Murray J. Clifford. Angiogenesis Protocols - Methods in Molecular Medicine, Vol. 46
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202 Burbridge and West
which will equilibrate at pH 7.4 in atmospheric CO
2
, concentration of 2%. If
conditions of 5% CO
2
are used, pH may be corrected by adding NaHCO
3
to a
final concentration of 25 mM, or by adding NaOH until equilibrium is reached at
pH 7.4, the net result in terms of NaHCO
3
concentration and buffering capacity
being the same [Freshney (22)]. Maintenance of pH at 7.4 is of utmost impor-
tance for optimal growth.
6. Where fibrin gels are used, 300 µg/ml ¡-amino-n-caproic acid should be added to
culture medium to prevent spontaneous fibrinolysis.
7. For optimal contrast and depth of field, reduce the aperture as far as possible.
8. Use a black and white camera for optimal image resolution, especially when
automated image analysis is required.
9. The image analyzer used in our laboratory is the BIOCOM Visiolab™ 2000,
using specialized software developed specifically for analysis of the 3D aortic
ring cultures.
10. The use of Permanox plastic ensures minimal autofluorescence when using fluo-
rescent markers.
11. In view of the large species heterogeneity of von Willebrand factor, antibodies
raised to the human antigen may be unsuccessful, even though species cross-
reactivity is claimed.
12. Cell culture-treated dishes are preferred to ensure reduced adherence of agarose
for easier manipulation.
13. At this stage, agarose may be stored at 4°C for up to three days; any longer, and
agarose tends to dehydrate.
14. Using bacteriological plastic here improves adherence of agarose to the base of
the dish so wells do not become detached during culture.
15. The rat aortic endothelium is exceptionally well retained to the intima, and unlike
that of larger species, the formation of clotted blood will not lead to a stripping
off of ECs if aortas are rinsed within 15 min of removal from the animal.
16. If more than one aorta is used in an experiment, this stage should be completed
for each aorta before moving on to the next.
17. It is not necessary to keep Petri dishes on ice during dissection procedures under
the dissecting microscope. Frequent replenishment of ice-cold medium is suffi-
cient to maintain integrity of the aorta and its cellular components. Total proce-
dure may last up to 4 h with no detrimental effects on the subsequent spontaneous
angiogenic response.
18. The optimal length of aortic rings is found to be 1 mm. Shorter lengths are more
difficult to handle and position correctly, and give rise to a lower level of out-
growth, probably because there is a lower local concentration of growth factors.
Longer lengths, although giving satisfactory outgrowth, would obviously lead to
a lower yield per aorta.
19. While all preceding steps may be performed on the workbench in a culture labo-
ratory, from here on work under a laminar-flow hood.
20. The polymerization of the collagen gel will vary with pH.
3D Model of Angiogenesis 203
21. Prepared collagen solutions may be kept on ice up to 30 min; at room tempera-
ture they will gel rapidly (3 min).
22. As for collagen, a concentration of 1.5 mg/mL is found to be optimal.
23. Take care to work at pH 7.4 for satisfactory gel polymerization.
24. The time required for the analysis of each aortic ring is between 2 and 5 min,
depending on the extent of outgrowth.
25. Care should be taken when using such automated image analysis techniques
because artifacts in the captured image (such as incomplete removal of
fibroadipose tissue, or bubbles, or other defects in the collagen or fibrin gels)
may significantly alter results if not carefully monitored. Quantification is also
highly sensitive to illumination level during image capture.
Acknowledgments
D. C. W. acknowledges the support of the North West Cancer Research
Fund and Servier. Dr. Gordon Tucker and Prof. Ghanem Atassi have contrib-
uted greatly to the development of these techniques.
References
1. Auerbach, R., Auerbach, W., and Polakowski, I. (1991) Assays for angiogenesis:
a review. Pharmacol. Ther. 51, 1–11.
2. Montesano, R. and Orci, L. (1985) Tumor-promoting phorbol esters induce
angiogenesis in vitro. Lab. Invest. 42, 469–477.
3. Madri, J. A., Pratt, B. M., and Tucker, A. M. (1988) Phenotypic modulation
of endothelial cells by transforming growth factor beta depends upon the compo-
sition and organization of the extracellular matrix. J. Cell Biol. 106, 1375–1384.
