School of Biological Sciences, University of East Anglia, Norwich, NR4
7TJ, UK
* Present address: St. Vincent's Institute of Medical Research, 9 Princes
Street, Fitzroy, Victoria 3065, Australia
Author for correspondence (e-mail:
dylan.edwards{at}uea.ac.uk)
Accepted 10 June 2002
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Summary |
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Key words: MT-MMPs, Fibrin gels, Tubulogenesis, Endothelial, Angiogenesis
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Introduction |
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The MMPs are a family of Zn2+-binding, Ca2+-dependent
endopeptidases that play a crucial role in the degradation of the components
of the extracellular matrix (ECM)
(Coussens and Werb, 1996). The
MMPs are involved in a variety of physiological processes and also diseases
characterised by pathological tissue destruction
(Nelson et al., 2000
), all of
which are associated with angiogenesis and vascular remodelling. There are
more than 20 members of the MMP family presently described, and they are
subdivided into five main categories on the basis of their structural
similarities and substrate preferences. These are the collagenases, the
gelatinases, the stromelysins, the membrane-type MMPs (MT-MMP) and a
heterogeneous subgroup (i.e. MMP-7, MMP-12, MMP-19, MMP-20)
(Murphy and Knäuper,
1997
; Murphy et al.,
2000
). The MT-MMP subgroup comprises six members, four of which
(MT1-, MT2-, MT3- and MT5-MMP) possess a transmembrane domain and a short
cytoplasmic tail at the C-terminal region of the protein. The transmembrane
domain anchors the enzymes to the cell surface, whereas the cytoplasmic tail
may interact with intracellular proteins that regulate function or
localisation. The transmembrane MT-MMPs are also distinctive as they have an
`MT-loop' in the catalytic domain that is not found in other MMPs
(English et al., 2001
).
MT4-MMP and MT6-MMP lack the MT-loop, the transmembrane domain and cytoplasmic
tail, but are localised to the cell membrane by a glycosylphosphatidylinositol
(GPI) anchor, suggesting that these enzymes represent a functionally distinct
branch of the MMP family (Murphy et al.,
2000
). Matrix remodelling by the MMPs can be regulated by four
tissue inhibitors of metalloproteinases (TIMPs)
(Murphy et al., 2000
).
Since the first 2D cultures of endothelial cells (EC) derived from the
human umbilical vein in the early 1970s
(Jaffe et al., 1973), great
improvements in our understanding of EC function and EC responses to various
factors have been achieved. The major disadvantage, however, to this type of
in vitro study was that the EC milieu was too simple, as EC normally reside
and interact within a 3D environment. Consequently, attention has shifted to
culturing EC or tissue explants (with intact blood vessels) in several
different types of 3D matrices such as type I collagen gels, Matrigel and
fibrin gels in order to more closely mimic the in vivo cellular
micro-environment (Hiraoka et al.,
1998
; Ilan et al.,
1998
; Vernon and Sage,
1999
). When cultured within these 3D matrices, EC respond to known
angiogenic factors, that is, vascular endothelial growth factor (VEGF) and
fibroblast growth factor-2 (FGF-2)
(Anand-Apte et al., 1997
),
forming `tube-like structures' with patent lumens
(Meyer et al., 1997
;
Yang et al., 1999
). A fibrin
matrix is often found surrounding `leaky' tumour-associated blood vessels or
surrounding blood vessels at sites of vascular injury
(Nagy et al., 1989
). This
fibrin barrier acts as a provisional matrix, through which EC must penetrate
to facilitate capillary sprouting. Therefore culturing EC within a 3D fibrin
gel is a useful model system for studying EC sprouting.
MT1-MMP has been implicated in the neovascularisation process, as it can
act as a potent pericellular fibrinolysin that enhances invasion and formation
of tubular structures in fibrin gels when overexpressed in Madin-Darby canine
kidney (MDCK) cells (Hiraoka et al.,
1998; Hotary et al.,
2000
). Furthermore, MT1-MMP expression is increased in human
dermal microvascular EC by angiogenic factors such as VEGF, tumour necrosis
factor-
(TNF-
) and FGF-2, and EC assembly into capillary-like
structures within collagen gels was delayed by pre-incubation with
anti-MT1-MMP antibodies (Chan et al.,
1998
). Finally, in an in vivo corneal angiogenesis assay,
MT1-MMP-null mice failed to exhibit an angiogenic response to FGF-2 unlike
their wild-type littermates (Zhou et al.,
2000
). In the present work, we have explored further the
involvement of metalloproteinases in tubulogenesis in 3D fibrin gels. Our data
lend support to the idea that MT-MMPs play an essential role during
tubulogenesis within a fibrin matrix. These results define a subgroup of
MT-MMPs as crucial targets for the development of selective MMP inhibitors
that target potentially redundant `angiogenic MMPs' while sparing MMPs
involved in other normal physiological processes.
