Angiogenesis (2018) 21:425–532
https://doi.org/10.1007/s10456-018-9613-x
REVIEW PAPER
Consensus guidelines for the use and interpretation of angiogenesis
assays
Patrycja Nowak‑Sliwinska1,2 · Kari Alitalo3 · Elizabeth Allen4 · Andrey Anisimov3 · Alfred C. Aplin5 ·
Robert Auerbach6 · Hellmut G. Augustin7,8,9 · David O. Bates10 · Judy R. van Beijnum11 · R. Hugh F. Bender12 ·
Gabriele Bergers13,4 · Andreas Bikfalvi14 · Joyce Bischoff15 · Barbara C. Böck7,8,9 · Peter C. Brooks16 ·
Federico Bussolino17,18 · Bertan Cakir19 · Peter Carmeliet20,21 · Daniel Castranova22 · Anca M. Cimpean23 ·
Ondine Cleaver24 · George Coukos25 · George E. Davis26 · Michele De Palma27 · Anna Dimberg28 ·
Ruud P. M. Dings29 · Valentin Djonov30 · Andrew C. Dudley31,32 · Neil P. Dufton33 · Sarah‑Maria Fendt34,35 ·
Napoleone Ferrara36 · Marcus Fruttiger37 · Dai Fukumura38 · Bart Ghesquière39,40 · Yan Gong19 ·
Robert J. Griffin29 · Adrian L. Harris41 · Christopher C. W. Hughes12 · Nan W. Hultgren12 · M. Luisa Iruela‑Arispe42 ·
Melita Irving25 · Rakesh K. Jain38 · Raghu Kalluri43 · Joanna Kalucka20,21 · Robert S. Kerbel44 · Jan Kitajewski45 ·
Ingeborg Klaassen46 · Hynda K. Kleinmann47 · Pieter Koolwijk48 · Elisabeth Kuczynski44 · Brenda R. Kwak49 ·
Koen Marien50 · Juan M. Melero‑Martin51 · Lance L. Munn38 · Roberto F. Nicosia5,52 · Agnes Noel53 · Jussi Nurro54 ·
Anna‑Karin Olsson55 · Tatiana V. Petrova56 · Kristian Pietras57 · Roberto Pili58 · Jeffrey W. Pollard59 · Mark J. Post60 ·
Paul H. A. Quax61 · Gabriel A. Rabinovich62 · Marius Raica23 · Anna M. Randi33 · Domenico Ribatti63,64 ·
Curzio Ruegg65 · Reinier O. Schlingemann46,48 · Stefan Schulte‑Merker66 · Lois E. H. Smith19 · Jonathan W. Song67,68 ·
Steven A. Stacker69 · Jimmy Stalin66 · Amber N. Stratman22 · Maureen Van de Velde53 · Victor W. M. van Hinsbergh48 ·
Peter B. Vermeulen50,72 · Johannes Waltenberger70 · Brant M. Weinstein22 · Hong Xin36 · Bahar Yetkin‑Arik46 ·
Seppo Yla‑Herttuala54 · Mervin C. Yoder71 · Arjan W. Griffioen11
Published online: 15 May 2018
© The Author(s) 2018
Abstract
The formation of new blood vessels, or angiogenesis, is a complex process that plays important roles in growth and development, tissue and organ regeneration, as well as numerous pathological conditions. Angiogenesis undergoes multiple discrete
steps that can be individually evaluated and quantified by a large number of bioassays. These independent assessments hold
advantages but also have limitations. This article describes in vivo, ex vivo, and in vitro bioassays that are available for the
evaluation of angiogenesis and highlights critical aspects that are relevant for their execution and proper interpretation. As
such, this collaborative work is the first edition of consensus guidelines on angiogenesis bioassays to serve for current and
future reference.
Keywords Angiogenesis · Aortic ring · Endothelial cell migration · Proliferation · Microfluidic · Zebrafish · Chorioallantoic
membrane (CAM) · Vascular network · Intussusceptive angiogenesis · Retinal vasculature · Corneal angiogenesis ·
Hindlimb ischemia · Myocardial angiogenesis · Recombinant proteins · Tip cells · Plug assay · Myocardial angiogenesis ·
Vessel co-option
Table of contents
* Patrycja Nowak-Sliwinska
Patrycja.Nowak-Sliwinska@unige.ch
* Arjan W. Griffioen
aw.griffioen@vumc.nl
Extended author information available on the last page of the article
1. Introduction
2. Endothelial cell and monocyte migration assays
3. Endothelial cell proliferation assays
4. 3D models of vascular morphogenesis
5. Aortic ring assay
6. Tumor microvessel density and histopathological growth patterns
in tumors
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7. Assessment of intussusceptive angiogenesis
8. In vivo sprouting lymphangiogenic assay and AAV-mediated gene
transfer of vascular endothelial growth factor c (VEGFC)
9. Assay for pericyte recruitment to endothelial cell-lined tubes,
capillary assembly, and maturation
10. EC co-culture spheroids
11. Endothelial cell metabolism
12. Endothelial cell precursors
13. Microfluidic assays
14. Flow cytometry and cell sorting assays
15. Loss-of-function approaches in the developing zebrafish
16. Chorioallantoic membrane assays
17. Murine allantois assay
18. In vivo angiogenesis plug assay
19. In vivo vascular network forming assay
20. Developing mouse retinal vasculature—tip cells
21. Corneal angiogenesis assays
22. Mouse oxygen-induced retinopathy model
23. Laser-induced choroidal neovascularization mouse model
24. Transparent window preparations for angiogenesis studies in mice
25. The RIP1-Tag2 transgenic mouse model
26. The MMTV-PyMT breast cancer model
27. Tumor implantation models
28. Mouse hind limb ischemia model
29. Large animal models for myocardial angiogenesis
30. Guidelines for purity of recombinant proteins in angiogenesis
assays
31. Conclusions
Angiogenesis (2018) 21:425–532
angiogenesis, inherently implies accepting specific limitations. It is therefore crucial to understand the full potential
of these bioassays during their specific applications. These
assays have been instrumental in the study of vascular biology in growth and development [6–8] but also play a key
role in the design, development, and evaluation of drugs
that positively or negatively modulate vessel function for
the treatment of many diseases [9–11]. Some examples of
where the use of such bioassays has been imperative are:
(1) the development of angiostatic drugs for the treatment
of cancer, ocular diseases, and other pathologic conditions
where angiogenesis is implicated and also angiogenic treatment strategies in ischemic cardiovascular disease [12, 13],
(2) screening of natural anti-angiogenic compounds [14], (3)
the efforts to design combination therapies including angiogenesis inhibitors [15–20], (4) the unraveling of mechanisms
regulating lymphangiogenesis [21, 22], (5) the interrelationship of angiogenesis and immunity [23–25], (6) the development of imaging as diagnostic strategy [26], (7) the study
of drug resistance mechanisms [27–29], (8) development
of compounds and strategies for the revascularization of
ischemic injuries [30, 31], and (9) to improve the vascular
fitness in aging vessels [32, 33]. The current paper describes
a large collection of assays and techniques for the evaluation of angiogenesis and aims at explaining their respective advantages and limitations. In addition, we included
strategies to study angiogenesis in tissues, through means
of assessing and quantifying microvessel density (MVD),
vessel co-option, pericyte coverage, and tip cell behavior.
