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JPET #52167
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A Selective and Oral Small Molecule Inhibitor of VEGFR-2 and
VEGFR-1 Inhibits
Neovascularization and Vascular Permeability
Neela Patel, Li Sun, Deborah Moshinsky, Hui Chen, Kathleen M.
Leahy, Phuong Le, Katherine G. Moss, Xueyan Wang, Audie Rice, Danny
Tam, A. Douglas Laird, Xiaoming Yu, Qingling Zhang, Cho Tang,
Gerald McMahon, and Anthony Howlett
SUGEN, Inc., South San Francisco, CA 94080 (N.P., L.S., D.M.,
H.C., P.L, K.G.M.,
X.W., A.R., D.T., A.D.L., X.Y., Q.Z., C.T., G.M., A.H.),
Pharmacia Corp. , Chesterfield
MO 63017 (K.M.L.)
Copyright 2003 by the American Society for Pharmacology and
Experimental Therapeutics.
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DOI:10.1124/jpet.103.052167This article has not been copyedited and
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Running Title: Indolinone VEGFR Kinase Inhibitor
Address Correspondence to:
Neela Patel, Ph.D.
Associate Director, Discovery Biology
SUGEN, Inc.
230 E. Grand Ave.
South San Francisco, CA 94080
Email: [email protected]
Tel: (650) 837 3565
Fax: (650) 837 3348
35 text pages
2 tables
7 figures
36 refs.
235 words in abstract
743 words in introduction
1290 words in discussion
Abbreviations:
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AMD, age-related macular degeneration; bFGF, basic fibroblast
growth factor; DR,
diabetic retinopathy; FGFR-1, fibroblast growth factor
receptor-1; HUVEC, human
umbilical vein endothelial cells; IL-8, interleukin-8; lyn,
LCK/YES-related Novel
tyrosine kinase; HGFR, hepatocyte growth factor receptor;
PDGFRb, platelet derived
growth factor receptor beta; SCF, stem cell factor; TNF, tumor
necrosis factor; VEGF,
vascular endothelial growth factor; VEGFR-1, vascular
endothelial growth factor
receptor-1; VEGFR-2, vascular endothelial growth factor
receptor-2.
Recommended section assignment: Other
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ABSTRACT Vascular epithelial growth factor (VEGF) is a key
driver of the
neovascularization and vascular permeability which lead to the
loss of visual acuity in
diabetic retinopathy and neovascular age-related macular
degeneration. Our aim was to
identify an orally active, selective small molecule kinase
inhibitor of VEGFR-2 with
activity against both VEGF-induced angiogenesis and vascular
permeability. We utilized
a biochemical assay to identify SU10944, a pyrrole indolinone,
which is a potent ATP-
competitive inhibitor of VEGFR-2 (Ki 21± 5 nM). In cellular
assays SU10944 inhibited
VEGF-induced receptor autophosphorylation (IC50 227± 80 nM) as
well as downstream
signaling (IC50 102 ± 27 nM). In biochemical assays SU10944
exhibits potent inhibitory
activity against VEGFR-1, weak activity against other related
subgroup members
including SCF-R, PDGFRβ, and FGFR-1, and no detectable activity
against other protein
tyrosine kinases such as EGFR, Src, and HGFR. In cellular assays
the selectivity for
SU10944 to inhibit VEGFR is maintained compared to other
tyrosine kinases (IC50 SCF-
R 1.6±0.3 uM, PDGFRβ 30.6±13.3 uM, FGFR-1>50 uM, EGFR>50
uM). Upon oral
administration, SU10944 gave a clear dose response in the
corneal micropocket model
with an ED50 for inhibition of neovascularization of ~30 mg/kg
and a maximum
inhibition of 95% at 300 mg/kg. Similarly, upon oral
administration in the Miles assay,
SU10944 potently inhibited VEGF-induced vascular permeability.
Our data indicate that
small molecule inhibitors of VEGFR signaling have the potential
to ameliorate both
VEGF induced neovascularization as well as vascular
permeability.
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In diabetic retinopathy (DR) and exudative age-related macular
degeneration (AMD),
vascular endothelial growth factor (VEGF) is a driver of both
the neovascularization and
retinal vascular permeability which underlie the loss of visual
acuity. The temporal and
spatial expression patterns of VEGF in these diseases implicate
VEGF in ocular
neovascularization. VEGF expression levels and activity are
elevated in vitreous samples
from diabetic patients with active proliferative retinopathy, as
compared to individuals
with non-proliferative diabetic retinopathy, quiescent
proliferative retinopathy, non-
diabetic individuals, or individuals with non-ischemic ocular
disease (Aiello et al., 1994,
Adamis et al., 1994). The levels of VEGF are positively
correlated to the clinically
observed degree and stage of retinal neovascularization (Aiello
et al., 1994). In AMD,
VEGF is expressed in choroidal neovascular membranes (Frank et
al., 1996).
As with ocular neovascularization, elevated levels of VEGF are
associated clinically with
ocular edema (Funatsu et al., 2002, Vinores et al., 1995). In
pre-clinical models, VEGF
is likewise associated with changes in vascular integrity.
Increases in ocular VEGF in
diabetic animals correlate with elevated vascular permeability,
prior to observable retinal
proliferative changes (Sone et al., 1997, Gilbert et al.,1998).
