order of magnitude (from 8 mM to 0.56 mM), whereas only modest
changes were observed by the addition of P2 or P4. This was
consistent with previously observed substrate:enzyme binding
interactions with serine proteases, in which much of the binding
energy is contributed by the P1 amino acid interacting with the
S1 pocket of the protease. This also correlated with known
MT-SP1 substrate data and with crystallographic observations
about the S4 and S3 pockets (Bhatt et al., 2007; Friedrich
et al., 2002). Friedrich et al. (2002) observed that the deep S3
pocket of MT-SP1 could accommodate a Lys or Arg residue
from either P3 or P4 of a substrate whereas the other amino
acid is solvent exposed. This phenomenon could explain the
modest increase in inhibition seen when increasing from a tripep-
tide- to a tetrapeptide phosphonate.
A similar trend was seen in IC-50 values measured against
thrombin (Table 2, section C) and uPA (data not shown). Again,
it is notable that QRVBz-DPP (1), the ideal MT-SP1 inhibitor,
was more potent than the ideal thrombin inhibitor LTPBz-DPP
(2) by an order of magnitude (0.125 uM for 1 compared to 1.1
uM for 2). Sequence data alone does not seem to be sufficient
A
B
C
D
Figure 2. Structure of the Catalytic Domain
of MT-SP1 Bound to Benzamidine Phospho-
nate
(A) Numbered amino acid residues correspond to
chymotrypsin protease numbering.
(B) Close-up ribbon of thrombin (tan) overlaid onto
MT-SP1 (light blue). Thrombin L99 (purple), MT-
SP1 F99 (blue), catalytic triad (cyan).
(C) Thrombin (tan) surface overlaid onto MT-SP1
(light blue). Thrombin L99 (purple), MT-SP1 F99
(blue), cataly tic triad (cyan).
(D) Thrombin surface (tan) overlaid onto MT-SP1
(light blue) and uPA (pink). uPA His99 (yellow)
occupies prime-side pocket similarly as MT-SP1
Phe99 (blue). Thrombin Leu99 (purple), catalytic
triad (cyan). See also Figure S2.
to design ABPs that can distinguish
between these two proteases, though
other proteases with much more stringent
substrate binding pockets have shown to
be amenable to sequence-derived speci-
ficity (Mahrus and Craik, 2005).
To determine if potency continued to
improve beyond P4, a hexapeptide ABP
was synthesized and tested against
MT-SP1. The preferred P5 and P6 amino
acids for MT-SP1 were added to the
most stable inhibitor (GSGBz, 3) to create
13 (FTGSGBz). 13 is 25% more potent
than 3 against MT-SP1, verifying that
further increasing length improved
potency.
Interestingly, each of the ABPs synthe-
sized for this study inhibited thrombin
at much lower concentrations than
MT-SP1. We next sought to obtain
a structural basis for the improvement in inhibition observed
against thrombin.
Structure of MT-SP1 Bound to Bz-DPP
The structure of MT-SP1 bound to Bz-DPP phosphonate was
solved to 1.19 A
˚
resolution (Figure 2 and Table 3). The protein
was incubated with saturating ABP concentrations overnight
at room temperature and purified via size exclusion chromatog-
raphy. Attempts to crystallize MT-SP1 with Ac-QRVBz-DPP only
produced crystals containing Bz-DPP. A possible explanation is
that nonspecific hydrolysis of the peptide bond occurred during
the several rounds of seeding and crystal growth that were
necessary to obtain high-quality crystals, all of which took place
in an aqueous environment at room temperature over several
months. The truncated inhibitor may have also resulted from
incomplete purification of the Bz-DPP (12) starting material
from the full-length ABP.
This is the highest resolution MT-SP1 structure to date, which
allows for visualization of all bonds. The phosphonate bound in
the solvent exposed binding pocket in the expected conforma-
tion, with the phosphorous atom bound to the active site
Ser195 and Bz in the S1 pocket (Figure 2A). The positively
Chemistry & Biology
Potency Determinants of Peptide Phosphonates
Chemistry & Biology 18, 48–57, January 28, 2011 ª2011 Elsevier Ltd All rights reserved 51
charged guanidine group of Bz formed two hydrogen bonds with
the negatively charged Asp189. The N-terminal end of the inhib-
itor pointed down the substrate binding pocket, and the phenyl
ring is solvent exposed.
A structure of thrombin (PDB: 1QUR) (Steinmetzer et al., 1999)
was overlaid onto the MT-SP1-Bz-DPP structure, and the probe
fit similarly into the binding pocket (Figure 2B). Inspection of the
placement of the phenyl rings of the phosphonate revealed one
important difference in the active site architecture. In MT-SP1
residue Phe99 lay in the region where the leaving group phenyl
ring binds during catalysis. In thrombin, a smaller Leu99
occupied this location, forming a larger pocket than seen in
MT-SP1. The Phe99 was only 2 A
˚
away from the phosphorous,
thus we hypothesized that Phe99 caused steric interference
with the leaving phenyl ring, slowing orientation of the inhibitor
in the active site and slowing inhibition of MT-SP1 relative to
that of thrombin.