1 Introduction
The process of angiogenesis—the formation of new blood
vessels from preexisting ones—is a hallmark of tissue repair,
expansion, and remodeling in physiological processes, such
as wound healing, ovulation, and embryo development, and
in various pathologies including cancer, atherosclerosis,
and chronic inflammation [1–5]. Many of these conditions
share characteristics, for example the occurrence of hypoxia
or inflammation, recruitment of inflammatory cells, angiogenic growth factor production, basement membrane degradation, endothelial cell (EC) migration, proliferation and
differentiation, and modulation of vascular support cells.
However, depending on the tissue or disease under investigation, important details may differ considerably. Moreover, EC in different vascular beds exhibits organ-specific
heterogeneity associated with the differentiated specialized
functions of the tissue. It is often not possible to accurately
visualize the process of angiogenesis and its molecular
players. Therefore, different in vivo, ex vivo, and in vitro
bioassays and techniques have been developed to investigate the specific stages of the angiogenesis. However, the
use of bioassays that study a part of the process, with the
intention to extrapolate and understand the full process of
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2 Endothelial cell and monocyte migration
assays
EC migration is one of the hallmarks of angiogenesis and
one of the earlier steps in the angiogenic cascade. This
process is characterized by cell-autonomous motility property but in some cases, it acquires the features of collective
migration [34–37], in which a group of cells coordinate their
movements toward a chemotactic gradient and by establishing a precise hierarchy with leader and follower cells.
Therefore, dissection of the molecular mechanisms of EC
migration is critical to understand and to therapeutically
manipulate the process to either inhibit sprouting (e.g., in
tumors) or stimulate vessel formation (e.g., during tissue
regeneration or wound healing).
Likewise, migration assays have been successfully used
to assess the migratory responsiveness of monocytes.
Monocytes are actively involved in angiogenesis, and their
migratory response or potential correlates well with that of
endothelial cells. Most importantly, CD14-positive monocytes can easily be isolated and obtained from any individual, not only humans [38, 39], but also mice [40]. A number
Angiogenesis (2018) 21:425–532
of 2D and 3D cellular migration assays have been established as relatively simple in vitro readouts of the migratory/
angiogenic activity of EC in response to exogenous stimuli.
Depending on the specific scientific question, a range of
assays is available to quantitatively and qualitatively assess
EC migration. The most widely employed assays include
variations of the wound closure and the Boyden chamber
assays.
2.1 Types of assays
Cell culture wound closure assay Lateral migration assays
are performed to investigate the pro- or anti-migratory effect
of compounds, as well of specific gene perturbations, or to
describe phenotypes resulting from genetic manipulation
of EC. Although these assays can be used to characterize
chemokinesis (unidirectional migration) in response to a
given compound added to the cell culture medium, they do
not allow determination of directed migration rate toward or
away from a compound. Assessment of chemotaxis can only
be determined when a gradient is also provided.
The cell culture wound closure assay is one of the basic
readouts for characterizing the migratory activity of cells.
It is a measure of the lateral 2D migration of EC in cell culture to test compounds for pro-migratory or anti-migratory
activity. Depending on the migratory effect of the tested
substances, the assay is performed over 2–4 days. ECs are
grown to confluency in a cell culture dish and then scraped
with a razor blade/pipette tip [41], allowing the EC at the
wound edge to migrate into the scraped area. To really examine the motility contribution to the healing and to exclude
the component related to cell proliferation, ECs are incubated with the antimitotic agent mitomycin [42]. Large
genome-wide screens can also be assessed with the scratch
wounding technique. The use of precision wounding replicators with floating pins and a workstation robot enables large
numbers of scratches to be made with reduced coefficients
of variation [43, 44].
Wound healing assay connected with video-lapse microscopy allows studying in 2D dimension the role of collective migration in angiogenesis and vascular development
[34–36]. The use of aortic rings (see below) and that of specific microfluidic devices represent a further tool to describe
this process in a 3D architecture [45]. For instance, wound
healing assay exploited by single-cell analysis and by using
chimeric EC sheets obtained by infecting cells with different
fluorescent proteins [34, 45, 46] was instrumental to describe
the following steps of EC collective migration: (1) In resting
state, ECs undergo random cell motility in the monolayer
with a regulated dynamics of homotypic cell junctions; (2)
the presence of cell-free space (i.e., the wound) and a chemotactic gradient results in the appearance at the sheet margin of leader cells, which is characterized by an aggressive
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phenotype with prominent stress fibers, ruffling lamellipodia
and enlarged focal adhesions, formation of peripheral actin
cables, and discontinuous adherens junctions, which indicate
mechanical coupling between leader and follower cells in
the migrating cluster [47]; (3) as leaders start to migrate in
the free space, a follower phenotype appears within cells of
the monolayer.