4. Nicosia, R. F. and Ottinetti, A. (1990) Growth of microvessels in serum-free
matrix culture of rat aorta—A quantitative assay of angiogenesis in vitro. Lab.
Invest. 63, 115–122.
5. Schor, A. M., Ellis, I., and Schor, S. L. (1999) Collagen gel assay for angiogen-
esis: induction of endothelial cell sprouting. In: Methods in Molecular Medicine:
Angiogenesis Protocols (Murray, J. C., ed.), Humana Press Inc., Totowa, NJ,
pp XX.
6. Madri, J. A., Merwin, J. R., Bell, L., Basson, C. T., Kocher, O., Perlmutter, R.,
and Prinz, C. (1992) Interactions of matrix components and soluble factors in
vascular responses to injury. In: Endothelial Cell Dysfunctions, (Simionescu, N.
and Simionescu, M., eds.), Plenum Press, New York, pp11–30.
7. Pröls, F., Loser, B., and Marx, M. (1998) Differential expression of osteopontin,
PC4, and CEC5, a novel mRNA species, during in vitro angiogenesis. Exp. Cell
Res. 239, 1–10.
8. Montesano, R., Pepper, M. S., and Orci, L. (1990) Angiogenesis in vitro: mor-
phogenetic and invasive properties of endothelial cells. News. Physiol. Sci. 5,
75–79.
204 Burbridge and West
9. Knedler, A. and Ham, R. G. (1987) Optimized medium for clonal growth of human
microvascular endothelial cells with minimal serum. In Vitro Cell Dev. Biol. 23,
481–491.
10. Nicosia, R. F., Bonanno, F., and Villaschi, S. (1992) Large-vessel endothelium
switches to a microvascular phenotype during angiogenesis in collagen gel cul-
ture of rat aorta. Artherosclerosis 95, 191–199.
11. Nicosia, R. F. and Bonanno, F. (1991) Inhibition of angiogenesis in vitro by
arg-gly-asp-containing synthetic peptide. Am. J. Pathol. 138, 829–833.
12. Nicosia, R. F., Nicosia, S. V., and Smith, M. (1994) Vascular endothelial growth
factor, platelet-derived growth factor and insulin-like growth factor-1 promote rat
aortic angiogenesis in vitro. Am. J. Pathol. 145, 1023–1029.
13. Nicosia, R. F., Lin, Y. J., Hazelton, D., and Qian, X. H. (1997) Endogenous regu-
lation of angiogenesis in the rat aorta model—role of vascular endothelial growth
factor. Am. J. Pathol. 151, 1379–1386.
14. Derringer, K. A. and Linden, R. W. A. (1998) Enhanced angiogenesis induced by
diffusible angiogenic growth factors released from human dental pulp explants of
orthodontically moved teeth. Eur. J. Orthod. 20, 357–367.
15. Wakabayashi, T., Kageyama, R., Naruse, N., Tsukahara, N., Funahashi, Y., Kitoh, K.,
and Watanabe, Y. (1997) Borrelidin is an angiogenesis inhibitor; Disruption of
angiogenic capillary vessels in a rat aorta matrix culture model. J. Antibiot. 50,
671–676.
16. Bocci, G., Danesi, R., Benelli, U., Innocenti, F., Di Paolo, A., Fogli, S., and
Del Tacca, M. (1998) Inhibitory effect of suramin in rat models of angiogenesis in
vitro and in vivo. Cancer Chemother. Pharmacol. 43, 205–212.
17. Chen, C. H., Cartwright, J., Li, Z., Lou, S., Nguyen, H. H., Gotto, A. M., and
Henry, P. D. (1997) Inhibitory effects of hypercholesterolemia and Ox-LDL on
angiogenesis-like endothelial growth in rabbit aortic explants—Essential role of
basic fibroblast growth factor. Arterioscler. Thromb. Vasc. Biol. 17, 1303–1312.
18. Burbridge, M. F., West, D. C., Atassi, G., and Tucker, G. C. (1999) The effect of
extracellular pH on angiogenesis in vitro. Angiogenesis 3, 281–288.