To test this hypothesis, MT-SP1 Phe99 was mutated to Ala to
create MT-SP1 F99A, and IC
50
measurements with the same
series of inhibitors were repeated (Table 2, section B). The IC
50
values for MT-SP1 F99A fell between that of MT-SP1 and
thrombin in each case, indicating the size of the binding pocket
for the leaving group in MT-SP1 influenced phosphonate
inhibition.
When the structure of uPA (PDB: 3KGP) (Zhang et al., 2010), an
S1A protease with slow kinetics of phosphonate inhibition similar
to those of MT-SP1 was overlaid with thrombin, a similar steric
hindrance was observed (Figure 2D). At the same pocket in
uPA, His99 protruded exactly where Phe99 did in MT-SP1.
Thrombin had a larger binding pocket than either MT-SP1 or
uPA, due to the Leu found at residue 99. The smaller pocket in
uPA, as with MT-SP1, may explain the slow inhibition of uPA
by DPPs. This reinforces the idea that the steric fit of the leaving
group on the prime side has an important effect on potency.
Labeling of Cell Surface Proteases by ABPs
Having determined the kinetic and labeling properties of the
ABPs in vitro, their potential for labeling active proteases on the
surface of live immortalized cancer cells was examined. The
most stable inhibitor (13) and the most potent inhibitor (1) were
used to label the cell surface proteases of two different cell lines
grown in culture. These cell lines represented epithelial cancer
types from two different species: PC3 cells, derived
from human prostate cancer, and PDAC2.1 cells, derived from
a transgenic mouse model of pancreatic ductal adenocarcinoma
(Nolan-Stevaux et al., 2009). Live cells were incubated with phos-
phonate overnight in serum free media at 37
"
C. Streptavidin
conjugated with AlexaFluor488 dye was used to fluorescently
tag the ABPs. Cells were examined for the presence of labeled
proteases either by fluorescent microscopy or by flow cytometry.
As seen in Figure 3, green fluorescent labeling was observed in
PC3 and PDAC cells that had been exposed to ABPs, indicating
the presence of active S1A proteases on the cell surface.
PDAC2.1 (Figure 3A) cells labeled discrete foci on the cell
surface. In the case of PC3 cells (Figure 3B), these labeled prote-
ases were localized to one end of each cell. In addition to cell
surface labeling, punctate labeling was also observed just
internal to the plasma membrane. These puncta colocalized
with membrane staining dye, indicative of internalization of
labeled surface proteases. Internalization was also occasionally
observed in PDAC2.1 cells and more frequently observed in
MCF7 cells, a human breast cancer cell line (Figure S2) This
result indicates that ABPs can visualize and follow active serine
proteases on the surface of live cells, a novel use for peptide
phosphonates.
After demonstrating the ability to label cell surface proteases,
the ability to quantify the active form of these proteases was
tested. Cells were grown, exposed to phosphonate overnight
at 37
"
C, and fluorescently tagged with labeled streptavidin. Fluo-
rescence was quantified using flow cytometry on the live cells.
Fluorescence increased relative to background when cells had
been incubated with an ABP, and this labeling was reduced in
the presence of protease inhibitors (Figure 3E). These results
demonstrate that ABPs can be used to quantify active proteases
on the surface of live cells, and that labeling can be directly
attributed to proteolysis.
Collectively, the cell-based experiments show that peptide
phosphonates can be used to quantitatively label active cell
surface proteases on different cell types. Live cell imaging can
Table 3. Data Collection and Refinement Statistics
Structure MSTP1-covalent adduct
PDB ID 3NCL
Data set ALS290709
Data statistics
Wavelength 1.11587 A
˚
Space group C2
Cell dimensions a = 79.9 A
˚
b = 80.2 A
˚
c = 40.5 A
˚
b = 95.8
"
Resolution (last shell) 54.6–1.19 A
˚
(1.25–1.19 A
˚
)
Unique reflections 58,355 (1597)
Redundancy 6.7 (3.8)
Completeness 76.4% (14.5%)
Mean I/s (I) 21.0 (5.0)
R
sym
6.5% (15.0%)
Refinement statistics
Resolution range 54.6–1.19 A
˚
Reflections used word (test) 56,653 (1530)
R
free
/R
fac
20.3%/19.4%
Overall figure of merit 0.915
Overall B
wilson
12.7A
˚
2
Protein atoms 1900, 21.8 A
˚
2
Ligand atoms 21, 34.8 A
˚
2
Solvent atoms 221, 38.9 A
˚
2
Rmsd bonds 0.016 A
˚
Rmsd angles 1.010
"
Ramachandran analysis
Residues in preferred regions 96.4%
Residues in allowed regions 3.6%
Outliers 0%
Rmsd = root-mean-square deviation.