Trans-well cell migration assay—Boyden chamber
assay The Boyden chamber assay is a useful tool to study
chemotaxis and cell invasion. It was originally introduced
by Steven Boyden in the 1960s for the analysis of leukocyte
chemotaxis [48], and a modified version of the assay has
recently been used extensively for the assessment of monocyte migration [39]. Indeed, today, a large range of Boyden
chamber devices, adapted to individual needs, are commercially available. The assay is based on a chamber of two
medium-filled compartments separated by a microporous
membrane of defined pore size and can be used to distinguish positive chemotaxis (migration toward the attractant)
and negative (migration away from a repellent) chemotaxis.
Briefly, ECs are placed in the upper compartment and are
allowed to migrate through the pores of the membrane into
the lower compartment. The chemotactic agent of interest or
cells-secreting chemotactic agents are present in the lower
compartment. The membrane between the fluid-filled compartments is harvested, fixed, and stained after a defined
incubation time, and the number of cells that have migrated
to the bottom side of the membrane is determined by staining and subsequent microscopic analysis. Because a chemical gradient cannot be maintained for extended periods,
Boyden chamber assays are limited to 2–6 h.
Boyden chamber assays are also utilized to measure different types of chemotaxis, including haptotaxis, transmigration, and cell invasion. Angiogenesis and transendothelial
migration are special forms of haptotaxis, as the trigger
for migration requires not only a chemokine, but also the
presence of cell surface or extracellular matrix (ECM)
molecules. In this case, the insert on the bottom of the
Boyden chamber is coated with purified cell surface ligands
(e.g., ICAM1, VCAM1) or ECM proteins (e.g., collagens,
fibronectin), evaluating the migration of cells exposed to
specific adhesion sites. Transmigration describes the migration of cells, such as leukocytes or tumor cells, through the
vascular endothelium and toward a chemoattractant. Therefore, the assay measures transmigration of cells through a
confluent, tight EC layer. Angiogenesis requires the invasion of EC through the basement membrane to form sprouting capillaries. Invasion processes can also be modeled in
a Boyden chamber assay by coating the well membrane
with a layer of Matrigel or collagen. In this case, cells must
secrete matrix metalloproteases to degrade matrix proteins
and migrate (invade).
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The bioactive molecules in Boyden chamber assays can
be provided directly by adding recombinant proteins or small
molecule compounds or by plating cells-secreting specific
factors in the bottom chamber. Manipulation of test cells
(gain-of-function/loss-of-function) can be introduced into
the assay. Migration assays are often performed in co-culture
with tumor cells, pericytes or fibroblasts.
The real-time random migration assay The use of videolapse microscopy allows measuring migration not only as
an endpoint result, but gives information on single-cell
parameters, on morphological changes, and on the influence
exerted by specific substratum. Subconfluent ECs are plated
on plastic surface coated with specific extracellular matrix
proteins (i.e., collagens, fibronectin, vitronectin) allowed
to adhere and then observed with an inverted microscope
equipped with thermostatic and CO2-controlled chamber
(e.g., Leica, DMi8 platform; Nikon, TE microscope). Images
of motile ECs are captured with a 5 min time interval over
4 h. Images were then processed with DIAS software (Solltech). A recent review on tracking algorithms offers a wide
and comprehensive selection of the available tools to analyze
cell motility [49]. Generally, data are displayed as a centroid
plot showing the location of the geometrical center of the
cell as a function of time. Directional persistence was calculated by determining the ratio between the net path length
and the total path length. Furthermore, other parameters
such as the total and net distance, the speed, the feature of
turning angle can be calculated. Single-cell trajectories were
plotted using MATLAB software and displayed in windrose
graphs [50, 51].
2.2 Limitations and challenges
Standardization of techniques is one of the most critical
issues to ensure the reproducibility of experimental results,
and one has to be aware that cellular in vitro systems represent only a surrogate of the in vivo conditions. Nevertheless, compared to in vivo experiments, in vitro assays are
relatively simple to perform and they offer the possibility to
pursue high-throughput screens of compounds or supernatants of tumor cells affecting EC migration, e.g., supernatants of tumor cells. Nonetheless, the assay has limitations.
The cell culture conditions must be standardized, and pure
populations of EC are required. More frequently than not,
human umbilical vein ECs (HUVECs) are used for these
assays; however, these are derived from a large vessel,
whereas angiogenesis occurs in microvessels. HUVECs are
primary cells and are only viable for a limited time, and like
other cells in culture, they change their expression profile
and therefore their phenotype and behavior over time and
through repeated passage events. Furthermore, reproducibility of scratch assays relies strongly on the initial degree of
confluency [52]. In addition, scratch assays must be carefully
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evaluated to rule out the possibility that experimental conditions have influenced proliferation of EC, rather than their
migration (this can be easily done using cell proliferation
assays such as expression of pHH3, see below). Another
important consideration is that ECs in vivo are exposed to
shear stress as well as hypoxia gradients, i.e., the drag force
induced by blood flow to the endothelium, which are absent
in the standard static cell culture models. While the scratch
assay is a straightforward cell culture assay to analyze EC
chemokinesis, it does not have a high degree of sensitivity,
but it is a useful tool to perform large-scale screening experiments. One downside of this method is that the width of the
scratch introduced into the cell layer is difficult to control
and cannot be easily standardized. Moreover, wounding of
the monolayer with a sharp object may scratch the surface
of the cell culture dish and additionally damage the EC at
the migration front.
To obtain more reliable and reproducible results, fencing techniques that allow for controlled release of a confluent monolayer have been developed. These enable lateral
migration without wounding of the cells or the underlying
matrix [53]. Cells are grown as a monolayer in a culture dish
containing a silicon template of defined size prior to seeding
the cells. Once the culture reaches confluency, the silicon
template is removed, thus allowing the cells to migrate laterally into the area previously occupied by the silicon template. Precise microscopic quantification of lateral migration
is then possible over 2–4 days. Ideally, the duration of the
assays should be kept under 2 days, because over longer
incubation times, cell proliferation will contribute to wound
closure, thereby confounding the migration effect.