19. Burbridge, M. F. (2000) The rat aortic ring model of angiogenesis in vitro as an
assay for angiogenic modulators. Ph.D. Thesis, University of Liverpool,
Liverpool, UK.
20. Brown, K. J., Maynes, S. F., Bezos, A., Maguire, D. J., Ford, M. D., and Parish, C. R.
(1996) A novel in vitro assay for human angiogenesis. Lab Invest. 75, 539–555.
21. Antes, L. M., Villar, M. M., Decker, S., Nicosia, R. F., and Kujubu, D. A. (1998)
A serum-free in vitro model of renal microvessel development. Am. J. Physiol.
274, F1150–F1160.
22. Freshney, R. I. (1987) In: Culture of Animal Cells. A Manual of Basic Technique.
Alan R. Liss, Inc., New York.
In Vitro Matrigel Assay 205
205
From:
Methods in Molecular Medicine, Vol. 46: Angiogenesis Protocols
Edited by: J. C. Murray © Humana Press Inc., Totowa, NJ
14
In Vitro Matrigel Angiogenesis Assays
M. Lourdes Ponce
1. Introduction
A variety of in vivo and in vitro methods have been used to study angiogen-
esis, the process of blood vessel formation. Two widely accepted but techni-
cally difficult assays include the cornea implant assay and the chick
chorioallantoic membrane assay. The cornea assay requires special equipment
and a skilled person to implant beads containing the test compound in the eyes
of animals; only a small number of samples can be tested due to cost and time.
The chorioallantoic membrane assay requires a large number of samples on
account of the variability of the system and its difficulty in quantitation. In our
laboratory, we have developed a quick and highly reliable method for testing
numerous compounds for angiogenic and/or antiangiogenic activity. The
method is based on the differentiation of ECs on a basement membrane matrix,
Matrigel, derived from the Engelbreth-Holm-Swarm tumor (1). ECs from
human umbilical cords as well as from other sources differentiate and form
capillary-like structures on Matrigel in the presence of 10% bovine calf serum
(BCS) and 0.1 mg/mL of endothelial cell growth supplement (ECGS) (2),
which is a mixture of both acidic and basic fibroblast growth factor (Fig. 1,
Panel C). The formation of tube-like vessels under these conditions can be
used to assess compounds that either inhibit or stimulate angiogenesis. In
the assay, substances that affect angiogenesis, such as the laminin-1 peptide
containing the IKVAV sequence, disturb the formation of capillary-like struc-
tures and the distinctive morphological cell characteristics resulting are indicative
of the potential activity of the compound (Fig. 1, Panel D). However, it can not
be determined from the morphological characteristics whether a compound is
angiogenic or antiangiogenic.
206 Ponce
Tubes do not form when both serum and ECGS concentrations are reduced
(Fig. 1, Panel A). This allows one to determine whether a substance is angio-
genic. Under these circumstances human umbilical vein endothelial cells
(HUVEC) do not completely differentiate; instead, they form incomplete, short
tube-like structures. In the presence of angiogenic compounds, such as fibro-
blast growth factor or hepatocyte growth factor, the ECs differentiate into well-
defined tube-like structures (Fig. 1 Panel B). An additional advantage of these
assays is that they can be scaled down and 48- or even 96-well plates can be
used if many compounds need to be tested or their quantities are limited. The
use of either one or both of these assays can help to quickly and efficiently
identify compounds with angiogenic or antiangiogenic activity. Using both of
these assays, a number of new compounds that affect angiogenesis have been
identified including hepatocyte growth factor (HGF), haptoglobin, estrogen,
IP-10, and numerous laminin-1 peptides among others (3–6). It should be noted
that the activity in all cases should be confirmed by an additional assay.
Fig. 1. Endothelial cells on Matrigel. HUVEC were plated on Matrigel in the pres-
ence of HUVEC media containing either 5% (A,B) or 10% BCS (C,D). Cells incu-
bated with 25 ng/mL of hepatocyte growth factor (HGF) formed tubes in serum
reduced media (B) as compared to their control (A). Cells that were plated with
0.1 mg/mL of a laminin-1 peptide that contains the IKVAV sequence did not form
complete tube-like structures (D) as compared to their negative control devoid of pep-
tide but containing 10% BCS (C). Original magnification ×5.