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Materials and Methods |
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Angiogenic factors used included: VEGF (Chemicon International Inc.,
Temucula, CA), FGF-2 (R&D Systems, Abingdon, UK), TNF- (Chemicon
International, Inc.), transforming growth factor-ß1 (TGF-ß1, R&D
Systems), epidermal growth factor (EGF, Chemicon International Inc.),
transforming growth factor-
(TGF-
, R&D Systems), hepatocyte
growth factor/scatter factor (HGF/SF, a kind gift from Alba Warn, University
of East Anglia, Norwich UK), interleukin-1
(IL-1
, R&D
Systems) and angiogenin (R&D Systems). Fibrinogen [plasminogen- and
urokinase plasminogen activator (uPA)-depleted] was obtained from Calbiochem
(Beeston, UK), whereas thrombin, pepstatin, E64
[trans-epoxysuccinyl-L-leucylamido (4-guanidino)-butane] and aprotinin were
obtained from Sigma-Aldrich (Poole, UK).
Cell culture
Primary human umbilical vein endothelial cells (HUVECs) from pooled donors
and primary human dermal microvascular endothelial cells (HDMEC) were
purchased from TCS Biologicals (Buckingham, UK) and grown on type I collagen
(Sigma-Aldrich) coated flasks (60 µg/ml), in medium supplied by the
manufacturer [which included 2% volume/volume (v/v) foetal bovine serum
(FBS)]. All experiments were performed on cells between passage number one and
five.
U87 human glioma cells, U251N human glioma cells, A10 smooth muscle cells, were obtained from the American Tissue Culture Collection (ATCC, Manassas, USA). MDCK epithelial cells where obtained from Morag Park, McGill University, Montreal, Canada. These cells were maintained in Dulbecco's modified Eagle's medium (DMEM-F12, Gibco BRL) with 10% (v/v) heat-inactivated FBS (Gibco BRL), 2 mM L-glutamine (Gibco BRL), 1x non-essential amino acids (Gibco BRL), and 1 mM sodium pyruvate (Sigma-Aldrich).
In vitro angiogenesis assay
Twenty-four-well format
HUVECs or HDMEC were embedded within fibrin gels at a concentration of
1.5x106 cells/ml. Cells (4.5x105) were
centrifuged gently at 170 g for 10 minutes in 1.5 ml eppendorf
tubes and the cell pellets resuspended in 300 µl of a mixture of 2.5 mg/ml
plasminogen- and uPA-free human fibrinogen (made in serum-free medium).
Thrombin (0.5 U/ml) was then added to the fibrinogen mixture and quickly
pipetted into the wells of a 24-well plate (covering the entire surface of the
wells) and allowed to clot at 37°C for 30 minutes. Serum-containing EC
medium was then added to the wells with angiogenic factors to induce tube
formation. Protease inhibitors and TIMPs were also added to the gels (prior to
polymerisation) and to the culture medium where indicated. Fibrin gels were
incubated for several days at 37°C and 5% (v/v) CO2. Images of
tubular structures were taken after 3 days, using a JVC TK-S340 video camera
attached to a Nikon microscope. Five different fields were evaluated for each
treatment, each image being selected on the basis of the optimal focal plane
that had the majority of cells in focus. The tubular structures were traced,
and the total length was analysed using LUCIA G/Comet software.
Six-well format
A six-well format was used to study the effects of co-culturing EC with
other cell types on EC tube formation within a 3D matrix. In this system, two
fibrin gels (containing two different cell types) were placed within the same
well of a six-well dish, covered with the same culture medium, but with no
direct physical contact. Fibrin gels were performed as above with
1.5x106 cells/ml (HUVECs) within 300 µl gels. Gels were
pipetted as a drop culture in the bottom of a six-well plate (not covering the
entire surface of the well). Similarly, an equivalent number of another cell
type (i.e. U87 cells or MDCK cells) was also embedded in the same way within a
separate fibrin gel and placed next to the EC (but without physical contact).
The gels were allowed to clot at 37°C for 30 minutes. The wells were then
flooded with serum-containing HUVEC medium and incubated at 37°C and 5%
(v/v) CO2 for several days. One half of the culture medium was
replaced with fresh serum-containing HUVEC medium every second day. Images of
the tubular structures were analysed as described above.
Preparation of fibrin gels for transmission electron microscopy
(TEM)
After 72 hours in culture, HUVECs within fibrin gels were fixed overnight
in 2.5% weight/volume (w/v) glutaraldehyde in 0.1 M cacodylate buffer pH 7.2
at room temperature. After fixation, the gels were placed in 0.1% (w/v) osmium
tetroxide for 1 hour at room temperature, after which they were washed with
distilled water and dehydrated using a series of graded ethanol dilutions.