As previously mentioned, a potential drawback is the difficulty of standardizing the wound areas, but this can be
overcome using silicon templates [53]. Likewise, several
commercial suppliers have developed robust assays that also
circumvent this problem. For example, Essen Biosciences
marketed a mechanical pin tool, the WoundMaker™,
which applies equal pressure to create 96 homogeneous
700–800 µm scratches in each microplate well (www.essen
biosc ience .com/en/produ cts/incuc yte). Wound healing is
visualized in real time using the IncuCyte ZOOM™, and
cell tracing is fully automated, thus facilitating analysis and
reproducibility. A similar device, providing automated hardware and analysis software, is provided by Peira Scientific
Instruments (Beerse, Belgium) [54]. Electric cell-substrate
impedance sensing (ECIS, Applied Biophysics Inc, New
York, USA) can also be used to automatically create wounds
and simultaneously measure cell migration in multiple wells,
and V&P Scientific has developed a wounding replicator
using floating pins that deliver a precise scratch (www.vpscientific.com/wounding_tissue_culture_exper iments.php)
by a Sciclone ALH 3000 workstation robot (Caliper Life
Sciences, Hopkinton, MA). The additional precision of the
Angiogenesis (2018) 21:425–532
scratches using the robot over manual wounding replicators makes the latter instrument the method of choice when
performing the scratch wound assay for large-scale screens
on drug or genome-wide siRNA/sgRNA screens [43, 44].
It should be stressed that the Boyden chamber assay is
somewhat delicate and it requires experience in handling.
The most critical issue is the possible trapping of air bubbles in the lower and upper chambers during assembly. Air
bubbles appear as empty spaces on the filter at the conclusion of the assay, because they hinder cell migration. For
manual analysis, this may not be of major importance, but
it becomes relevant if an automated analysis is performed
since trapped cells would be undistinguishable from the
absence of migration. It is important to invest substantial
time into the setup and troubleshooting of the assay in order
to yield robust and reliable results. It is also recommended
to include a checkerboard analysis to distinguish between
chemotaxis and chemokinesis effects. To this end, different
dilutions of the compound to be tested should be titrated in
the upper and lower chamber. Equal concentrations in the
upper and lower chamber should lead to the same migration behavior as in the control for a compound that strictly
responds to a gradient (i.e., chemotaxis).
2.3 Concluding remarks
In summary, the lateral scratch wound assay [55] and the
Boyden chamber assay are both robust and reliable platforms
to study EC migration. They are suitable for scaled-up purposes in order to perform manual or automated large-scale
compound screens. Various vendors provide scratch and
transwell assay systems. Although these systems offer good
reproducibility and adequate throughput capacity, variation
between individual EC isolations can occur resulting in variable results. Pooling several EC isolates may reduce this
variability. In combination with time-lapse microscopy, it
is also a powerful tool for tracing the migratory behavior of
individual EC.
3 Endothelial cell proliferation assays
Many regulators of angiogenesis have been identified, validated, and developed based on their effects on EC proliferation. ECs are among the most quiescent cells in the body,
with proliferation rates approaching zero under steady-state
conditions. Only after stimulation, usually as a consequence
of injury, inflammation, or pathological processes such as
malignant growth, can they initiate cell cycle entry [56, 57].
The ideal assay to measure EC proliferation should be rapid,
reproducible, and reliable and wherever possible should
exclude inter-operator variability, for example through
quantitative computational readout rather than qualitative
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researcher-dependent observations [57]. This section presents different methods and elaborates on problems and
pitfalls.
3.1 Types of proliferation assays
A number of different approaches to address cell proliferation have been developed in the last decades. In general,
these include assessment of cell number, detection of DNA
synthesis by incorporation of labeled nucleotide analogs,
measurement of DNA content, detection of proliferation
markers and metabolic assays (Fig. 1). Depending on the
broadness of the definition of cell proliferation, which can
range from the narrow description, for example, “the fraction of cells dividing over time” to the more general “the
doubling time of a population,” several different assays may
be pursued. Apart from that, means and equipment available will also dictate the choice of a particular method. As
all methods focus on a particular aspect of the process, it is
highly recommended to verify results with a complementary
assay.
Cell counting Cell counting is considered the gold standard for proliferation. Moreover, at least in theory, it is one
of the most straightforward procedures for measuring proliferation of a cell population. It can be done using automated
cell counters (e.g., Beckman Coulter) or by using a hemocytometer after removal of the cells from the culture vessel
[57, 58]. More recently, different automated platforms have
entered the market that allow analysis of cells while present
in microplates, such as plate cytometers, automated microscope, or high-content screening platforms, that are compatible with cell counting-like procedures. With these, cells can
be monitored over time but frequently require staining for
detection, calibration, and (computation-assisted) quantification by, for example, staining of nuclei. Moreover, real-time
cell analysis (referred to as RTCA) platforms have emerged
that allow label-free, automated, real-time monitoring of cellular properties during incubation based on electrical resistance measurements. The equipment, however, requires considerable investment, beyond reach for many laboratories.
DNA labeling During S phase of the cell cycle, DNA is
synthesized and subsequently divided between the daughter cells (2 N → 4 N → 2 N; N = number of a complete set
of chromosomes). Addition of modified nucleotides to the
culture medium will result in their incorporation into the
newly synthesized DNA. Adhering to the narrow definition of proliferation as stated above, this type of assay most
closely reflects a means of measuring the fraction of actively
dividing cells. It should be noted that this technique does not
directly measure cell division or population doublings, but
exclusively incorporation of a tracer into DNA synthesis.
3
H-thymidine has been used in proliferation assays for
decades [19, 56, 58, 59]. Briefly, cells are pulsed with
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3
H-thymidine for several hours and radioactivity is measured by liquid scintillation counting. This provides a very
accurate representation of DNA synthesis and it is highly
sensitive since the amount of incorporated 3H-thymidine is
directly proportional to the rate of DNA synthesis [57, 58].