Chemistry & Biology
Potency Determinants of Peptide Phosphonates
52 Chemistry & Biology 18, 48–57, January 28, 2011 ª2011 Elsevier Ltd All rights reserved
also be used in conjunction with ABP labeling to determine the
localization of active proteases in the context of a cell or group
of cells.
DISCUSSION
This study presents additional guidelines for the synthesis,
design, and application of peptide phosphonate ABPs to those
described previously (Oleksyszyn and Powers, 1994). Our
results show that by focusing on peptide length, stability, and
leaving group sterics, the lifetime and utility of these ABPs can
be improved dramatically. By following these guidelines, a novel
protease imaging application using ABPs was developed. These
experiments demonstrate that ABPs can be used to quantify,
image, and follow proteases on cell surfaces.
We have improved the utility of phosphonate ABPs by
combining synthetic, kinetic, and structural data. Obtaining
reagent quantities of material has been a significant barrier to
using these molecules in numerous applications. The improved
synthesis methodology presented here both increases reproduc-
ibility of ABP production and increases yields up to seven fold
compared to previous methods. The robust generation of ABPs
greatly improves their utility, and synthesizing greater quantities
of these probes enabled a systematic study of their potency.
Three major factors were found to contribute to peptide phos-
phonate potency: peptide stability, length, and leaving group
sterics. Peptide sequence is often viewed as a primary determi-
nant of potency, and reports of sequence-based selectivity exist
for Granzymes A and B, two S1A proteases with highly selective
A
C
B
D
E
Figure 3. Cell Surface Labeling of Active Serine
Proteases
PDAC2.1 and PC3 cells were imaged in the presence
(A and C) or absence (B and D) of ABP. Cell membranes
were stained with tetramethylrhodamine-conjugated
wheat germ agglutinin (red), and labeled proteases were
stained with streptavidin-AlexaFluor 488 (green). Colocal-
ization of labeled proteases with membrane is seen at the
cell surface and in discrete puncta, but only in the pres-
ence of ABP. Flow cytometry was used to quantify labeled
streptavidin on the surface of cells (E). Averaged mean
fluorescence intensity values are plotted normalized to
background staining of labeled streptavidin in the absence
of ABP. P values correspond to a two-sample t test, and
error bars correspond to the addition of the relative stan-
dard error values for the two averages multiplied by the
normalized mean of each sample.
and differing substrate specificities at P2-P4
(Mahrus and Craik, 2005). However, for the
majority of trypsin-fold proteases, much of the
substrate binding energy of is contributed via
binding of a basic P1 amino acid in the deep
S1 pocket of the active site. Because of the
strong P1 contribution, the P2-P4 sequence
often plays a minor role. Peptide stability,
however, was found to contribute significantly
to ABP functionality. Inhibitors containing the
amino acid Pro were especially unstable and
problematic for both synthesis and purification,
whereas long, charged side chains like Arg were found to react
with the electrophilic phosphonate, subsequently inactivating
the inhibitor. Additionally, the use of basic residues like Arg
upstream of P1 can result in a secondary cleavage site for
trypsin-like proteases, which removes the probe from the reac-
tive diphenylphosphonate moiety. Poorly fitting residues can
have a detrimental effect on inhibition, as seen by the slow
kinetics of the EPIBz (4) probe against MT-SP1, thrombin, and
uPA (Table 2; data not shown). Therefore, the optimal peptide
element should be composed of the most stable residues toler-
ated by the protease.
Peptide length, rather than sequence, had a more noticeable
contribution to improving inhibitory capabilities. Increasing
length increased potency, most notably at the P3 position. This
agrees with earlier findings (Oleksyszyn and Powers, 1991).
The data suggests that when designing peptide phosphonate
inhibitors, including a longer peptide will improve inhibition
more reliably than modifying the sequence. The sequence
composition data and stability observations indicate that phos-
phonate inhibitors of trypsin-fold proteases should contain at
least a tetrapeptide composed solely of stable amino acids.
The third factor contributing to potency is the steric fit of the
phosphonate leaving group in the binding pocket of the
protease. Atomic-resolution structural data showed this pocket
is smaller in MT-SP1 than in thrombin. This pocket is adjacent
to the catalytic S195, exactly where the leaving group phenyl
ring would reside. This smaller pocket in MT-SP1 is due to the
presence of a bulky Phe at residue 99. Mutagenesis experiments
confirmed that this pocket is an important binding determinant
Chemistry & Biology
Potency Determinants of Peptide Phosphonates
Chemistry & Biology 18, 48–57, January 28, 2011 ª2011 Elsevier Ltd All rights reserved 53
for DPPs and MT-SP1. The data indicates that the leaving group
of the phosphonate can have a large effect on inhibition. When
designing a phosphonate ABP specific for a protease, multiple
leaving groups should be tested on a stable peptide scaffold to
find the best inhibitor.