Embedding involved transfer of the gels from 100% ethanol into a 50%/50% (v/w)
resin/ethanol mix for 1 hour after which they were immersed in 100% LR resin
(Agar Scientific, Essex, UK) overnight at room temperature. The gels were then
polymerised at 60°C for 12-18 hours. The embedded gels were trimmed and
sectioned with an LKB Nova ultramicrotome. Sections of 70-100 nm were cut,
mounted onto copper grids and stained with uranyl acetate and lead citrate.
The sections were viewed at an accelerating voltage of 80 kV in a JEOL 2000 EX
electron microscope and suitable images were photographed.
Serial sections of fibrin gels for toluidine blue staining
After 72 hours in culture, HUVECs within fibrin gels were fixed for 1 hour
in 2.5% (w/v) glutaraldehyde in 0.1 M cacodylate buffer pH 7.2. After
fixation, the gels were placed in 0.1% (w/v) osmium tetroxide (in 1%
cacodylate buffer pH 7.2) for 1 hour at room temperature. The gels were then
infiltrated and polymerised in LR resin. Sections of 2 µm were cut, stained
with 1% toluidine blue and photographed.
RNA extraction
Total RNA from HUVECs cultured either as a monolayer or within the fibrin
gels was harvested by solubilising the gels with RNazol B (Biogenesis Ltd,
Poole, UK), and the RNA extracted as per the manufacturer's instructions. RNA
was then resuspended in H2O and stored at -70°C. The quality
and quantity of the RNA was established by reading the optical density (OD) of
each sample at 260 nm and 280 nm using a Cecil CE2041 Spectophotometer (2000
series).
Taq Man real-time reverse transcription-polymerase chain reaction
(RT-PCR)
RT reactions contained 1 µg of total RNA, 1xPCR buffer (10 mM
Tris-HCl pH 9.0, 50 mM KCl and 1.5 mM MgCl2) (Gibco BRL), 1 mM each
deoxynucleotide triphosphates (dATP, dGTP, dCTP, and dTTP), 20 units placental
ribonuclease inhibitor (RNAguard, Amersham), 100 pmol of random hexamer
oligodeoxynucleotides and 200 units of reverse transcriptase (Superscript II,
Gibco BRL). The final reaction volume was 20 µl. Each reaction was
pre-incubated at 20°C for 10 minutes and the room temperature reaction was
performed at 42°C for 50 minutes. Each sample was then heated to 95°C
for 5 minutes to terminate the room temperature reaction and then cooled to
4°C and samples stored at -20°C.
For the PCR reactions, standard curves were prepared for both the target gene (MMP) and the endogenous control (rRNA) by amplifying serial dilutions (in triplicate) of one of the samples known to contain the mRNA of interest. The quantity of the experimental samples for the MMP was then determined from the standard curve and divided by the quantity of the endogenous control (rRNA). The quantities of the MMP samples were thus expressed as an n-fold difference relative to the endogenous control.
Each PCR reaction contained: 1x TaqMan buffer A, 5.5 mM MgCl2, 0.05% (w/v) gelatin (Sigma-Aldrich), 200 µM dATP, 200 µM dCTP, 200 µM dGTP, 400 µM dUTP, 100 nM probe (for the MMP or rRNA), 200 nM forward primer (MMP or rRNA), 200 nM reverse primer (MMP or rRNA), 0.01 U/µl AmpErase UNG, 0.05 U/µl AmpliTaq Gold, 5 ng of reverse transcribed RNA and H2O for a total reaction volume of 25 µl. PCR reactions were carried out in microAmp optical 96-well plates in a Applied Biosystems ABI PRISM 7700 Sequence Detection System. The rRNA probe/primer set and all PCR reagents (except where indicated) were purchased from Applied Biosystems (Warrington, UK). The sequences for the human MMP probes and primers were designed using Primer Express 1.0 (Applied Biosystems, Warrington, UK), and these sequences are copyright of Applied Biosystems.
Cell lysate isolation
Conditioned medium from the HUVECs grown within the fibrin gels (3D
culture) was removed, and the gels washed with phosphate-buffered saline
(PBS). 250 µl lysis buffer [10 mM Tris-HCl pH 7.6, 10 mM NaCl, 3 mM
MgCl2, 1% (v/v) nonidet-P40 and 100 µM
phenylmethylsulphonylfluoride (PMSF)] was added to the gels and transferred to
a 1.5 ml eppendorf tube. The mixture was then homogenised using an ultraturrax
homogeniser. The homogenates were kept on ice for 1 hour with periodic mixing,
after which they were centrifuged for 10 minutes at 6000 rpm, 4°C. The
supernatant was collected and stored at -20°C for subsequent analysis.
Cell lysate isolation of 2D cultures of HUVECs was performed as above except
the cells were scraped from the dish with a cell scraper then lysed with 50
µl of lysis buffer.