In Vitro Matrigel Assay 207
2. Materials
1. Matrigel (Sigma Chemical Co., Saint Louis, MO or Becton Dickinson, Inc., Ru-
therford, NJ).
2. 24-Well tissue-culture plates.
3. Pipetmen and sterile tips.
4. Human umbilical vein endothelial cells (HUVEC) prepared from umbilical veins
or commercially obtained from Clonetics Corporation, San Diego, CA (see Note 1).
5. RPMI 1640 media (1 L, Life Technologies, Gaithersburg, MD).
6. Table centrifuge.
7. 50-mL Conical tubes.
8. Hematocytometer.
9. Trypsin-EDTA solution containing 0.05% trypsin, 0.53 mM ethylenediamino-
tetraacetic acid in Hanks’ Balanced Solution (Life Technologies, Gaithersburg, MD).
10. Incubator with 95% air/5% CO
2
mixture.
11. HUVEC medium:
• 500 mL RPMI 1640
• 100 mL of bovine calf serum (BCS), defined and supplemented (HyClone
Laboratories Inc., Logan, Utah)
• 0.1 g ECGS (Collaborative Biomedical Products, Bedford, MA)
• 2,500 U of heparin sodium (Fisher Scientific, Fair Lawn, NJ)
•5 mL of fungizone solution (Life Technologies, Gaithersburg, MD)
• 1.25 mg gentamicin (Life Technologies, Gaithersburg, MD)
•5mL of penicillin/streptomycin solution (10,000 U/mL of penicillin G sodium
and 10 mg/mL streptomycin sulfate in 0.85% saline; Life Technologies,
Gaithersburg, MD).
12. Diff-Quik fixative and solution II (Baxter Scientific Products, McGaw, IL)
3. Methods
3.1. Testing Angiogenic Compounds
1. Allow Matrigel to thaw on ice at 4°C; once thawed keep on ice at all times or it
will solidify at room temperature.
2. Under sterile conditions, coat 24-well plates with 300 µL per well of Matrigel
without introducing air bubbles. Let the plates sit at room temperature for at least
15 min to allow gelling of Matrigel.
3. Rinse a confluent 150-mm dish or equivalent containing 3–4 × 10
6
ECs in 5 mL
of RPMI 1640 medium.
4. Add 5 mL of trypsin-EDTA solution and incubate until the cells detach from the
plate (approx 5 min).
5. Place the cells in a 50 mL conical sterile tube and add 5 mL of HUVEC media to
neutralize the trypsin.
6. Centrifuge the tube on a table top centrifuge for 5 min at 50g; remove the super-
natant, tap the pellet gently and resuspend the pellet in 2 mL HUVEC medium.
7. Set aside a 0.2 mL cell aliquot as control.
208 Ponce
8. Count a 10 µL aliquot in a hemocytometer or use a Coulter Counter to determine
the total number of cells in the tube.
9. Adjust the volume with HUVEC media (containing 20% BCS) in order to have
2 × 10
6
cells per mL (see Note 2).
10. To reduce simultaneously the concentration of serum to 5% and the number of
cells to 0.5 × 10
6
per mL, add an amount of RPMI 1640 media equal to three
times the volume present in the tube.
11. Plate 192 µL of the cell mixture, containing 48,000 cells, into each Matrigel-
coated well.
12. Add the test compound in a total volume of 8 µL (see Note 3).
13. Controls that will contain 10% serum are similarly prepared from the aliquot that
was set aside, except that 1 × 10
6
cells per mL (instead of 2 × 10
6
cells per mL)
are suspended in HUVEC media (20% serum) and are subsequently diluted to
double their volume with RPMI 1640 medium.
14. Gently swirl the dishes to evenly mix the test compound(s) with the cells.
15. Incubate dishes overnight at 37°C in a 5% CO
2
/95% air incubator.
16. The next day gently aspirate the medium from each well, add approx 200 µL of
Diff-Quik fixative per well and incubate for 30 sec.