The data shown here provide important information for the
design of peptide phosphonate ABPs with both broad and
narrow specificity. We show that the peptide element (in stability
and length) and the leaving group (in reactivity and sterics)
contribute to inhibitory potency of phosphonate ABPs. When
designing a broad-specificity phosphonate ABP, one should
start with a scaffold containing at least four stable amino acids
and then consider varying the leaving group if improved potency
is desired. When designing an inhibitor for a specific S1A
protease, substrate cleavage data should be viewed as a starting
point. Varying the leaving group may result in larger differences in
potency, even for closely related proteases. Interestingly, kinetic
data indicates that varying the reaction time may label different
sets of proteases, with longer incubations resulting in larger
sets of labeled proteases. Therefore, reaction time may also
influence selectivity. However, these experiments suggest that
only in rare instances will true specificity be engineered by
varying peptide composition alone.
By following the guidelines presented here, several broad-
spectrum ABPs for S1A proteases were produced in high yields.
We used the most potent (1) and stable (13) ABPs to develop
a novel method of imaging proteases on the surface of cancer
cells. ABP-labeled proteases can be visualized and quantified
with fluorescently conjugated streptavidin. This approach found
that levels of protease activity vary by cell type, and that the
location of labeled proteases can be tracked through the cell
(Figure 3). Protease activity and localization information can be
leveraged to obtain new information about protease function.
The ability to study the localization of active proteases is
a novel use of phosphonate activity-based probes, and offers
a promising way to examine the biology of cell surface protease
activity. Previous studies with cysteine protease ABPs have
been successful at labeling and visualizing lysosomal cathepsins
and extracellular cathepsins in tumors (Blum et al., 2005; Joyce
et al., 2004). The current study extends the use of ABPs to serine
proteases on the surface of live cells. Differences in labeling and
localization varied between the cell types, a conclusion
previously observed with MT-SP1 using specific antibody-based
probes (Darragh et al., 2010). These broad-spectrum ABPs may
be used to highlight cell-specific differences in global S1A
protease function.
For example, cell surface proteases have been implicated in
metastasis, and it is interesting to note that active protease
localization was polarized. Notably, high concentrations of
enzymes were observed at one edge of the cells distal to cell-
cell junctions. In addition, correlating the degree of labeling
with a cellular phenotype such as metastatic potential may yield
new information about cancer cell biology.
Although the pattern varied, internalization of proteases was
observed in all cell types, as evidenced by the colocalization of
membrane and ABP in puncta just inside the cell membrane.
We hypothesize two explanations for this observation. First,
many proteins undergo natural trafficking to and from the
cell surface, modulating cellular interactions with the outside
environment. Trafficking has been implicated previously in the
regulation of proteases in the cell (Ghosh et al., 2003). ABPs could
thus be used to track the movement and function of active prote-
ases in many complex biological systems, including cancer
biology, where protease activity is frequently dysregulated. Alter-
natively, the ABPs themselves may cause proteases to become
internalized. Ligand-dependent endocytosis has been observed
with other cell surface receptors (Behrendt, 2004; Ghosh et al.,
2003). The molecular mechanism of cell surface protease
internalization observed via ABPs invites further investigation.
In summary, through synthetic, kinetic, and structural insights,
we have developed additional guidelines that define and expand
the use of peptide phosphonate ABPs. By following these
guidelines, we have created a potent pan-S1A protease probe
and developed a general methodology for the study of active
proteases at the surface of cells. This technology could allow
for future insights into the role of proteases in cancer, and has
potential applications to other fields in biology.
EXPERIMENTAL PROCEDURES
Materials
MT-SP1 was expressed and purified as described previously and stored at
#20
"
C in 50 mM Tris pH 8.0, 50 mM NaCl, 10% glycerol (Takeuchi et al.,
1999). Mutants for crystallography and kinetic studies were expressed, puri-
fied, and stored in the same manner. Thrombin was purchased from Sigma
and stored at #20
"
C in 50 mM Tris pH 8.0, 50 mM NaCl, 0.1 mg/ml BSA.
The MT-SP1 substrates spectrazyme-tPA and spectrafluor-tPA were
purchased from American Diagnostics and st ored at #20 at 10 mM in H
2
O.
The materials for peptide synthesis including PyBOP, EDAC, HOBt, Fmoc-
PEG
20atom
-OH, and 2-chlorotrityl chloride resin were purchased from
NovaBiochem. All other chemicals were purchased from Sigma unless other-
wise noted. Solvents including anhydrous ethanol (EtOH), chloroform (CHCl
3
),
1,4-dioxane, diethyl ether (Et
2
O), dichloromethane (DCM), triflouroethanol
(TFE), trifluoroacetic acid (TFA), and DMF were used as received. The peptides
were synthesized following standard Fmoc-SPPS procedure on 2-chlorotrityl
chloride resin.