Constraints on using radioactive compounds and the rise of
alternative methods have limited its use. In a similar manner, the incorporation of 5-bromo-2′-deoxyuridine (BrdU) or
EdU (5-ethynyl-2′deoxyuridine) can be measured. BrdU or
EdU can be (in)directly detected and subsequently be (semi-)
quantified using ELISA, flow cytometry, or immunohistochemistry [57, 58, 60]. The latter two quantification techniques allow one to determine the fraction of dividing cells.
These uridine analogs can be combined with DNA dyes (see
below) to gain additional cell cycle information [57].
Another approach is the measurement of cellular DNA
content using intercalating dyes such as PI (propidium
iodide) or DAPI (4′,6-diamidino-2-phenylindole). Using
flow or plate cytometry, a profile of the distribution of cells
over the different phases of the cell cycle can be visualized,
represented by DNA contents of 2 N (G1/0), 4 N (G2/M),
or mixed (S). In addition, this method allows for the detection of apoptotic cells that would exhibit a subG1/0 (< 2 N)
DNA content.
An alternative method to study EC cycle is based on the
use of Fucci (fluorescent, ubiquitination-based cell cycle
indicator) technology (Thermo fisher). It consists of a fluorescent protein-based system that employs both a red and
a green fluorescent protein, respectively, fused to cdt1 and
geminin, which are two regulators of cell cycle. These two
proteins are ubiquitinated by specific ubiquitin E3 ligases in
a specific temporal sequence. In the G1 phase, geminin is
degraded; therefore, only cdt1 is present and appears as red
fluorescence within the nuclei. In the S, G2, and M phases,
cdt1 is degraded and only geminin remains, resulting in cells
with green fluorescent nuclei. During the G1/S transition,
when cdt1 levels are decreasing and geminin levels increasing, both proteins are present, giving a nuclear yellow fluorescence. More recently, Fucci probe was reengineered to
generate a triple color-distinct separation of G1, S, and G2
phases extending the use of this technology to quantitative
analyze the interphase of cell cycle [61].
Fig. 1 Endothelial cell proliferation assays. a Phase-contrast image
(left) and binarized image of HUVEC grown in a regular 96-well
plate. Simple software solutions can be used to count features in the
image. b Example of MTT assay, with color intensity correlating with
cell number. c DNA staining profile of HUVEC using PI, measured
on a plate cytometer. d Cell viability of HUVEC exposed to sunitinib,
measured using a luminescent assay
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Proliferation markers Cell division is a highly coordinated process in which specific proteins act in concert with
allow progression through different stages of cell division.
Detection of these proteins, usually through immunochemical procedures, allows the estimation of the fraction of dividing cells. This approach can be used in in vitro endpoint
assays, but can additionally be used to evaluate active EC
proliferation in tissue sections (see separate section). The
most frequently used markers are phosphorylated histone H3
(PH3), which marks cells in S phase, or PCNA and KI67,
which label cells in all cell cycle stages except G0.
Metabolic assays Gradually, the use of cell viability
assays has taken a dominant position in addressing cell proliferation [62]. While not reflecting this property in its narrowest sense, if properly conducted, these assays accurately
represent the number of live cells. They are readily available
and require minimal handling and infrastructure. The most
well known is the MTT assay (3-(4,5-dimethylthiazol-2-yl)2,5-diphenyltetrazolium bromide), in which this yellow salt
is taken up by metabolically active cells and converted by
mitochondrial dehydrogenase to insoluble purple formazan
crystals. As the amount of the converting enzyme is highly
stable in a given cell population, the generation of formazan
(and hence color intensity) is proportional to the number
of viable cells. This is subsequently quantified by solubilization of the crystal-containing cells and spectrophotometry [63]. Variations in this method, for example, involving
less toxic reagents, simplified reaction steps, or alternative
readouts such as cellular ATP levels, have also been widely
used [54, 58, 64]. In particular, the water-soluble tetrazolium salts, such as MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium),
allowing the detection of the colored product directly in cell
culture media without solubilization, facilitate high-throughput and pharmacological screenings [65].
3.2 Limitations and challenges
Endothelial cell culture considerations Studying EC proliferation in vitro requires a purified population of EC
compatible with the assay setup. As already mentioned
before, HUVECs are a widely available source, but have
limitations regarding phenotype and life span in vitro. Therefore, other sources of EC are necessary for confirmation of
results. Foreskin-derived human dermal microvascular ECs
(HDMECs) or human dermal blood microvascular ECs
(HDBECs) are good alternatives; however, the user should
be aware that they represent mixed populations of blood and
lymphatic endothelial cells [66].
It is essential to standardize the protocol of cell culture
and propagation, by adhering to a fixed scheme of passaging,
in order to use passage number as a surrogate for population doublings [67]. In all cases, cell density needs to be
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carefully controlled. Assay linearity can be compromised
when cells are plated at too high (e.g., 50,000 cells/cm2) or
too low (e.g., 5000 cells/cm2) a density. Loss of cell–cell
contact is a potent stimulus for EC to proliferate, whereas
ECs enter a quiescent state upon confluency, a process
known as contact inhibition [57, 58, 63]. Optimization of
the dynamic performance should involve synchronization
of cells by exposure to low serum conditions (when studying pro-angiogenic molecules), or by stimulation prior to
the addition of angiostatic factors or anti-angiogenic drugs
[6]. Though immortalized EC can pose a helpful alternative, it should be recognized that the immortalization itself
will likely alter growth control and survival mechanisms in
these cells [67]. As such, care must be taken to address the
generalizability of assay outcome.