Gelatin zymography
Gelatin zymography was performed using a 5% polyacrylamide stacking gel and
a 10% polyacrylamide resolving gel co-polymerised with 1 mg/ml gelatin
(Sigma-Aldrich). Equal amounts of sample were mixed with sodium dodecyl
sulphate (SDS) sample buffer [final concentration: 50 mM Tris-HCl pH 6.8, 1%
(w/v) SDS, 0.025% (w/v) bromophenol blue, and 10% (v/v) glycerol] under
non-reducing conditions and loaded onto the gel. After electrophoresis, the
gels were washed in 50 mM Tris-HCl (pH 8.0), 5 mM CaCl2 and 2.5%
(v/v) Triton X-100 overnight and then incubated in 50 mM Tris-HCl (pH 7.5) and
5 mM CaCl2 for 24 hours at 37°C. Gels were stained with 2.5
mg/ml Coomassie Brilliant Blue R-250 in 10% (v/v) acetic acid and 10% (v/v)
isopropanol, then destained in 10% (v/v) acetic acid and 10% (v/v)
isopropanol. Gelatinolytic activity appeared as a clear band on a blue
background.
Western analysis
Equivalent amounts of total cellular protein from each sample were prepared
in SDS sample buffer with 100 mM dithiothreitol (DTT) and boiled for 5 minutes
prior to loading on the gel. Protein samples were separated using a 5%
polyacrylamide stacking gel and a 10% polyacrylamide resolving gel. After
electrophoresis, proteins were transferred to a polyvinylidene fluoride (PVDF)
membrane (NEN Life Science Products Inc. Boston, MA) in 10 mM
3-(cyclohexylamino)-1-propanesulphonic acid (CAPS) buffer pH 11 with 10% (v/v)
methanol using a BioRad semi-dry blotting apparatus at 250 mA for 30 minutes.
Membranes were left to air dry, then blocked in 5% (w/v) skimmed milk powder
in 0.1% (v/v) PBS-Tween-20 for 1 hour at room temperature. The membranes were
then probed with a sheep anti-human MT1-MMP polyclonal antibody (N175, 5
µg/ml) overnight at 4°C. The next day, the membranes were washed
2x15 minutes and 5x5 minutes in 0.1% (v/v) PBS-Tween-20. Membranes
were then probed with a horseradish peroxidase (HRP)-conjugated donkey
anti-sheep secondary antibody for 1 hour at room temperature. Membranes were
washed again for 2x15 minutes and 5x5 minutes in 0.1% (v/v)
TBS-Tween-20. Detection of the secondary antibody was performed using the
ECL+plus system as per the manufacturer's instructions (Amersham, Little
Chalfont, UK). Membranes were exposed to ECL hyperfilm (Amersham) and
developed using a Xograph Imaging Systems Compact X4 automatic developer.
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Results |
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Angiogenic factors and glioma-derived factors both increased
endothelial tubulogenesis
Endothelial tubulogenesis within fibrin gels was found to be dependent on
the presence of either angiogenic factors or tumour cells since HUVECs and
HDMECs grown within fibrin gels in standard serum-containing EC growth medium
with no exogenous added growth factors produced very few tubular structures.
Individual angiogenic factors and an angiogenic cocktail (containing VEGF,
FGF-2, EGF, TNF-, TGF-
, TGF-ß1, HGF/SF, IL-1
and
angiogenin) were used to evaluate their effect on HUVEC tube formation within
fibrin gels. Fig. 2A
illustrates that the angiogenic cocktail efficiently and significantly induced
tubulogenesis in HUVECs, demonstrating the synergistic action of these
angiogenic factors. Significant increases in tube formation were also seen
with VEGF, FGF-2 and HGF/SF on their own
(Fig. 2A). None of the other
angiogenic factors tested on their own were capable of increasing
tubulogenesis in this system. VEGF and FGF-2 also demonstrated additive
effects when added in combination (Fig.
2A). Similarly, we tested a smaller panel of angiogenic factors on
HDMEC tubulogenesis and found that both VEGF and FGF-2 could induce tube
formation within the fibrin gels (data not shown).
|
Since angiogenic factors were able to increase EC tubulogenesis within
fibrin gels, we looked at the effect of co-culturing glioma cells with HUVECs
to see if the former, a rich source of angiogenic factors
(Plate et al., 1992;
Takano et al., 1996
), could
influence HUVEC tubulogenesis. HUVECs were co-cultured with an equivalent
number of either glioma cells or non-tumorigenic cells by growing each cell
type in separate fibrin gels in a single well of a six-well plate. The two
gels were not in contact but were covered with serum-containing medium (with
no exogenous angiogenic factors added). Aprotinin, a serine protease
inhibitor, was also added to these co-cultures in order to prevent gel
degradation by soluble serine proteases secreted by the tumour cells.