Reactions were analyzed by LC-MS performed on a Waters Alliance liquid
chromatography system with a Waters Micromass ZQ single-quadrupole
mass spectrometer. HPLC purifications were carried out using an Agilent
1200 series system with C18 reversed-phase columns (Waters). Mobile phase
consisted of 99.9%:0.1% water/trifluoroacetic acid (solvent A) and
95%:4.9%:0.1% acetonitrile/water/trifluoroacetic acid (solvent B). All final
compounds were characterized by Matrix-assisted laser desorption ionization
time-of-flight (MALDI-TOF) mass spectrometry using an ABI 4700 MALDI-
TOF-TOF mass spectrometer.
Compound numbers in bold refer to the structures shown in Figure 1.
Diphenyl [N-(benzyloxycarbonyl)a mino](4-cyanophenyl)methanephosphonate
(9) and NHS-biotin were synthesize d according to literature procedure. No
attempts were made to resolve the D,L-(4-AmPhGly)
P
(OPh)
2
diastereomers.
Diphenyl-[N-(benzyloxycarbonyl)amino](4-ethylesterphenyl)
methane phosphonate Hydrochloride (10)
A batch of 9 (3.0g, 6.0mmol) was dissolved in 100 ml CHCl
3
and placed in a 0
"
C
bath. Next Ar was passed over the solution and under the flow of Ar, 3 ml of
anhydrous EtOH (60 mmol) and 100 ml of 4 M HCl in dioxane were added.
The reaction was stirred under argon at 4
"
C for 5 days, after which 10 formed
as a fine white precipitate formed. The precipitate was filtered, washed with
Et
2
O, and dried under reduced pressure (2.32 g, 72%). Mass calculated for
C
30
H
29
N
2
O
6
P: 544.18; found 545.45 (M + H)
+
.
Diphenyl Amino(4-amidinophenyl)methanephosphonate,
Trifluoroacetic Acid Salt (12)
A batch of 10 (1.0g, 1.8 mmol) was dissolved in 25 ml 1,4-dioxane and 25 ml
anhydrous EtOH. Next the reaction was purged with Ar and 10 ml of 0.5M
Chemistry & Biology
Potency Determinants of Peptide Phosphonates
54 Chemistry & Biology 18, 48–57, January 28, 2011 ª2011 Elsevier Ltd All rights reserved
NH3 in dioxane (5 mmol) was added dropwise. The reaction was stirred
under argon at 23
"
C for 2 days, and the solvent was removed completely
under vacuum to afford the gummy white crude intermediate, diphenyl
[N-(benzyloxycarbonyl)amino](4-amidinophenyl) methanephosphonate 11.
This intermediate was dissolved in anhydrous EtOH (80 ml) and concentrated
HCl (305 mL, 3.7 mmol) and hydrogenated over 10% palladium on activated
carbon for 5 hr at room temperature. The catalyst was separated by filtration,
and the solvent was removed under reduced pressure. The white powder thus
isolated was dissolved in HPLC solvent (water with 0.1% TFA) with the aid of
DMF and purified by reverse-phase HPLC. Lyophilization of fractions contain-
ing product afforded 0.49 g (27%) of 12 as a white powder. Mass calculated for
C
20
H
20
N
3
O
3
P: 381.12, m/z found: 382.01 (M + H)
+
.
Coupling of the Biotinylated Peptide to 12: General Procedure
To a DCM/TFE (5:2) solution containing the biotin-PEG-peptide (0.03 mmol),
EDAC (0.03 mmol) and HOBt (0.03 mmol), was added H-Bz-DPP 12
(0.03 mmol). After 2 hr, an additional equivalent of EDAC and HOBt was added
and the mixture stirred for 8 hr. Next the solvent was removed, the isolated oil
was dissolved in MeCN/water (1:3), and purified by reverse-phase HPLC. The
fractions with product were lyophilized down to yield the desired product as
a white powder. The final probes were dissolved in DMSO and stored at
#20
"
C. The concentration of the solution was determined by HABA biotin
quantification kit (Pierce).
Biotin-PEG-Gln-Arg-Val-Bz-DPP 3 TFA (1)
Reaction of Biotin-PEG-Gln(Trt)-Arg(Pbf)-Val-OH (0.10 g, 0.11 mmol), with 12
yielded 39 mg (72% yield) of Biotin-PEG-Gln(Trt)-Arg(Pbf)-Val-DPP as a white
powder. Mass calculated for C
92
H
119
N
14
O
18
PS
2
: 1802.80, m/z found: 1802.10
(M + H)
+
. Next the powder was resuspended in 95% TFA, 2.5% TIS, 2.5%
water and agitated for 2 hr, to obtain the desired probe 1. The product was
precipitated into cold ether, pelleted by centrifugation, and purified by
reversed-phase HPLC. Lyophilization of fractions containing product af-
forded 22 mg (79%) afforded 1 as a white powder. Mass calculated for
C
60
H
89
N
14
O
15
PS: 1308.61, m/z found: 656.34 (M + 2H)
2+
.