Assay choice considerations Each type of EC proliferation assay described here has limitations. Though cell
counting is the most straightforward method, it can be prone
to sampling error when cell detachment is required. Furthermore, it can be labor-intensive and it requires relatively
large samples [57, 58]. However, it can be executed in the
absence of toxic, mutagenic, or radioactive compounds, such
as metabolic, DNA labeling, and DNA incorporation-based
assays [57, 58, 63]. Of note, the outcome of these assays
is adversely affected by EC death. From a methodological
point of view, each assay has its strengths and weaknesses.
For example, the indirect detection of antigens (e.g., PCNA
or BrdU) requires careful procedural optimization. Although
the alternative “click” chemistry, by which the analogous
EdU can be detected directly, circumvents this issue of
indirect detection, making EdU preferable [57, 58, 60].
Nonetheless, assay readout and interpretation are important
to consider. When measuring incorporation of nucleotide
analogs, one should realize that DNA synthesis is not confined in all situations to chromosomal duplication during S
phase [57, 58]. For example, during DNA repair, nucleotides
are excised and replaced, which is especially relevant when
addressing the action of compounds with a potential DNA
damaging effect. With DNA-intercalating dyes, care must be
taken that doublets are excluded in the gating procedure or
with readout in plate-based systems. By nature, this type of
assay is mostly suited for truly diploid cells, and not for cells
that may display alternative karyotypes. Although the latter
is not a common trait of EC, a few reports have addressed
this matter in tumor-derived EC [68], and personal observations also indicate this may be the case with EC lines.
Finally, test reagents may interfere with readout chemistry;
thus, compounds that affect mitochondrial function are less
compatible with metabolic assays. Surrogate assays such
as the VEGFR-BaF3 cell lines which bind VEGF ligands
and signal through chimeric receptors in reporter cell lines
have been very useful (both academically and in industry)
for quantifying the presence of major angiogenic factors
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from human, mouse, and viral sources that stimulate the
VEGFR-2 and VEGFR-3 pathways [69, 70].
3.3 Concluding remarks
The choice for a particular EC proliferation assay is determined by a number of considerations. Endpoints, test compounds, laboratory infrastructure, scale, required throughput, convenience, and cost all influence the applicability of
an assay system. The growth of a cell population is influenced by both cell division and death, which are difficult to
monitor simultaneously. Most important is the researchers’
awareness of strengths and weaknesses of each individual
assay. Importantly, interpretation of data must be done with
care, and whenever possible, results are validated with an
alternative method.
4 3D models of vascular morphogenesis
The emergence of vascular networks either through vasculogenesis or angiogenesis requires the association of cells
into stable 3D tubes in a process that involves differentiation,
migration, proliferation, aggregation, and rearrangement of
these cells to form cords that then undergo lumen formation. Taken together, this process is referred to as vascular
morphogenesis. Subsequently, ECs recruit perivascular stromal cells (pericytes) to stabilize this newly formed network
and minimize leakage upon blood perfusion. Importantly,
not all sprouts become functional vessels. Pruning serves to
selectively remove redundant or non-functional vessels to
optimize fluid flow through the network [71].
In vitro assays have played a valuable role in our understanding of vascular morphogenesis. These assays provide
a simpler platform than animal models for dissecting individual steps within the process while also incorporating 3D
matrix to mimic native in vivo tissues. Here, we present
several of the most reliable and informative assays developed to date and highlight the strengths and limitations of
each (Table 1). While many types of EC can be used in these
assays, the most commonly used are HUVEC and human
endothelial colony-forming cell-derived EC (ECFC-EC),
which generally have a higher proliferative potential. Mouse
ECs are not generally used in these assays as they are notoriously difficult to maintain in culture. While we use “EC” to
reference to cells derived from both species, assays using a
specific EC source are annotated accordingly.
4.1 Types of assays
Fibrin Bead Assay Traditional Matrigel cord-forming or
collagen I angiogenic invasion assays are insufficient to
mimic the complexity of angiogenesis, as these assays are
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two-dimensional and ECs in these assays often form incomplete lumens. Moreover, lumen formation in Matrigel is not
unique to EC as several non-EC cell types (e.g., human
prostate carcinoma and glioblastoma cells) also form cords,
complicating the interpretation of results from these assays
[57]. By contrast, the fibrin bead assay provides a platform for testing EC sprouting and lumen formation over an
extended period (2–3 weeks) and incorporates a 3D, ECM,
and multiple cell types (i.e., stromal pericytes) to model
native angiogenesis. ECs (HUVECs) are first allowed to
adhere to collagen I-coated Cytodex beads to generate an
EC monolayer that mimics the vessel wall of native vessels.
These EC-coated beads are then embedded into a fibrin gel
with human stromal cells either embedded within the gel
or plated as monolayer on the surface of the gel. Tip cells
are observed 2–3 days post-plating, and elongating sprouts
appear 2–4 days thereafter (Fig. 2a, b). When maintained
in pro-angiogenic EGM-2 medium (Lonza), lumens form
within a week and the cells remain viable up to 3 weeks, at
which point anastomoses between sprouts are often apparent [72]. A detailed, video protocol of this assay is available
[73].
Angiogenic sprouting from individual beads is evaluated by phase-contrast microscopy allowing for quantification of sprout number, length of sprouts, percentage of
sprout lumenization, and number of anastomoses. Genetic
approaches (siRNA, lentiviral transduction, CRISPR/Cas9)
[74] can modify gene expression in individual cell types
to dissect cell-autonomous components of the angiogenesis
process. Protein expression and localization are measured by
fixing bead assays and using modified immunofluorescent
staining techniques. More detailed gene expression analyses
are made possible by harvesting individual cell types to track
RNA expression changes over time through various stages
of sprouting angiogenesis.