Aprotinin, however, did not have any adverse effects on EC tubulogenesis
within the fibrin gels (see below). Soluble factors produced either by the
tumour cells or the non-tumorigenic cells could therefore interact with the
HUVECs and influence their behaviour within the fibrin gels. U87 glioma cells
were able to significantly increase HUVEC tube formation within the fibrin gel
(Fig. 2B) (as well as U251N
glioma cells, data not shown), whereas non-tumorigenic cells, such as MDCK
epithelial cells (Fig. 2B) (and
A10 smooth muscle cells, data not shown), did not increase HUVEC tube
formation. Furthermore, the addition of U87 glioma cells to the co-culture was
dose dependent, as increasing the amount of U87 glioma cells resulted in an
increase in tubulogenesis (Fig.
2B). These results indicated that tumour cells specifically
secreted soluble factors that could influence EC tube formation within a 3D
environment. Since both the angiogenic factors and the tumour-derived factors
increased tubulogenesis, the angiogenic factors VEGF and FGF-2 were added (as
the angiogenic stimulator) to subsequent experiments to simplify the culture
conditions.
HUVEC tubulogenesis is metalloproteinase dependent
We then set out to characterise further the mechanism of HUVEC tube
formation within fibrin gels by looking at the involvement of proteolytic
enzymes during this process. Firstly, four different types of protease
inhibitors were tested, each blocking proteolytic activity against one of the
four classes of proteases (metallo-, serine, cysteine and acidic proteases),
which function in the extracellular environment. The protease inhibitors used
were TIMP-2 (natural MMP inhibitor), aprotinin (serine protease inhibitor),
E64 (cysteine protease inhibitor) and pepstatin (acidic protease inhibitor).
As can be seen in Fig. 3A, the
serine, cysteine and acidic protease inhibitors had no effect on HUVEC tube
formation within fibrin gels. Only TIMP-2 demonstrated a significant
inhibitory effect, suggesting an essential role for MMPs, and not other
classes of proteolytic enzymes, during HUVEC tubulogenesis within fibrin
gels.
|
Establishing that the endogenous MMP inhibitor, TIMP-2, was able to inhibit
HUVEC tube formation within fibrin gels prompted us to look at other TIMPs
(TIMP-1 and -4) to see if these family members would similarly manifest
inhibitory effects in this culture system. TIMP-4 behaved similarly to TIMP-2,
significantly inhibiting HUVEC and HDMEC tube formation (induced by VEGF and
FGF-2) at similar concentrations to TIMP-2
(Fig. 3B for HUVECs and
Fig. 3C for HDMEC) and was
found to be dose dependent (Fig.
3D for HUVECs). TIMP-1, however, had no inhibitory effect on HUVEC
(Fig. 3B) and HDMEC
(Fig. 3C) tube formation at the
same concentration used for TIMP-2 and TIMP-4. TIMP-2 also blocked U87
glioma-induced tubulogenesis (data not shown). The fact that TIMP-2 and -4,
but not TIMP-1, blocked HUVEC and HDMEC tubulogenesis within these fibrin gels
suggests that the principal MMP required for allowing these EC to invade and
arrange into tubular structures may be a member of the MT-MMP family (as
TIMP-1 is a very poor inhibitor of several MT-MMP family members)
(Butler et al., 1997;
Llano et al., 1999
;
Matsumoto et al., 1997
;
Shimada et al., 1999
;
Will et al., 1996
). HUVEC
viability and proliferation were not negatively effected by TIMP-2 and TIMP-4
(data not shown), demonstrating that the inhibitory actions of the TIMPs are
primarily caused by their MMP inhibitory functions.
Mutant N-TIMP-2 proteins show selective effects on HUVEC
tubulogenesis
Previously, the mechanism of inhibition of MMPs by TIMP-2 has been explored
through the generation of a panel of mutant TIMP-2 N-terminal domain
(N-TIMP-2) proteins (Butler et al.,
1999). The N-terminal domain of TIMP-2 is necessary and sufficient
for MMP inhibition, although loss of the C-terminal domain results in a lower
association rate constant with MMPs than the full-length wild-type TIMP-2 has
(Butler et al., 1999
;
Willenbrock et al., 1993
).
Several N-terminal TIMP-2 proteins carrying point mutations in defined
residues have been generated, and the interactions of these proteins with
purified MMPs have been dissected kinetically
(Butler et al., 1999
). In
particular, one mutant of notable importance is the Y36G mutant. The Y36G
mutant was generated by substituting a Gly for Tyr36 in the flexible AB loop
of TIMP-2, thus generating a net negative charge in this region. This
substitution resulted in an elevation of the apparent inhibition constant
(Kiapp) for MT1-MMP while maintaining similar
Kiapp to the wild-type N-TIMP-2 for all other MMPs
tested (Butler et al., 1999
;
Williamson et al., 2001
).
Tyr36 interacts directly with the MT-loop, which is an eight-residue insertion
between strands II and III only found in MT1-, MT2- and MT3-MMP
(Butler et al., 1999
). This
mutant therefore is a very poor inhibitor of MT1-MMP, while still maintaining
good inhibitory action against most other MMPs.