Biotin-PEG-Leu-Thr-Pro-Bz-DPP (2)
The protected peptide Biotin-PEG-Leu-Thr(tBu)-Pro-OH was reacted with 12
and resulted in Biotin-PEG-Leu-Thr(tBu)-Pro-Bz-DPP as a white powder
(22 mg, 56% yield). Mass calculated for C
63
H
93
N
10
O
15
PS: 1292.63, m/z found:
1293.29 (M + H)
+
. Next the powder was resuspended in 95% TFA, 2.5% TIS,
2.5% water and agitated for 2 hr, to obtain the desired probe 2. The product
was precipitated into cold ether, pelleted by centrifugation, and purified by
reversed-phase HPLC. Lyophilization of fractions containing product afforded
8 mg (36% yield). Mass calculated for C
59
H
85
N
10
O
15
PS: 1236.57, m/z found:
1237.12 (M +H)
+
.
Biotin-PEG-Gly-Ser-Gly-Bz-DPP (3)
Reaction of Biotin-PEG-Gly-Ser-Gly-OH with 12 resulted in 3 as a white
powder (22 mg, 65%). Mas s calculated for C
51
H
71
N
10
O
15
PS: 1126.46, m/z
found: 1126.92(M + H)
+
.
Biotin-PEG-Glu-Pro-Ile-Bz-DPP (4)
Biotin-PEG-Glu(OtBu)-Pro-Ile-OH was reacted with 12 to afford 23 mg (57%
yield) of Biotin-PEG-Glu(OtBu)-Pro-Ile-Bz-DPP. Mass calculated for
C
64
H
93
N
10
O
16
PS: 1320.62, m/z found: 1321.43 (M + H)+. Then the powder
was resuspended in 95% TFA, 2.5% TIS, 2.5% water, and agitated for 2 hr
to obtain the desired probe. The product was precipitated into cold ether,
pelleted by centrifugation, and pu rified by reversed-phase HPLC. Lyophiliza-
tion of fractions containing product afforded 20 mg (90% yield) of 4. Mass
calculated for C
60
H
85
N
10
O
16
PS: 1264.56, m/z found: 1266.31 (M + H)
+
.
Biotin-PEG-Bz-DPP (5)
Reaction of Biotin-PEG-OH with 12 yielded 5, which was isolated as white
powder (7 mg, 25%). Mass calculated for C
44
H
60
N
7
O
11
PS: 925.38, m/z found:
926.17 (M + H)
+
.
Biotin-PEG-Val-Bz-DPP (6)
A batch of Biotin-PEG-Val-OH was reacted with 12 to afford 6 as a powder
after HPLC purification (7 mg, 20%). Mass calculated for C
49
H
69
N
8
O
13
PS:
1024.45, m/z found: 1025.82 (M + H)
+
.
Biotin-PEG-Arg-Val-Bz-DPP (7)
Reaction of Biotin-PEG-Arg(Pbf)-Val-OH with 12 afforded Biotin-PEG-Arg
(Pbf)-Val-OH as a white powder (34 mg, 77%). Mass calculated for
C
68
H
99
N
12
O
17
PS
2
,: 1450.64, m/z found: 1451.24 (M + H)
+
. Then the powder
was dissolved in 95% TFA, 2.5% TIS, 2.5% water, and agitated for 2 hr to
obtain the desired probe. The product was precipitated into cold ether,
pelleted by centrifugation, and purified by reversed-phase HPLC. Lyophiliza-
tion of fractions containing product afforded 24 mg (90% yield) of 7. Mass
calculated for C
55
H
81
N
12
O
14
PS: 1180.55, m/z found: 1181.64 (M + H)+.
Biotin-PEG-Phe-Thr-Gly-Ser-Gly-Bz-DPP (13)
Reaction of Biotin-PEG-Phe-Thr-Gly-Ser-Gly-OH with 12 afforded 13 as
a white powder (45 mg, 40%). Mass calculated for C
64
H
87
N
12
O
18
PS:
1374.57, m/z found: 1376.27 (M + H)
+
.
Inhibition Assays
All kinetic fluorescence measurements were taken in duplicate using a Spec-
traMax Gemini fluorescence spectrometer (Molecular Devices) with an excita-
tion wavelength of 380 nm, an emission wavelength of 460 nm, and a 435 nm
cutoff filter. A solution of inhibitor was serially diluted over an appropriate
concentration range and incubated with enzyme. Substrate was added at
the end of 4 hr to initiate the reaction, and IC
50
s were calculated. MT-SP1
was used at 0.2 nM in a buffer containing 50 mM Tris pH 8.0, 50 mM NaCl,
0.01% Tween-20, with 200 mM spectrafluor-tPA as the substrate. Thrombin
was used at 0.5 nM in a buffer containing 50 mM Tris pH 8.0, 50 mM NaCl,
0.01% Tween-20, 0.1 mg/ml BSA, with 200 mM Boc-b-benzyl-Asp-Pro-Arg-
AMC as the substrate. All reactions were run in duplicate.