Local delivery of VEGF by
intravitreal implants result in significant increases in retinal
permeability by day 3 while
retinal neovascularization is observed only after 14 days
(Alikacem et al., 2000). A
similar breakdown of the blood-retinal barrier occurs in
primates administered VEGF by
intravitreal implant (Ozaki et al., 1997).
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Intervention in VEGF signalling, either by decreasing local
concentrations of ligand with
antisense oligodeoxynucleotides (Robinson et al., 1996) or
soluble chimeric receptors
(Aiello et al., 1995), or by inhibiting receptor signaling by
small molecules (Ozaki et al.,
2000) decrease ocular neovascularization, thus confirming the
central role of VEGF in
this process. Neovascularization in DR and AMD may differ
significantly from
angiogenesis in other pathological contexts such as tumor
angiogenesis where multiple
targets have been implicated, including PDGF, FGF, and IL-8
(Laird et al., 2000; Rofstad
and Halsor 2000; Shaheen et al., 2001). Current data strongly
suggests that in diabetic
retinopathy and exudative age-related macular degeneration, VEGF
is the key driver. In a
pre-clinical model of diabetic retinopathy, hypoxia-driven
retinal neovascularization,
VEGF inhibitors are efficacious but PDGFR inhibitors are not
(Ozaki et al., 2000);
similarly in a model of AMD, injury-induced choroidal
neovascularization, a VEGF
inhibitor decreases choroidal neovascularization ~ 85% but
administration of a PDGF
inhibitor does not decrease choroidal neovascularization (Kwak
et al., 2000). The
contribution of FGF to neovascularization is less well
understood. While bFGF is
expressed in neovascular membranes from AMD and DR patients
(Frank et al., 1996) and
intravitreally administered FGF has a synergistic effect with
VEGF in producing retinal
hemorrhage (Wong et al., 2001), retinal neovascularization
occurs upon hypoxic
challenge even in the absence of bFGF (Ozaki et al., 1998).
Moreover, overexpression of
bFGF in a transgenic mouse does not produce retinal
neovascularization nor increase the
degree of neovascularization upon hypoxia (Ozaki et al.,
1998).
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Inhibition of PDGF could have deleterious effects, particularly
in the context of DR.
PDGF appears to play a special role in retinal vasculature.
Recent reports suggest that
PDGF signaling is important for survival of retinal vasculature,
specifically pericytes,
under conditions of hypoxic or metabolic stress (Kodama et al.,
2001, Hammes et al.,
2002). In vivo, PDGF plays a role in retinal capillary coverage:
PDGF-B deficient mice
(PDGFB +/-) have fewer retinal pericytes and more acellular
retinal capillaries than wild-
type controls, differences which are more pronounced in diabetic
animals. In the
hypoxia-induced model of neovascularization, new vessels are
twice as prevalent in
PDGFB +/- mice compared to wild-type animals (Hammes et al.,
2002), suggesting that
pericyte deficiency renders endothelial cells more susceptible
to angiogenesis. Therefore,
a potential side effect of PDGF inhibition could be to
accelerate the loss of pericytes
which are implicated in the destabilization of blood vessels
during the early stages of
diabetic retinopathy. Therefore, given the lack of data
supporting a role for PDGF in
retinal and choroidal neovascularization as well as the risk of
increasing pericyte dropout,
a VEGF-selective inhibitor is likely the best choice for
treatment of retinopathies.
Our goal was to identify and characterize an orally available
selective VEGFR inhibitor,
since angiogenesis and increased vascular permeability in DR and
exudative AMD is
primarily or solely driven by VEGF in these settings. A
selective compound should
minimize the potential toxicities resulting from the inhibition
of additional kinases and be
more likely to give a sufficient therapeutic index for the
treatment of non-life threatening
disease.
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MATERIALS AND METHODS
Chemicals. SU10944 was synthesized as described in patent
WO00/08202. Its chemical
structure is shown in Fig. 1. Stock solutions of SU10944 were
made in DMSO and stored
at –20°C. Dilutions for assays were made fresh prior to use.
Kinetic Analysis. The catalytic portion of mouse VEGFR-2 was
expressed as a GST
fusion protein following infection of Spodoptera frugiperda
(sf9) cells with engineered
baculoviruses by standard methods (King and Possee, 1992).
GST-VEGFR-2 was
purified to homogeneity from infected sf9 cell lysates by
glutathione sepharose
chromatography (Smith and Johnson, 1988). GST-fusion
preparations were analyzed by
gel electrophoresis and determined to be of high purity, with no
detectable breakdown
products as determined using protein staining and Western blot
analysis for the GST
moiety. Protein concentration was determined with the Bio-Rad
protein assay reagent kit
(Bio-Rad, Hercules, CA) using a standard curve with bovine serum
albumin. For the
SU10944 Ki determination against VEGFR-2, biochemical kinase
reactions were
performed as described below (Biochemical Assays) in the
presence of various ATP and
inhibitor concentrations. Rates were expressed as the increase
in TR-FRET counts per
minute of reaction within the linear reaction time. Ki values
were determined by
graphical analysis of the plot of the slopes from the double
reciprocal plot vs. inhibitor
concentration. The final Ki value represents the average + the
standard deviation from 5
independent experiments.