These N-TIMP-2 proteins (wild-type and mutant) were used as tools to
characterise the possible involvement of MT1-MMP in the tubulogenesis process.
HUVECs were thus embedded within fibrin gels and overlaid with standard
serum-containing EC medium with VEGF and FGF-2. The mutant and wild-type
N-TIMP-2 as well as the wild-type full-length TIMP-2 were added at the
concentrations indicated in Fig.
3, both within the gel and in the culture medium.
Fig. 3E shows that the
full-length wild-type TIMP-2 was more efficient at blocking HUVEC tube
formation than the N-terminal wild-type TIMP-2, which corresponded with
previously reported kinetic data (Butler et
al., 1999; Willenbrock et al.,
1993
). The Y36G N-TIMP-2 mutant, which was reported to be a very
poor inhibitor of MT1-MMP but not of other MMPs tested, could not suppress
HUVEC tubulogenesis within fibrin gels at the same concentration as the
wild-type N-TIMP-2 (Fig. 3E).
Therefore, these data further support the hypothesis that MT1-MMP, or an MMP
with a similar TIMP sensitivity, is the principal enzyme involved during HUVEC
tube formation within fibrin gels.
The C-terminal haemopexin domain of MMP-2 (otherwise known as PEX) inhibits
angiogenesis in several model systems
(Brooks et al., 1998;
Pfeifer et al., 2000
). This
may occur through competition for binding sites for MMP-2 on the cell surface
(via the integrin
vß3 or the MT1-MMP/TIMP-2
complex), thereby blocking pro-MMP-2 activation and localisation on the cell
surface (Brooks et al., 1998
).
The recombinant C-terminal haemopexin domain (up to 5 µM) had no effect on
HUVEC tube formation in fibrin gels (data not shown). Furthermore, a
monoclonal antibody raised against the human MMP-2 haemopexin domain (VB3),
which prevents MT1-MMP-mediated pro-MMP-2 activation in cell-based assays
(V.K., L. Bailey, M., Patterson, G. M., unpublished) also failed to block
HUVEC tube formation (data not shown). Together these observations suggest
that active MMP-2 is not essential in this process; an observation is
consistent with the insensitivity of tubulogenesis to inhibition by
TIMP-1.
MMP expression during tubulogenesis
We investigated the expression of the MT-MMPs and additional MMPs in HUVECs
undergoing tubulogenesis within fibrin gels compared to cells in conventional
2D cultures using quantitative real-time TaqMan RT-PCR
(Fig. 4). Expression of MT1-,
MT2- and MT3-MMPs increased in HUVECs during culture within the fibrin gels,
whereas cells in 2D culture showed no appreciable changes in expression of any
of the MMPs that we examined. In some experiments, increases in MT4- and
MT5-MMP mRNA levels were also observed in the fibrin gel cultures, but this
was not seen consistently. MT6-MMP expression was very low (>35 threshold
cycles, which is the standard detection limit that we have employed in TaqMan
quantitative RT-PCR assays) and did not vary during culture. MMP-2 (but not
MMP-19) was also found to be upregulated in HUVECs cultured within the fibrin
gels. Although MMP-19 is also poorly inhibited by TIMP-1
(Stracke et al., 2000), its
expression profile is inconsistent with a major role in HUVEC tubulogenesis.
Therefore, several MT-MMPs and MMP-2 are upregulated in the 3D fibrin gel
model, consistent with the possible collaboration of multiple MMPs in in vivo
angiogenesis.
|
As shown by zymography in Fig. 5A, the levels of cell-associated MMP-2 protein increased as the cells were cultured in the 3D fibrin matrix but not in 2D monolayers, which agrees with the mRNA data. Moreover, consistent with the increased activity of MT-MMPs, the amount of active MMP-2 also increased in the 3D fibrin gel cultures. However, steady-state protein levels of MT1-MMP did not appear to increase in 2D or 3D cultures as demonstrated by western blot analysis of whole cell lysates (Fig. 5B).
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Discussion |
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We demonstrated that although unstimulated HUVECs and HDMEC could undergo
tubulogenesis within fibrin gels, tube formation was significantly increased
with the addition of angiogenic factors such as VEGF, FGF-2 and HGF/SF.