Steady-state kinetics were used to determine the observed rate constants
for the inhibition reaction. The inhibitors were serially diluted in a 96-well plate
at an appropriate range of concentrations. Enzyme was added in hour inte rvals
over 8–10 hr. The reaction was initiated by the addition of 200 mM substrate,
and the V
max
recorded on a SpectraMax fluorescence spectrometer. K
obs
was determined at each inhibitor concentration by plotting V
max
versus time,
and K
Iapp
was determined by plotting K
obs
versus [I]. K
inact
/K
I
was determined
using the equation
k =
k
2
½I
o
%
½I
o
% + K
i
&
!
1 +
½S
o
%
K
m
"
where K
i
&
=
k
inact
K
I
:
Crystallization
Crystals were grown at room temperature by vapor diffusion in hanging drops.
A combination of micro and macro seeding was used to grow large single
crystals in 4.0 M Na Formate at pH = 7.0 and 25 mM FeCl
3
as additive. Crystals
belong to monoclinic space group C2 with one protease-inhibitor complex in
the asymmetric unit corresponding to a solvent content of 50% and diffracted
to >1.2 A
˚
resolution.
Structure Resolution and Refinement
Data were collected at beamline 8.3.1 at the Advanced Light Source in Berke-
ley on a single crystal cryoprotected in mother liquor supplemented with 20%
glycerol. The data were indexed, scaled, and reduced using Mosflm and Scala
in Elves (Holton and Alber, 2004). The structure was solved by molecular
replacement using Phaser (McCoy et al., 2007) with the previously solved
protease structure (PDB: 3BN9) as search probe (Farady et al., 2008). Auto-
matic building and refinement were performed in Phenix (Adams et al., 2002)
using Phenix elBow to generate the covalently bound ligand. Manual building
was carried out in Coot (Emsley and Cowtan, 2004). The stereochemistry of the
final model was validated using MolProbity (Davis et al., 2007).
Western Blot Labeling Analysis
For recombinant protease labeling, enzyme was combined with varying
concentrations of phosphonate inhibitor (3 mM–30 mM) at room temperature
Chemistry & Biology
Potency Determinants of Peptide Phosphonates
Chemistry & Biology 18, 48–57, January 28, 2011 ª2011 Elsevier Ltd All rights reserved 55
overnight. The reaction was stopped by the addition of SDS loading buffer and
boiling for 10 min. Western blots were developed using the Vectastain ABC
elite kit (Vector Labs).
Fluorescent Substrate Synthesis
Substrates corre sponding to each inhibitor were synthesized by solid phase
peptide synthesis. ACC-Rink-amide resin was obtained from Kimia Corp.
The first amino acid was coupled using five equivalents each of amino acid,
HATU, and collidine in dry DMF under argon for 16 hr with agitation. The full-
length peptide was synthesized using a Symphony Quartet peptide synthe-
sizer (Protein Technologies), acetylated with eight equivalents each of acetic
anhydride and DIPEA, and cleaved with 95% TFA/2.5% water/ 2.5% triiso-
propyl silane. Cleaved peptides were precipitated into cold ether, collected
by centrifugation, and purified by HPLC.
Cell Culture and Propagation
PC3 cells were obtained from the American Type Culture Collection (ATCC)
and propagated in F12K Nutrient mixture with Kaighn’s Modification (13)
and L-Glutamine (GIBCO). Media was supplemented with 10% FBS and 13
penicillin/streptomycin. MCF7 cells were obtained from the ATCC and
propagated in Dulbecco’s modified Eagle’s medium with high-glucose
(D-ME H21) without phenol red (GIBCO). Media was supplemented with
10% FBS, 10 ug/ml Insulin, and 13 penicillin/streptomycin. PDAC2.1 cells
were isolated from p48-Cre/+, LSL-KrasG12D/+, Trp53F/+ transgenic mice
according to (Nolan-Stevaux et al., 2009). PDAC2.1 cells were propagated in
D-ME H21 (GIBCO) supplemented with 10% FBS and 13 penicillin/
streptomycin.