The use of stromal cells (e.g., lung fibroblasts) is critical
to the success of this assay, as these cells secrete angiogenic
factors necessary for EC sprouting and lumen formation,
including growth factors [hepatocyte growth factor (HGF),
transforming growth factor alpha (TGF-α), and angiopoietin-1 (Ang-1)], as well as matrix molecules, matrix-modifying proteins and matricellular proteins [e.g., procollagen
C endopeptidase enhancer 1, secreted protein acidic and
rich in cysteine (SPARC), transforming growth factor-βinduced protein ig-h3 [βIgH3], and insulin growth factor
binding protein 7 (IGFBP7)]. These factors act to locally
stiffen the matrix, which supports sprouting and lumen formation [75]. This assay represents a significant improvement over conventional, single-cell-type angiogenic assays,
as the inclusion of multiple cell types more closely mimics
the physiological environment. Nevertheless, as this assay
uses primary cultures of cells, rather than cell lines, it is
important to remember that batch-to-batch variations in
Angiogenesis (2018) 21:425–532
Table 1 Comparison of
3D models of vascular
morphogenesis
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Assay
Fibrin bead assay
Collagen lumen assay
Retina explant assay
Vascularized micro-organ
Process
Sprouting
Lumen
formation
Anastomosis
Pericyte
Perfusion
recruitment
✓
✓
✓
✓
✓
✓
✓
✓
✓
✓
✓
✓
✓
✓
Pruning
✓
stromal cells (and EC) can significantly affect assay results.
To partially overcome these issues, it should be appropriate
to use ECs pooled from 5 to 10 umbilical cords. Generally,
it is crucial to identify stromal cell-HUVEC pairs that yield
optimal angiogenic sprouts.
Collagen Lumen Assay To investigate EC lumen formation mechanisms, early assays seeded EC in monolayers on
plastic dishes coated with ECM proteins (i.e., collagen I,
collagen III, fibrin, or Matrigel). While these 2D assays are
sufficient to induce EC cord formation [76–78], they cannot
reproduce the necessary cues for true lumen formation found
in native, 3D tissues. Collagen sandwich assays surround
EC within a 3D matrix by seeding the cells in monolayer
on collagen I matrix and then covering them with a second
layer of collagen [79]. Nevertheless, tube formation fails to
occur in a random, 3D growth pattern, forming only in the
X–Y plane of the initial gel layer and not in the z-axis. As
this does not adequately recapitulate normal vessel growth
in a true 3D environment, George Davis and others further
optimized these assays, opting instead of embedding single
EC (HUVEC) randomly throughout a collagen I matrix. In
the simplest version of these assays, HUVECs are seeded
at low density (7 × 105 cells/ml) under serum-free growth
conditions and with the addition of minimal growth factors (phorbol ester, VEGF), and fibroblast growth factor-2
(FGF-2). After 48 h, the embedded ECs form intact tubes
throughout the gel, with clearly demarcated lumens (Fig. 2c,
d). Several variations on this assay have since enhanced and
optimized lumen formation. First, the addition of several
other growth factors, including a cocktail of stem cell factor (SCF), IL-3, stromal-derived factor-1α (SDF-1α), and
FGF-2, further promotes lumen formation while maintaining serum-free growth conditions. Second, when simultaneously seeded within the same matrix, stromal pericytes
are recruited by EC, recapitulating a key step in vascular
morphogenesis. Lastly, to understand the process of EC
sprouting and angiogenesis, EC can be seeded on top of
a 3D collagen gel containing the same growth factors and
Fig. 2 Three-dimensional assays of vascular morphogenesis. a A
fibrin bead assay uses collagen I- and EC-coated Cytodex beads
embedded within a 3D fibrin gel matrix to measure EC sprouting and
lumen formation. b These features are readily resolved using phasecontrast microscopy. c EC tube formation can be measured by embedding EC within a collagen I matrix. d Once formed, these tubes can
be visualized by toluidine blue staining and bright-field microscopy.
e Whole-mount, dissected retinas from postnatal mice are mounted
within collagen I-Matrigel matrix mix and cultured in pro-angiogenic
medium to stimulate EC sprouting. f Sprout and lumen formation are
resolved using phase-contrast microscopy. g The vascularized microorgan (VMO) approach utilizes “arteriole” (high pressure) and “venule” (low pressure) microfluidic channels to drive medium diffusion
and flow across a cell chamber where microvasculature forms. h The
formed microvasculature (EC, red) can be measured for leak by perfusion with 70 kDa FITC-dextran (green)
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invasion of the underlying gel layer can be quantified. A
detailed protocol of the collagen lumen assay and its variations is available for further reading [80].
Real-time imaging of tube formation can be achieved
using fluorescent protein-transduced EC. Alternatively, fixed
vessels can be stained with 0.1% toluidine blue and imaged
using bright-field microscopy (Fig. 2d). More in-depth analyses can be carried out on these fixed vessels using immunofluorescence staining of relevant protein markers or transmission electron microscopy to resolve structural details of
formed lumen and remodeled ECM.
Regular users of collagen gels will note that the viscosity, pH, and contraction of these gels can hinder successful
execution of assays in the hands of new users. As a result,
special care should be taken when pipetting (such as when
mixing cells and growth factors) and plating gels to ensure
even gel coating of the bottom of the well plate. Perhaps
most significantly, early gel contraction can limit the useful length of these assays. Users will note that plating gels
only in wells within the center of the 96 half-area wells and
adding medium or water to the outer wells of the plate will
minimize gel contraction, by maintaining local humidity
levels. Additionally, seeding fewer ECs within the collagen
(1.5 × 103 cells/ml) can minimize gel contraction and prolong the assay.
Retinal Explant Assay Although in vitro assays are high
throughput and can mimic major steps in vascular morphogenesis, they do not fully recapitulate the in vivo, wholeorgan environment [72]. Several in vivo animal models,
such as mouse retina or zebrafish fins, are valuable tools
for studying vascular (re)-establishment in a physiologically
relevant context [57, 81]. However, the added complexity
of these systems makes it more difficult to ascertain the
role of individual proteins and growth factors and cell types
in the vascular morphogenesis process, relying on genetic
manipulations or system-wide administration of pharmacologic inhibitors to dissect molecular pathways [57, 82].