Additive effects were also observed during VEGF and FGF-2 co-stimulation, and
synergistic effects were observed when treated with an angiogenic cocktail of
nine known angiogenic factors. Soluble factors derived from glioma cells could
also significantly increase HUVEC tube formation in a dose-dependent fashion,
which is similar to the angiogenic factors, although factors derived from
non-tumorigenic cell types could not. It is very likely that angiogenic
factors produced by the glioma cells were responsible for influencing HUVEC
tubulogenesis, as glioma cells secrete high levels of angiogenic factors such
as VEGF, FGF-2 and EGF (Plate et al.,
1992; Rooprai et al.,
2000
; Shweiki et al.,
1992
), although the precise nature of these glioma-derived
secreted factors is unknown. In our culture system, both macrovascular EC
(HUVECs) and microvascular EC (HDMEC) behaved similarly in terms of tube
formation in fibrin gels and MMP production (data not shown), which
demonstrates that although EC sprouting occurs primarily from the
microvasculature in vivo (Folkman and D'
Amore, 1996
), macrovascular EC can also undergo an angiogenic
phenotype if given the proper cues and growth conditions.
Since HUVECs and HDMEC used in these studies were embedded within an
insoluble 3D matrix, proteolytic activity would be a prerequisite in order for
the cells to invade, migrate and organise into a complex network of
interconnecting structures. A series of metallo-, serine, cysteine and acidic
protease inhibitors were therefore tested for their ability to alter EC
tubulogenesis. MMP inhibitors were the only class of inhibitors that
significantly blocked EC tube formation and their effect was dose dependent.
This was found for recombinant TIMP-2 and -4 but not recombinant TIMP-1. A
high concentration of recombinant TIMPs (10 µg/ml) was required to
achieve inhibition of tubulogenesis [occurring in 2% (v/v) serum-containing
medium], which has also been observed by others
(Anand-Apte et al., 1997
;
Hiraoka et al., 1998
). This
may have been caused by the high levels of MMPs in the serum, which may act as
a sink to limit the amount of free TIMP available to inhibit cellular MMPs.
Also, some MMPs (i.e. MT1-MMP and MMP-2) are localised to invadopodia during
cellular invasion (Nakahara et al.,
1997
). It is possible that MMPs in these specialised structures
might require a high TIMP concentration for inhibition owing to the high local
MMP concentration or to steric effects that limit TIMP access to these
regions. We also demonstrated that both TIMP-2 and TIMP-4 did not inhibit
HUVEC tube formation via growth inhibitory effects. Therefore, we conclude
that the main inhibitory action of TIMP-2 and TIMP-4 in this process was
likely to be caused by the prevention of proteolytic activity, thereby causing
HUVECs and HDMEC to remain stationary within the fibrin matrix. However,
alternative roles for the MT-MMPs in this process cannot be ruled out, such as
the liberation of cryptic integrin-binding sites within the fibrin molecule,
shedding of cell-surface EC growth factors or the degradation of inhibitory
proteins such as IGF binding proteins, which have been previously reported for
other MMPs (d'Ortho et al.,
1997
; Giannelli et al.,
1999
; Wu et al.,
1999
; Xu et al.,
2001
).
Although the uPA/plasmin system is involved in the degradation of fibrin
(van Hinsbergh et al., 1997),
this enzymatic cascade did not appear to play a role during HUVEC
tubulogenesis within fibrin gels owing to its insensitivity to the serine
protease inhibitor aprotinin, which inhibits uPA/plasmin
(Pepper et al., 1996
). Other
reports have demonstrated that EC from muscle explants generated from
plasminogen-/- or uPA-/-/tPA-/- mice embedded
within a fibrin matrix successfully sprouted and formed tubular structures
within the 3D matrix (Hiraoka et al.,
1998
). Natural and synthetic MMP inhibitors (TIMP-2 and BB94) were
able to block EC sprouting into the surrounding matrix but not that of
non-endothelial mesenchymal cells (Hiraoka
et al., 1998
). The fact that TIMP-2 and -4, but not TIMP-1,
inhibited HUVEC and HDMEC tube formation within fibrin gels suggests that
members of the MT-MMP family (specifically MT1-, MT2-, MT3- and MT5- MMP)
might be the main MMPs involved in this process, since these MT-MMP members
are very poorly inhibited by TIMP-1 (Butler
et al., 1997
; Llano et al.,
1999
; Matsumoto et al.,
1997
; Shimada et al.,
1999
; Will et al.,
1996
). Further support for this suggestion comes from our data
using the mutant N-terminal TIMP-2 proteins, which indicate that a target that
is not inhibited by Y36G N-TIMP-2 is involved. This result is highly
suggestive of MT1-MMP (Butler et al.,
1999
; Williamson et al.,
2001
). However, as the Kiapp values of Y36G
N-TIMP-2 for all MMPs are not known at present, we cannot conclusively state
that EC tubulogenesis is strictly MT1-MMP dependent. Furthermore, it is clear
that expression of other TIMP-1-insensitive MMPs (MT2-MMP and MT3-MMP) is
upregulated in our 3D fibrin cultures, suggesting they may also participate.