Biochemical Assays. VEGFR-2 and PDGFRβ time-resolved
fluorescence (TR-FRET)
autophosphorylation assays were performed as described
(Moshinsky et al., in press).
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Briefly, 1nM VEGFR-2 or 5nM PDGFRβ was added to a reaction
buffer composed of
50mM Hepes, pH 7.4, 1mM MnCl2, 0.01% BSA, and 1mM DTT,
containing twice the
apparent Km concentration of both ATP and N-terminal
biotinylated peptide (KY-tide:
KYKKYKKKYKKKKYKYK) in 50uL total volume. Reactions were allowed
to
proceed within the linear reaction time then terminated by the
addition of 20uL 90mM
EDTA. Eu-W1024-labeled antiphosphotyrosine PY20 and
Streptavidin: SureLight –
Allophycocyanin (Perkin Elmer Life Sciences, Foster City, CA)
were diluted in TBS
containing 0.02% BSA and 0.1% Tween20 and added to a final
concentration of 0.5nM
and 1.6nM, respectively. After incubation for at least 10
minutes, samples were excited
at 340nM and emissions were read at 665nM using an LJL Analyst
(LJL Biosystems,
Sunnyvale, CA). The increase in signal for both assays was
determined to be time
dependent with a requirement for ATP, peptide, kinase, and
divalent metal cation. FGFR-
1 and EGFR autophosphorylation reactions were performed using
immuno-captured
kinases as described (Laird et al., 2000). The HGFR and VEGFR-1
assays were
performed using poly(Glu, Tyr) 4:1 as a substrate as described
(Blake et al., 2000). For
the cdk2/cyclin A assay, a Scintillation Proximity Assay method
was used (Amersham
Pharmaceutical Assays Development Group, 1995). The SCF-R, src,
lyn, and fyn assays
were performed in standard TR-FRET format (Kolb et al., 1998)
using peptide substrates
found by screening an internal peptide substrate library.
Conditions for compound
testing were saturating peptide concentration and an ATP
concentration of 2*Km.
VEGFR-2 Cell Autophosphorylation Assay. Full-length mouse
VEGFR-2 was cloned
into the C-terminal 3xFLAG-tag expression vector p3xFLAG-CMV-14
(Sigma, St.
Louis, MO). Human embryonic kidney 293T cells were grown in DMEM
supplemented
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with 10% fetal bovine serum and grown in a 37°C humidified
incubator with 5% CO2.
The construct was transfected into 293T cells using
LipofectAmine2000 (Invitrogen,
Carlsbad, CA) under manufacturer’s recommendations. Following
transfection, cells
were starved in DMEM containing no serum and 0.1% BSA for 24
hours. Cells were
then split into 96-well plates and treated with compound in a
final concentration of 1%
DMSO for 2 hours. Cells were lysed by the addition of HNTG (50mM
Hepes, pH 7.4,
150mM NaCl, 1.5mM MgCl2, 10% glycerol, 1% Triton X-100, 1mM
EGTA), and lysates
were transferred to polystyrene 96-well plates that had been
pre-coated with 1ug per well
of M2 anti-FLAG monoclonal antibody (Sigma, St. Louis, MO) to
capture the 3xFLAG
tagged kinase. Quantification of phosphorylation was performed
by incubating with
HRP-labeled anti-phosphotyrosine PY99 (Santa Cruz Biotechnology,
Santa Cruz, CA),
followed by detection with an ABTS color readout.
VEGFR-2 Phosphorylation Detected by Western Blot. NIH/3T3 cells
stably
expressing VEGFR-2 were grown to confluence in DMEM with 10%
heat-inactivated
calf serum and then incubated in serum-free medium containing
different concentrations
of SU10944 for 20 hr. After stimulation with human recombinant
VEGF165 (R&D
Systems, Minneapolis, MN) at 50 ng/ml for 10 min, cells were
lysed in lysis buffer
containing 50 mM Hepes, 150 mM NaCl, 10% glycerol, 0.5% Triton
X-100, 100 mM
PMSF, 1 mM sodium vanadate and 2 µg/ml leupeptin and aprotinin.
VEGFR-2 protein
was isolated with a monoclonal anti-mouse VEGFR-2 antibody made
at SUGEN
designated L4. Phosphorylation of VEGFR-2 was then analyzed by
SDS-PAGE
followed by western blotting using biotin-labeled
anti-phosphotyrosine PY99 (Santa Cruz
Biotechnology, Santa Cruz, CA). Total VEGFR-2 levels were
assessed by stripping and
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re-probing the membrane with anti-VEGFR-2 antibody L4 (SUGEN,
South San
Francisco, CA).
VEGFR-2 Cellular Functional Assay. The assay is an immunoassay
for the quantitative
detection of human tissue factor. HUVECs were seeded at 50,000
cell/well in growth
medium EMB (Endothelial Cell Basal Medium, BioWhittaker,
Walkersville, MD) +
10%FBS with complete supplements containing hEGF 0.5 ml/500 ml,
hydrocortisone
0.5 ml/500 ml, gentamicin sulfate amphotericin-B 0.5 ml/500 ml,
bovine brain extract 2
ml/500 ml (BioWhittaker, Walkersville, MD) in 24 well plate
coated with 0.2% gelatin.