Flow Cytometry
Cells were grown to confluence in 6-well cell culture-treated dishes using
complete media appropriate for each specific cell line. Media was then
aspirated and replaced with Opti-MEM serum-free medium (GIBCO). All
experiments were done in triplicate. FTGSGBz-DPP (13) was added to the
appropriate wells at a final concentration of 50 mM. To test proteolysis, ABP
was added in the presence of 13 Complete Protease Inhibitor Cocktail (Roche)
dissolved in Opti-MEM. The cells were then incubated at 37
"
C for !20 hr. After
incubation, ABP-containing media was aspirated and cells were washed three
times with Opti-MEM. Cells were detached from the surface of the wells with
Enzyme-free Cell Dissociation Buffer (GIBCO) for 15 min. Detached cells were
washed two times with Opti-MEM, resuspended in 200 ml, and incubated for
20 min with Streptavidin/AlexaFluor 488 conjugate (Invitrogen) at 4
"
C. Cells
were washed three times with Opti-MEM and resuspended in 500 ml Opti-MEM
and assayed for fluorescence using a BD FACSCalibur (BD Biosciences). Data
was analyzed and population mean fluorescence values were obtained using
FlowJo Flow Cytometry Analysis Software (TreeStar, Inc.).
Microscopy
Cells were grown to confluence in 9.5 cm
2
glass bottom microwell dishes
(MatTek) in complete media. Media was then aspirated and replaced with
Opti-MEM. Phosphonate was added to each dish to a final concentration of
50 mM. FTGSGBz-DPP (13) was incubated for 20 hr at 37
"
C and QRVBz-
DPP (1) was incubated for 3 hr at 37
"
C, respectively. Cells were washed three
times with Opti-MEM and then incubated for 20 min with Streptavidin/Alexa-
Fluor 488 conjugate and tetramethylrhodamine-conjugated wheat germ
agglutinin (Invitrogen), simultaneously, at 4
"
C. Cells were then washed three
times with Opti-MEM and imaged. Fluorescence microscopy was carried
out in the wide field using a Nikon Diaphot with a Nikon 603 lens, numerical
aperture 1.4, objective and standard interference filter sets (Omega Optical).
Images were collected using a 12-bit cooled charge-coupled device camera
(Princeton Instruments) interfaced to a computer running Micro-Manager 1.3
software (http://micro-manager.org). Images were processed using Adobe
Photoshop to assemble dual color image files. Brightness/contrast was
adjusted, where necessary, to improve image quality and clarity.
SUPPLEMENTAL INFORMATION
Supplemental Information includes two figures and two tables and can be
found with this article online at doi:10.1016/j.chembiol.2010.11.007.
ACKNOWLEDGMENTS
Mass spectrometry was provided by the Bio-Organic Mass Spectrometry
Resource at UCSF (A. L. Burlingame, Director) supported by the Bio medical
Research Technology Program of the NIH Natio nal Center for Research
Resources (NIH NCRR P41RR001614 and NRCC RR014606). This work was
supported by a NIH/NIGMS award (K12GM081266) to A.E.-R., a NIGMS-
IMSD award (R25-GM56847) to C.T., and by the National Institutes of Health
(NIH R01CA128765).
Received: June 25, 2010
Revised: October 26, 2010
Accepted: November 5, 2010
Published: January 27, 2011
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    • "The beads were pelleted by centrifugation for 30 sec at 3000 rcf, and the soluble portion was moved to a new microfuge tube. The probe biotin-nVPL-O(Ph) 2 (biotin-PEG-norleucyl-valylprolyl-leucyl-diphenyl phosphonate) was synthesized following a previously described method, added for a final concentration of 5 mM, and incubated for 1 hour at room temperature [15]. To stop the reaction, 400 mL 1 M guanidine-HCl was added and the entire reaction was run over a 5 kDa Amicon filter (Millipore, Bedford, MA). "
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  • [Show abstract] [Hide abstract] ABSTRACT: α-Aminoalkylphosphonate diaryl esters are potent, irreversible, and highly selective site-directed inhibitors of serine proteases. The structure of the phosphonate group resembles the transition state observed during a peptide bond hydrolysis and therefore phosphonates are referred as transition state analogues. They react with the hydroxyl group of the active site serine residue leading to formation of a stable enzyme-inhibitor complex. Moreover, incorporation of a peptidyl chain at the N-terminus as well as an introduction of electron withdrawing or electron donating substituents within the ester ring structure allows for a generation of specific inhibitors that react only with target serine protease. The great advantage of the aminophosphonate diaryl esters over other classes of inhibitors is their stability in aqueous solutions, no toxicity and lack of reactivity with cysteine, threonine, aspartyl and metalloproteinases. The above resulted in their application as convenient tools to study proteases function and activity using in vivo and in vitro assays of different pathological disorders (diabetes, cancer metastasis, pulmonary diseases or hypertension); to determine the cellular localization of the proteinases (activity based probes), to be used in proteomic approach or as the reactive antigens to develop a catalytic function within the antibodies binding site. Herein we present the development of α-aminoalkylphosphonate diaryl esters as inhibitors of several serine proteases including dipeptidyl peptidase IV, cathepsin G, human neutrophil elastase, mast cell chymase and urokinase-type plasminogen activator. We have provided a historical perspective as well as a comprehensive report of the most recent studies in this field.
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