The lack of increase in MMP-19 expression in HUVECs in 3D fibrin cultures
argues against its involvement, and it is also clear that tubulogenesis does
not depend upon active MMP-2 (even though when undergoing tubulogenesis, MMP-2
mRNA and protein levels are increased and activated MMP-2 is present) on the
basis of its insensitivity to TIMP-1, recombinant PEX and a function-blocking
antibody to MMP-2. Using membrane preparations, we showed previously that
MT1-MMP was the principal activity required for pro-MMP-2 activation in
HUVECs, on the basis of the ability of specific anti-MT1-MMP antibodies to
block this process (Lafleur et al.,
2001
). There is certainly increased activation of pro-MMP-2
occurring during 3D tubulogenesis in fibrin gels, so it was somewhat
surprising that levels of MT1-MMP protein did not rise in parallel with the
mRNA abundance. However, it is possible that MT1-MMP turnover via
internalisation and degradation increases during tubule formation, such that
steady-state levels remain relatively constant. We can nevertheless not rule
out the possibility that other MT-MMPs are responsible for fibrin degradation
and pro-MMP-2 activation in HUVEC undergoing tubulogenesis.
In other studies, EC and epithelial cell sprouting have been suggested to
be dependent on the action of membrane-anchored MT1-MMP, as MT1-MMP was
demonstrated to possess potent fibrinolytic activity and transfection of
non-invasive MDCK cells with MT1-MMP enhanced invasion and a tubulogenic
response when cultured on fibrin gels
(Hiraoka et al., 1998).
Another group has also demonstrated that MT1-MMP was specifically responsible
(using antisense oligonucleotides) for the formation of HGF/SF-induced
branching and tubulogenesis in MDCK epithelial cells
(Kadono et al., 1998
).
Similarly, a capillary-like structure formation in 3D type I collagen gels of
human microvascular EC was delayed when pre-treated with anti-MT1-MMP
antibodies (Chan et al., 1998
).
Finally, Gálvez et al. have generated function-perturbing anti-MT1-MMP
monoclonal antibodies and have demonstrated that these antibodies could
inhibit phorbol 12-myristate 13-acetate (PMA)-induced EC migration and
invasion of collagen and fibrin gels and EC tubulogenesis when seeded on
Matrigel (Galvez et al.,
2001
). However, although these antibodies inhibit EC migration,
invasion and tubulogenesis, this inhibition was partial, which may reflect the
ability of other members of the MT-MMP family to compensate for the lack of
MT1-MMP activity. Although these groups have specifically linked MT1-MMP in
either EC or kidney epithelial tubulogenesis, we have not been able to
completely block MT1-MMP activity in our culture system. Attempts have been
made with several polyclonal and monoclonal anti-MT1-MMP antibodies, and
although we have previously demonstrated that we could block MT1-MMP activity
using these antibodies in a cell-free system
(Lafleur et al., 2001
), these
antibodies never proved inhibitory in cell-based assays in our hands (data not
shown). This may reflect participation of multiple TIMP-1-insenstive MT-MMPs
in HUVEC tubulogenesis.
Taken together, the results presented suggest that the principal enzymes
involved in HUVEC tubulogenesis within fibrin gels are members of the MT-MMP
subfamily and may be specifically MT1-MMP, MT2-MMP or MT3-MMP. Other groups
(Hiraoka et al., 1998;
Hotary et al., 2000
) have
previously presented data linking MT1-MMP to tubulogenesis in vitro, and our
results are consistent with a significant role for MT1-MMP. However, it is
also clear from our RNA expression data that MT2-MMP and MT3-MMP are also
upregulated during tubule formation, and these enzymes could therefore
contribute to the process. MT1-MMP involvement in angiogenesis in vivo in at
least some tissue types is indicated by studies on MT1-MMP-/- mice,
where defective vascular invasion of cartilage was observed and there was a
lack of an angiogenic response induced by FGF-2 in the corneal micropocket
assay (Zhou et al., 2000
).
However, MT1-MMP cannot be essential for all types of angiogenesis as the
MT1-MMP-knockout would be expected to result in mid-gestational lethality, as
is the case for VEGF-/- or VEGFR-/- mice
(Carmeliet, 2000
). Therefore it
may be the redundancy of the MT-MMP subfamily (or the MMP family in general)
that could be responsible for the more subtle vascular defects seen in
MT1-MMP-null mice. These results have implications for the design of
anti-angiogenic therapies that target the MMPs. Although certain matrix
components will often be encountered by invading EC (such as the basement
membrane or fibrin), the angiogenic response might recruit different MMPs
depending on the tissue type (and therefore ECM composition) being
vascularised. Therefore, perhaps a more efficient method to block the
angiogenic process would be to selectively target the key redundant MMPs
involved during angiogenesis, such as the MT-MMPs, while sparing other MMPs
that may be involved in other normal processes and in the production of
angiogenesis inhibitors such as angiostatin (i.e. MMP-3, MMP-7 and MMP-12)
(Cornelius et al., 1998
;
Lijnen et al., 1998
;
Patterson and Sang, 1997
).
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