Cells were grown for 2 days >90% confluence, and then treated
with SU10944 and
human recombinant VEGF165 (R&D Systems, Minneapolis, MN) at
50 ng/ml or PMA
(phenylmethylsulfonyl fluoride, Sigma, St. Louis, MO) at 10 nM
for 4 hours; negative
control cells received media only. Cells were lysed with HNTG
lysis buffer (50mM
Hepes, pH 7.4, 150mM NaCl, 1.5mM MgCl2, 10% glycerol, 1% Triton
X-100, 1mM
EGTA), plus EDTA (1:100), protease inhibitor (1:100), and PMSF
(1:100) in 125 ul per
well for 20 minutes at 4 °C. Cell lysates were were processed
using an IMUBIND Tissue
Factor ELISA Kit (American Diagnostica, Greenwich, CT). Briefly,
lysates were
transferred to 96 well format precoated micro-test strips and
incubated at 4 °C overnight.
The micro-test strips were washed 4 times with wash buffer, 100
uL of biotinylated anti-
human tisuue factor antibody added to each well, and incubated
for 1 hour at room
temperature. After washing 4 times, 100 uL of diluted
streptavidin-horseradish
peroxidase conjugated antibody was added to each well, and
incubated for 1 hour at room
temperature. The wells were washed 4 times with wash buffer and
incubated with 100 ul
of TMB (tetramethyl-benzidine, Sigma, St. Louis, MO) substrate
solution for 20 minutes
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at room temperature. The reaction was stopped by the additional
of 50 ul of H2SO4
solution, and absorbance read on a microplate reader at a
wavelength of 450 nm.
Functional Cellular Assays for SCF-R, EGFR, PDGFRβ, and FGFR-1.
For EGFR,
PDGFRβ, and FGFR-1, the functional activity of the receptor was
measured by a ligand-
induced BrdU incorporation assay in 3T3 cells which endogenously
express both FGFR-
1 and PDGFRβ but not EGFR. A single cell line stably transfected
with EGFR was used
for all assays. Cells seeded in a 96-well plate were made
quiescent by serum-deprivation
for 24 hr and then stimulated with FGF2/bFGF (1.5 nM), EGF (4
nM) or PDGF (3.8 nM)
in the absence or presence of the indicated concentrations of
SU10944 for 20 hr. BrdU
was added for a two hr labeling period, and the cells were
fixed. The amount of BrdU
incorporation was determined with an anti-BrdU-peroxidase
conjugated antibody using
an ELISA kit (Boehringer Mannheim). Cell viability following
exposure to compounds in
the assay format was assessed by substituting the addition of
BrdU with resazurin (1
mg/ml) at 1:100 after the 20 hour incubation. Following a 3 hr
incubation, the absorbance
of the samples are measured at 630 nm in "dual wavelength" mode
with a filter reading at
450 nm, as a reference wavelength) on a Dynatech ELISA plate
reader. Compound was
added to wells in the dilutions used to determine IC50 values,
including a negative
control in which all components were included except for cells
and a positive control in
which all components except compound were added. The resazurin
assay is based upon
the conversion of the dark blue resazurin dye to a pink dye in
proportion to the metabolic
activity of the cells (O’Brien et al. 2000). For each IC50
value, the standard deviation is
reported.
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The functional activity of SCFR was assessed using a
SCF-dependent cell proliferation/
survival assay. MO7e cells were grown and expanded in RPMI with
10%FBS in the
presence of IL-3 (10ng/ml) and GM-CSF (10 ng/ml). After
counting, cells were pelleted
by centrifugation and washed twice with PBS. Cells were
resuspended in medium
containing either IL-3 or SCF (100ng/m) and aliquoted into
96-well plates at 50,000 cells
per well along with varying concentrations of SU10944. After
incubation for three days
at 37°C in a humidified incubator with 5% CO2, live cells were
quantified by their ability
to metabolize resazurin as described above.
Corneal Angiogenesis Model. An intrastromal pocket was
surgically created in one or
both corneas of each anesthetized Sprague Dawley female rat. A
slow release
hydron/sucralfate pellet containing 150 ng human recombinant
VEGF165 (PeproTech,
Rocky Hill, NJ) was inserted into the pocket as previously
described (Kenyon 1996), the
pocket closed to self-seal, and the rats given analgesia and
topical antibiotic ointment
applied once to the eye. The rats recovered from anesthesia on a
warming pad, and were
returned to their cages. SU10944 was administered daily by
gavage in a 1.0 ml
suspension of 0.5% methylcellulose (Sigma Chemical, St. Louis),
0.025% Tween-20
(Sigma) to four to six rats per dose group beginning the day
before implant, and
continuing the length of the study. Four days after surgery, the
corneas of the re-
anesthetized rats were examined under a slit lamp microscope and
the neovascular
response was quantified by measuring the average new vessel
length (VL), the corneal
radius (r = 2.6 mm), and the contiguous circumferential zone (CH
= clock hours where 1
CH = 30 degrees), and applied to the formula: Area (mm2)=
(CH/12) x 3.14(r2- (r-VL)2 ).
The rats were then immediately euthanized. Eyes from rats that
developed infection as a
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result of the surgery were not included in the study. Six to
eight eyes were included per
dosing group. The neovascular areas of the vehicle and 250mg/kg
dosed rat corneas were
dissected from the eye, flat mounted, and photographed at 4x
with a digital camera
mounted on a microscope. Rat corneas implanted with pellets
containing no growth
factor (placebo pellets) generated no new blood vessel growth
(Leahy et al). All animal
treatment protocols were reviewed by and were in compliance with
Pharmacia’s
Institutional Animal Care and Use Committee.