17. Aspirate the fixative and stain the cells for 2 min with Diff-Quik solution II that
has been diluted 1/1 with water (see Notes 4 and 5).
18. Observe tube structures under a microscope and photograph.
3.2. Screening Compounds That Influence Angiogenesis
This method is a variation of the assay described in Subheading 3.1. and
can be used for screening compounds that either induce or inhibit angiogen-
esis.
1. Follow protocol as in Subheading 3.1. up to step 8.
2. Suspend cells in HUVEC media containing 20% BCS to yield 1 × 10
6
cells per mL.
3. Dilute cell suspension with an equal volume of RPMI 1640 medium lacking
serum to reduce by half the amount of serum and the number of cells per mL.
4. Continue assay as described in Subheading 3.1. Controls should include cells in
medium containing 5 and 10% serum (see Notes 6–8).
4. Notes
1. HUVEC should not be older than 6 passages because they are primary cell cul-
tures and tend to lose their differentiated characteristics. They should always be
handled with gloves since they are of human origin.
2. The amount of HUVEC media in which the cells are suspended in step 9 will
vary from experiment to experiment according to the total number of cells har-
vested from each plate.
3. Several concentrations of each compound should be tested at least in triplicate.
Include 6 wells for controls (3 positive and 3 negative).
In Vitro Matrigel Assay 209
4. Staining of the Matrigel with Diff-Quik solution II for longer than 2 min or with-
out previous dilution will excessively stain the Matrigel making it difficult to
observe the tubular structures.
5. Dishes containing stained cells can be stored wrapped with plastic for a week or
longer at 4°C. Pictures should be taken within a few days since the Matrigel tends
to dry up.
6. To perform assays on 48-well dishes, wells are coated with 200 µL of Matrigel
and 24,000 cells are plated in 150 µL of medium. Ninety-six well plates are coated
with 100 µL of Matrigel and plated with 14,000 cells in 100 µL of medium.
7. Unknown samples should be tested using both serum concentrations.
8. HUVEC can form tubes even in the absence of serum in some commercially
available batches of Matrigel. It is recommended that a control experiment be
carried out using low serum concentrations each time a new Matrigel lot is used.
References
1. Kubota, Y., Kleinman, H. K., Martin, G. R., and Lawley, T. J. (1988) Role of
laminin and basement membrane in the morphological differentiation of human
endothelial cells into capillary-like structures. J. Cell Biol. 107, 1589–1598.
2. Grant, D. S., Kinsella, J. L., Fridman, R., Auerbach, R., Piasecki, B. A., Yamada,
Y., et al. (1992) Interaction of endothelial cells with a laminin A chain peptide
(SIKVAV) in vitro and induction of angiogenic behavior in vivo. J. Cell Physiol.
153, 614–625.
3. Grant, D. S., Kleinman, H. K., Goldberg, I. D., Bhargava, M. M., Nickoloff, B. J.,
Kinsella, J. L., et al. (1993) Scatter factor induces blood vessel formation in vivo.
Proc. Natl. Acad. Sci. USA 90, 1937–1941.
4. Cid, M. C., Grant, D. S., Hoffman, G. S., Auerbach, R., Fauci, A. S., and
Kleinman, H. K. (1993) Identification of haptoglobin as an angiogenic factor in
sera from patients with systemic vasculitis. J. Clin. Invest. 91, 977–985.
5. Ponce, M. L., Nomizu, M., Delgado, M. C., Kuratomi, Y., Hoffman, M. P., Powell, S.,
et al. (1999) Identification of endothelial cell binding sites on the laminin a1 chain.
Circ. Res. 84, 688–694.
6. Malinda, K. M., Nomizu, M., Chung, M., Delgado, M., Kuratomi, Y., Yamada, Y.,
et al. (1999) Identification of laminin _1 and `1 chain peptides active for endot-
helial cell adhesion, tube formation, and aortic sprouting. FASEB J. 13, 53–62.