Miles assay for vascular permeability. The Miles assay for
vascular permeability
(Miles and Miles 1952) was adapted to athymic mice as follows.
Mice were given a
single oral dose of SU10944 or vehicle alone. Simultaneously or
at designated later
timepoints, 100ul of 0.5% Evan’s blue dye (Sigma-Aldrich, St.
Louis, MO) in PBS was
administered intravenously via the tail vein. One hour later,
mice were injected
intradermally (in duplicate sites on their backs) with 400 ng of
VEGF (human
recombinant VEGF165, R&D Systems, Minneapolis, MN) dissolved
in 20 ul of PBS or (in
adjacent duplicate sites) with PBS alone. After an additional 30
minutes, VEGF-
dependent dye leakage from the vasculature into skin was
assessed visually and scored
semi-quantitatively (100, 50, or 0% inhibition for each spot).
Two spots per animal
allowed each animal to be scored as representing 100%, 75%, 50%,
25%, or 0%
inhibition. Low-level background effects from time-matched
vehicle-treated groups were
subtracted out.
Determination of SU10944 Plasma Levels. Plasma samples (100 µl),
SU10944 standard
or quality control samples in mouse blank plasma were mixed with
acetonitrile (300 µl)
containing DL-propranolol hydrochloride (internal standard) in a
96-well polypropylene
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plate (Orochem Technology, Westmont, IL). The plate was mixed by
vortex for 1 minute
and the samples were centrifuged for 10 minutes at 4000 rpm. Ten
µL of the supernatant
was injected onto the LC/MS/MS system where separation occurred
on a BDS
HYPERSIL C-18 (5 µm, 100 x 4.6 mm) reverse-phase HPLC column
(Keystone
Scientific, Foster City, CA). The amount of SU10944 and the
internal standard in each
mouse plasma sample was quantified based on standard curves
generated using known
amounts of compound ranged from 5 to 10,000 ng/mL. Standard
curve samples and
quality control samples of SU010944 were prepared by spiking 10
µL of stock standard
solutions with 90µL of blank plasma.
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RESULTS
We identified SU10944 (Fig. 1) as a potent inhibitor of VEGF-R
as part of our efforts to
synthesize and characterize indolin-2-ones as inhibitors of
class III receptor tyrosine
kinases (Yarden and Ullrich, 1988). Previous structure-activity
relationship studies
revealed that modifications on the indolin-2-one core could
generate compounds with
different kinase selectivity profiles (Sun et al. 1998, 1999,
2000). Modifications of the
core have also been found to have a dramatic impact on the
pharmaceutical properties of
the compounds in terms of solubility, metabolic stability,
permeability, plasma protein
binding, and pharmacokinetic properties.
Following the identification of SU10944 as an inhibitor of
VEGFR-2 in biochemical
assays (IC50 96 ± 20 nM), we went on to further assess the
biochemical activity of the
compound against other kinases, as well as its potential to act
as a competitor for ATP.
SU10944 exhibited competitive inhibition with respect to ATP for
VEGFR-2. This is
indicated by the fact that the lines from the double reciprocal
plot (Fig. 2) converge on
the Y-axis. The Ki was determined to be 21 + 5 nM. In a panel of
kinase assays,
SU10944 potently inhibited VEGFR-2 with an IC50 of 96 ± 20 nM
and exhibited even
greater activity against VEGFR-1 IC50 6±1 nM (Fig. 3). It showed
some activity against
other closely related family members, for example PDGFRβ and
FGFR-1 but exhibited
significantly less activity against other receptors (Table 1).
The compound is not a pan-
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kinase inhibitor as no discernible inhibition of more distantly
related tyrosine kinases was
evident e.g. EGFR and Src.
We then used a panel of three cell-based assays to determine
whether the compound
could cross the cellular membrane and inhibit VEGFR-2 within
cells. In 293T cells
transiently transfected with mouse VEGFR-2, SU10944 exhibited an
IC50 value of 227 ±
80 nM for receptor autophosphorylation as measured by ELISA
(Figure 4A). Similarly,
in an assay to assess the functional activity of endogenous
VEGFR-2 in HUVECs, tissue
factor production stimulated by VEGF was inhibited with an IC50
of 102 ± 27 nM (Figure
4B). However SU10944 did not inhibit TNF-stimulated release of
tissue factor (Figure
4B). Final confirmation that the compound inhibits VEGFR-2
receptor phosphorylation
was obtained by western blot analysis of 3T3 cells engineered to
express mouse VEGFR-
2 (Figure 4C). After cells were pre-treated with compound and
stimulated with VEGF,
VEGFR-2 was immunoprecipitated and then detected on the blot
with an anti-
phosphotyrosine antibody. SU10944 inhibited receptor
autophosphorylation confirming
the results of the 293 assay in a different cell type and assay
format.
For kinases against which SU10944 was active in biochemical
assays, cellular assays
were performed to determine whether the activity was maintained
in a more physiological
setting (Table 2). Similar to the observations in the panel of
biochemical assays,
significantly higher concentrations of compound were required to
inhibit closely related
subgroup of family members such as FGFR-1, SCF-R, and PDGFRβ
than was required
for the inhibition of VEGFR-2 in functional assays. The
functional activities of the
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FGFR-1, PDGFRβ, and EGFR were measured by ligand-induced cell
proliferation of
3T3 cells as measured by BrdU incorporation. For SCFR, the
activity was measured by
the SCF-dependent survival of MO7e cells. SU10944 did not
exhibit detectable inhibition
of EGFR or FGFR-1 (IC50 values >50 uM). Furthermore SU10944
was not cytotoxic: the
LD50 was >50 µM for 3T3 cells.
We then assessed the ability of SU10944 to inhibit angiogenesis
and vascular
permeability in vivo when administered by the oral route. In the
rat corneal micropocket
model of angiogenesis, a VEGF pellet is implanted in the cornea
to stimulate
neovascularization. In this model, the compound significantly
decreased the
angiogenesis, both the number of vascular sprouts as well as the
length of the sprouts
(Fig. 5). Moreover, SU10944 gave a clear dose response with an
ED50 of ~ 50 mg/kg for
inhibition of neovascularization (Fig. 6). In order to increase
potential exposure levels
upon oral administration by decreasing dissolution rate
limitations of the compound, we
prepared an in situ sodium salt of SU10944 and repeated the
experiment. The sodium salt
decreased the apparent ED50 slightly, to approximately 30 mg/kg.
In addition a
maximum inhibition of 95% was achieved at the highest dose of
300 mg/kg. Increased
exposure upon administration of the sodium salt was confirmed in
pharmacokinetic
studies (data not shown).
In the Miles assay for vascular permeability (Miles and Miles,
1952), a visualization dye
is administered intravenously, followed by a bolus
administration of VEGF by the
intradermal route. Leakage of the dye in the skin indicates a
local increase in vascular
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permeability. In this assay, SU10944 inhibited VEGF-induced
vascular permeability in a
time and dose dependent manner (Fig 7a). Maximum inhibition was
observed at the
earliest timepoint (1 hr), with decreasing levels of inhibition
over the course of 24 hr. At
100 mg/kg the maximum inhibition of vascular permeability was
sustained to 2 hr, and a
50% reduction in response was still apparent 24 hr post-dose. In
contrast, at the 30 mg/kg
dose maximum inhibition was seen at 1 hr and decreased
dramatically over 24 hr to zero.
Plasma concentrations of SU01944 were also determined and
correlated with inhibition
(Fig 7b). Based on the total data set that reflects a range of
doses and times post-
administration of compound, we conclude that SU10944 plasma
exposures of 250 ng/ml
(844 nM) result in 50% inhibition of VEGF-mediated vascular
permeability in vivo.
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DISCUSSION
We have identified and characterized a new small molecule
inhibitor of VEGF signaling
which inhibits both angiogenesis and vascular permeability when
administered orally.
The compound, SU10944, is a potent ATP-competitive inhibitor of
the kinase activity of
purified VEGFR-2 with a Ki of 21± 5 nM and an IC50 of 96± 20 nM.
SU10944 also
exhibits potent activity against VEGFR-1, with an IC50 of 6±
1nM. The compound
maintains activity in cellular assays against both the
mouse(flk-1) and human (KDR)
forms of VEGFR-2. In the cellular assay for inhibition of
VEGFR-2 autophosphorylation
the compound exhibited an IC50 of 227± 80 nM, a value in good
agreement with the IC50
obtained in our HUVEC assay for receptor function (IC50 102± 27
nM). In the functional
assay, we demonstrated that the inhibition of tissue factor
production resulted from
inhibition of VEGFR by showing that TNF-stimulated release of
tissue factor was not
affected by the compound. The HUVEC data suggest that the
compound will not only be
functionally active against human VEGFR-2 but should be active
in endothelial cells
which are the target for inhibition of angiogenesis and vascular
permeability. We were
further able to confirm the nature of the cellular activity of
SU10944 by visualizing
inhibition of receptor phosphorylation by Western analysis.
From a panel of biochemical and cellular assays we conclude that
SU10944 is a relatively
selective inhibitor with a strong preference for VEGFRs. The
kinase activity of both
VEGFR-1 and VEGFR-2 are potently inhibited by the compound, with
low to mid
nanomolar IC50 values. Some activity was observed in the
biochemical assays,
particularly against other members of the class III receptor
tyrosine kinases, for example,
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PDGFRβ 1000±� 83 nM and SCFR 1580± 270 nM. SU10944 exhibited
little or no
activity against the other kinases surveyed, which represent a
range of tyrosine kinases as
well as some serine/ threonine kinases. We limited our cellular
assays to those kinases
which had shown activity in biochemical assays, plus one more
distantly related kinase.
Consistent with the biochemical observations, the compound
displayed very limited cross
reactivity in the cellular functional assays. The most notable
cross reactivity occurred
against SCFR (IC50 1600± 300 nM; however, compared to VEGFR
functional readout
(IC50 102± 27 nM), there was an 18-fold selectivity. Selectivity
at the cellular level was
good compared to PDGFRβ (IC5030.6±�13.3 uM), with a ratio of
340x.