Microvessel ECs from Adipose Tissue 213
213
From:
Methods in Molecular Medicine, Vol. 46: Angiogenesis Protocols
Edited by: J. C. Murray © Humana Press Inc., Totowa, NJ
15
Microvessel Endothelial Cells
from Human Adipose Tissues
Isolation, Identification, and Culture
Peter W. Hewett
1. Introduction
Human endothelial cells (EC) can be readily prepared from large vessels
such as the aorta, and umbilical (1) and saphenous veins. ECs derived from
these vessels have proven an abundant, convenient, and useful tool for the
investigation of many aspects of endothelial biology. However, the majority of
the body’s endothelium (>95%) forms the microvasculature, and it is at the
level of the blood/tissue interface that many of the phenomena occur that we
wish to investigate. The high degree of functional diversity that the endothe-
lium demonstrates between organs, and within the different vascular beds of a
given organ, is manifest in considerable morphological, biochemical, and
molecular differences (2–6). This apparent heterogeneity has highlighted the
need for reliable methods for microvessel EC isolation and culture from a vari-
ety of tissues of different species in order to establish more realistic in vitro models.
Many techniques have been developed to enrich ECs from tissue
homogenates either directly or after a period in culture, and/or to selectively
promote their proliferation in vitro (6). Most methods use as their starting point
tissue homogenization and digestion, and most are often hampered by low EC
yield and the contamination of cell populations that readily adapt to culture.
Some tissues are inherently better suited to microvessel EC isolation such as
brain and adipose. Wagner and Matthews (7) first utilized adipose from the rat
epididymal fat pad for the isolation of microvessel endothelium. The major
214 Hewett
advantage of this tissue is the difference in buoyant densities between
adipocytes and the stromal component that permits their separation by cen-
trifugation. Similarly, high microvessel density, ease of separation, and the
resistance of the parenchyma to growth in culture have made brain a relatively
accessible source of microvessel endothelium (2,8). Nonetheless the isolation
of ECs from more complex tissues that contain a wide variety of cell types,
many of which readily adapt to cell culture, has until recently remained a sig-
nificant technical problem.
The use of super-paramagnetic beads (Dynabeads™) coupled to endothelial-
specific ligands to select the ECs from mixed cell populations represents a
major advance in the purification of ECs. The original technique described by
Jackson and colleagues utilized the lectin Ulex europaeus agglutinin-1 (UEA-1),
which binds specifically to _-fucosyl residues of glycoproteins on the surface
of endothelial cells (9,10). However, UEA-1 also binds to some epithelia and
mesothelial cells (1,11). We refined this technique by coupling antibodies
raised against platelet EC adhesion molecule (PECAM-1/CD31) (13), a pan-
endothelial marker (4,5), to Dynabeads (PECA-beads), and have used these to
prepare microvessel ECs from a variety of human tissues (12). Dynabeads con-
jugated to antibodies against other endothelial markers, including CD34 and
E-selectin (endothelial cell leukocyte adhesion molecule-1/ELAM-1) (14),
have also been used to isolate microvessel ECs from various tissues.
The identification of new markers such as VE cadherin, ICAM-2, the endothel-
ial restricted tyrosine kinases (Flt-1, Flt-4, KDR/Flk-1, Tie-1 and Tie-2/Tek),
and the increasing number of commercially available antibodies raised against
them have greatly improved the ease of EC characterization (5). Many EC
markers crossreact with subpopulations of hematopoietic cells underlining the
common developmental origins of these cell types. However, due to their lim-
ited viability in vitro these cells do not represent a significant problem for EC
culture. Mesothelial cells are a potentially difficult contaminant of EC cultures
isolated from tissues such as omentum, because they exhibit similar morphol-
ogy and marker expression. However, the absence of PECAM-1 and E-selectin
expression in human mesothelial cells readily distinguishes them from ECs
(11). The use of PECA-beads in principle should eliminate mesothelial cell
contamination of endothelial isolates from sites such as the lung and omentum
that are surrounded by mesothelium (12).
In this chapter we describe the use of super-paramagnetic beads (Dyna-
beads™) coupled to anti-PECAM-1 antibodies (PECA-beads) to isolate
microvessel endothelial cells from human adipose tissue obtained and methods
for the routine culture of these cells. In addition, we briefly discuss some of the
characteristics of these cells and outline the adaptation of this basic technique
to isolate microvessel endothelial cells from human lung and stomach (12,18).