While SU10944 inhibited the kinase activity of purified VEGFR-2
with an IC50 of 96± 20
nM, when the compound was tested in cellular assays the apparent
IC50 value shifted to
227± 80 nM in the autophosphorylation assay but was in very
close agreement with the
functional assay IC50 of 102 ±27 nM. A more notable discrepancy
was observed between
the IC50 values for the kinase activity of purified PDGFRβ
(1000±�83 nM) compared to
the cellular assay (IC5030.6±�13.3 uM). Modest discrepancies
between the biochemical
and cellular values are not uncommon findings with small
molecule enzyme inhibitors
since the physical properties of the compounds as well as the
assay format become
important when translating from biochemical to cellular
activity. To inhibit activity of
the kinase within a cellular context, the compound must cross
the cell membrane and
retain activity in the presence of cellular proteins. In
addition, differences in assay
formats can influence the observed IC50 values.
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In the context of potential treatments for exudative AMD and
diabetic retinopathy, a
therapeutic molecule should inhibit both neovascularization as
well as increased vascular
permeability in order to be maximally effective. In pre-clinical
models, SU10944
potently inhibited both VEGF-induced angiogenesis and vascular
permeability following
administration by the oral route. In the corneal micropocket
model of angiogenesis, we
observed a clear dose response with a maximal inhibition of
nearly 100%. Compound
levels obtained by oral administration clearly achieved
sufficient exposure levels to
inhibit the functional activity of the receptor. Similarly, in
the Miles assay of vascular
permeability, a time and dose dependent response was observed.
The maximum response
was nearly 100%, confirming that in a second species and
different model,
pharmacologically relevant levels of drug were achieved by oral
administration.
We note that there is a discrepancy between the in vitro potency
of SU10944 (227± 80
nM in the autophosphorylation assay, 102±27 nM in the functional
assay) and its in vivo
activity as measured in the vascular permeability assay (EC50
cutoff estimated to be 833
nM). While there can be many reasons for such observed
differences in the translation
from in vitro to in vivo results, the main source in this case
is likely to be high plasma
protein binding of the compound. Other compounds in the series
have been shown to
have high protein binding; our data suggest protein binding of
>98% (data not shown).
The unbound concentration of SU10944 in vivo would therefore be
roughly 16 nM, a
value in better agreement with our in vitro observations.
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To develop the compound for potential use in the treatment of
human ocular disease, the
efficacy of the compound in more relevant disease models, as
well as the potential safety
of the compound with systemic administration will require
careful investigation. The
activity of the compound in human retinal endothelial cells and
in more physiologically
relevant animal models such as retinal vascular permeability in
streptozoticin-induced
rats, hypoxia-driven retinal angiogenesis in mouse neonates, and
laser injury-induced
choroidal neovascular model of AMD should be determined. In
addition, the therapeutic
index of such compounds will be essential to establish because
of potential mechanism-
based toxicities with systemically administered VEGFR
inhibitors, particularly in the
context of diabetic co-morbidities. One area of particular
concern is whether VEGFR
inhibitors will further impair coronary collateral formation in
diabetic patients, in
response to myocardial ischemia. Collateral vessel development
occurs by the process of
arteriogenesis, the expansion of existing arterioles, a process
which is differentially
regulated from angiogenesis. Monocyte migration into the area of
ischemia is a key
process in the formation of the collateral vessels. Current
evidence suggests that
impaired collateral vessel formation in diabetic individuals
results from a signaling defect
downstream of VEGFR-1 in monocytes (Waltenberger, 2001), which
normally respond
to VEGF-A by increased migration. Since the signaling via
VEGFR-1 is already impaired
in diabetic patients, it remains to be determined whether a
VEGFR-1 inhibitor would be
additive or neutral for collateral formation, or whether a
selective VEGFR-2 inhibitor
would be preferred for treatment of diabetic retinopathy.
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Unlike VEGFR-2, VEGFR-1 has not been implicated directly in
mediating VEGF-
induced angiogenesis or vascular permeability. However,
inhibition of VEGFR-1 may be
beneficial in AMD where macrophage infiltration has been
suggested to play a role in the
etiology of the disease. VEGF produced by hypoxic retinal
pigmented epithelial cells has
been postulated to act as a chemotactic factor for macrophages,
which can then secrete
additional VEGF as well as other pro-angiogenic factors.
We have identified and characterized a novel small molecule
inhibitor of VEGFR-2,
SU10944. This compound is a potent, ATP-competitive inhibitor of
VEGFR-2
biochemical activity and is active in the nanomolar range in
cellular assays. SU10944 can
be administered in vivo by the oral route and achieves
sufficient exposure to inhibit
nearly all VEGF-stimulated neovascularization and vascular
permeability. A selective,
well-tolerated VEGFR inhibitor, administered orally or by local
delivery, should be of
therapeutic benefit in both diabetic retinopathy and exudative
age-related macular
degeneration. In addition we believe compounds of this nature
will be valuable in
delineating the role of VEGF in various forms of pathological
angiogenesis where
multiple kinases may play contributing roles. Clinical trials
will be necessary to show the
potential prophylactic or therapeutic utility of these novel and
selective VEGFR-
inhibitors in human eye diseases. The discovery of a multiple
VEGFR inhibitors
representing a variety of pharmacophores offers the opportunity
to generate new
molecules with increased potency or improved pharmaceutical
properties by rational
design based on co-crystals with VEGFR-2.
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ACKNOWLEDGMENTS
We thank Steve Settle for excellent technical work on the
corneal angiogenesis model,
the Discovery Technology Group at SUGEN for biochemical cross
screening data, Ken
Lipson for assistance with data analysis, and Anne-Marie
O’Farrell for suggestions
regarding the manuscript.
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Legends for figures
Fig. 1. Chemical structure of SU10944.
Fig. 2. Double reciprocal plot of SU10944 inhibition of VEGFR-2
at 160nM (∇), 80nM
( �����Q0�� ���DQG�QR��
��FRPSRXQG��7KH�LQVHW�GLVSOD\V�D�JUDSK�RI�WKH�VORSHV�IURP�WKH�
double reciprocal plot vs. inhibitor concentration. The –(Ki)
value is indicated by an
arrow. Data are representative from 5 independent
experiments.
Fig. 3. Dose response curves for SU10944 inhibition of VEGFR-2
(a) and VEGFR-1 (b).
Error bars represent standard deviations from at least 3
replicate measurements.
Fig. 4. SU10944 inhibits VEGFR-2 receptor phosphorylation and
functional activity in
cells. (a) 293T cells transiently transfected with mouse VEGFR-2
were exposed to
varying concentrations of SU10944 and receptor phosphorylation
detected by ELISA.
Error bars represent standard deviations from n=3. (b) HUVECs
were stimulated with
either PMAn or VEGFu. Production of tissue factor was measured
by ELISA. Error
bars represent standard deviations from duplicate samples. (c)
3T3 cells stably transfected
with mouse VEGFR-2 were stimulated with VEGF. Anti-mouse VEGFR-2
antibody was
used for immunoprecipitation from cellular lysates. Western
analysis of
immunoprecipated proteins was performed using
anti-phosphotyrosine antibody or anti-
mouse VEGFR-2 antibody.
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Fig. 5. SU10944 inhibits VEGF-induced neovascular growth in rat
corneas. Four days
after VEGF pellet implant, the neovascular area of the rat
cornea was measured, dissected
from the eye, flat mounted, and photographed at 4x with a
digital camera mounted on a
microscope. Panel A shows representative corneas from rats dosed
with vehicle. Panel B
shows corneas from rats dosed with SU10944 free acid at 250 mg
per kg.
Fig. 6. Dose response of sodium salt or free acid SU-10944 on
VEGF-induced
neovascular growth in rat corneas. Four days after VEGF pellet
implant, the neovascular
area of the rat cornea was measured using a calibrated eye piece
mounted on a slit lamp
microscope. Vessel length, and the contiguous circumferential
angle of
neovascularization were noted and used to calculate the area of
each cornea that was
covered with neovessels. This area was expressed as a percent of
the average vehicle-
dosed, VEGF-implanted (Control) corneas. Error bars represent
SEM for n=6-8 eyes.
*p
-
JPET #52167
35
three animals), with any low-level background effects from
time-matched vehicle-treated
groups subtracted out. Error bars represent SE. Vascular
permeability in animals treated
with SU10944 at 100 mg/kg and 30 mg/kg was significantly
inhibited (* indicates p
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JPET #52167
36
TABLE 1
Biochemical Activity of SU10944
Kinase Mean IC50,
µM
Kinase IC50, µM
VEGFR-2 0.096 ± 0.020
(n=16)
EGFR
(n=4)
>20
VEGFR-1 0.006 ±0.001
(n=5)
Src
(n=8)
>20
FGFR1 1.60±0.87
(n=5)
Lyn
(n=3)
>20
PDGFRβ 1.00±0.08
(n=4)
Fyn
(n=5)
>20
SCFR 1.58±0.27
(n=5)
Cdk2
(n=4)
>20
HGFR
(n=8)
>20
The IC50 values for SU10944 were determined by measuring
autophosphorylation or
substrate phosphorylation, as specified in Materials and
Methods.
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TABLE 2
Cellular Activity of SU10944
Receptor (cell type) IC50, µM
SCFRa (MO7e) 1.6 ±0.3
(n=2)
PDGFRβ b (3T3) 30.6±13.3
(n=6)
EGFRb (3T3) >50
(n=3)
FGFR-1b (3T3) >50
(n=3)
a Functional activity of SCF-R was measured in a survival assay
for MO7e cells.
bFunctional activity was measured by BrdU incorporation in 3T3
cells stably transfected
with the receptor.
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NH
O
NH
OHO
Figure 1
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0.001 0.01 0.1 1 10
0
50
100
Concentration (µM)
% in
hibi
tion
Figure 4
a
c
b
VEGF (100ng/ml)
anti-pTyr
anti-Flk-1
25 15 0.2
- +++++
SU10944 [µM]
0.001 0.01 0.1 1 100.0
0.5
1.0
1.5
Concentration (µM)
O.D
. at
450
nm
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Figure 5
VEGF pelleted corneas—systemic treatment with vehiclea
b VEGF pelleted corneas—systemic treatment with SU10944
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Figure 6
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1
10
100
1000
10000
100000
100 75 50 25 0
% InhibitionP
lasm
a S
U10
944
conc
. (ng
/mL)
250 ng/mL
Figure 7
0
10
20
30
40
50
60
70
80
90
100
0 4 8 12 16 20 24
100 mg/kg
30 mg/kg
3 mg/kg
% In
hibi
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Time Post Administration (h)
* * **
*
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