As such, there is a need to increase assay complexity and
physiological relevance while developing platforms amenable to ex vivo study in the laboratory. Retina explant assays
are one such ex vivo platform, whereby dissected retinas
are maintained and observed for vascular morphogenesis
over several weeks in the laboratory. While multiple versions of this assay have been published, a protocol published
by Sawamiphak et al. is most widely used for the study of
endothelial sprouting [83]. Briefly, retina cups from embryonic, postnatal, or adult mice are harvested and cut radially
to allow flat mounting of the retina interior surface onto a
membrane insert. After recovery in media for 2–4 h, the
explants can then be treated with stimulatory or inhibitory
agents for up to 4 h, followed by whole-mount microscopy
analysis to evaluate the (anti-) angiogenic effect of these
agents on vessel sprouting (Fig. 2e). A trained researcher
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can harvest and dissect each pair of retinas within minutes.
Unfortunately, without the support of a 3D matrix, retinal
cells cannot survive for long periods, thus making studies
of later stages of angiogenesis impossible. To overcome
this, Rezzola et al. have improved the assay by embedding
the retinas in different matrices after dissection [84]. In this
approach, retinas can be crosscut into four equal pieces and
left in serum-free media overnight. The retina fragments are
then embedded in Matrigel, collagen I, or fibrin matrix and
fed every 2–3 days. Depending on the age of the mice and
the matrix used, sprouts can be observed between days 3 and
6 and anastomosis of neighboring sprouts similar to what
occurs in vivo can be observed in 10–14 days [85] (Fig. 2f).
These explants can be maintained up to 3 weeks before the
vessels eventually regress. When cultured for a longer period
however, vascular sprouts also start to form toward the chorioidea and not only in the retinal plane, making analysis
more complex.
Vessel formation can be analyzed in real time using timelapse imaging or immunofluorescence microscopy of fixed
explants at established experimental time points [85, 86].
Gene expression can be manipulated by genetic crossing of
the donor mice or, more transiently, by treating retinas with
lentivirus or siRNA or Crispr-Casp9 technology. Moreover,
embedded retinas can be treated with drugs over extended
periods to dissect individual signaling pathways.
Several factors are critical to consistently achieve sprouting from dissected retina explants. Firstly, the matrix proteins in which retinas are embedded can greatly influence
how vessels sprout. Matrigel is far superior to single matrix
proteins in inducing sprouting. However, the addition of
10–20% Matrigel in collagen I matrix is sufficient to stimulate sprouting compared to pure collagen I matrix. Secondly,
the use of pro-angiogenic EGM-2 medium yields more
sprouts as compared to basal medium alone. Lastly, as with
any tissue explant, the age of the mouse can influence the
degree of vessel sprouting. As such, special care should be
taken to select mice appropriate for the experimental question at hand. There are many similarities between the mouse
retinal explant assay and the traditional mouse/rat aortic ring
angiogenesis assay or rat vena cava explant assay [87–89].
However, retinal explant models more closely model true
capillary sprouting as the vasculature in these explants is
actively developing and remodeling. This makes the retinal
explant model uniquely suited to studying microvessel formation and its underlying mechanisms.
Vascularized Micro-Organ Platform To understand all
the steps of vascular morphogenesis in a single platform, a
vascularized micro-organ (VMO) approach has been developed to drive formation of a perfusable vascular network
within a 3D hydrogel matrix environment. In contrast to the
assays described above, VMO-embedded ECs are exposed
to and respond to shear stress, form lumenized vessels, and
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are perfused with a blood-substitute medium that delivers
nutrients to tissues within the matrix, just as in the body.
Specifically, this platform utilizes “arteriole” (high pressure)
and “venule” (low pressure) microfluidic channels that are
joined by a living microvascular network that forms by vasculogenesis in an intervening tissue chamber (Fig. 2g). A
pressure differential between the two channels is used to
drive interstitial flow through the fibrin gel matrix during
vessel development, induce vessel formation (through shearsensing), and drive convective flow through the mature vasculature once formed and anastomosed to the outer channels.
This pressure difference is induced by varying the level of
medium within fluid reservoirs at either end of the microfluidic channels, thereby creating hydrostatic pressure heads
that ensure continuous fluid convection across the cell chamber. To form vessels, human ECFC-EC and human lung
stromal cells are co-loaded within a fibrin matrix into the
central cell chamber through an independent loading tunnel. When maintained in pro-angiogenic EGM-2 medium,
vessels form within 4–6 days post-loading (Fig. 2h). When
perfused with 70 kDa rhodamine dextran, a molecule similar
in size to albumin, these vessels demonstrate minimal vessel
leakage—comparable to in vivo microvasculature. For readers interested in more information, a detailed protocol for
loading and maintaining the VMO platform is available [90].
The VMO platform is fabricated from polydimethylsiloxane (PDMS), an optically clear, biologically inert polymer
widely used in the microfluidics field [91]. The use of this
polymer and the dimensions of the platform ensure that live,
GFP-transduced endothelial progenitor cells (ECFC-EC) can
be imaged and quantified throughout vessel formation. Specific parameters such as vessel network length, branching,
and anastomosis can be measured in real time as can vessel permeability by perfusion with fluorophore-tagged dextran molecules of various molecular weights. Additionally,
immunofluorescent staining can be used to quantify expression of specific molecular markers or RNA can be collected
to measure changes in gene expression. Lastly, gene expression can be manipulated by treating individual cell types
with lentivirus or siRNA prior to loading in the platform.
To ensure robust and reproducible vascular network formation, several steps are critical. First, the fibrin gel matrix
must be consistently loaded within the VMO cell chamber.
During normal loading, perfusion burst valves at the interface between the tissue chamber and the microfluidic channels ensure a gel/air interface (later a gel/fluid interface)
is formed. To simplify loading and minimize specialized
training for new users of the platform, current iterations of
the VMO platform incorporate a pressure release valve at
the loading tunnel that minimizes unintended gel bursting
[92]. Second, robust vascular network formation requires
that vessels within the chamber anastomose with the outer
microfluidic channels. To facilitate the formation of these
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anastomoses, EC can be either seeded directly within the
microfluidic channels or induced to migrate from the gel by
coating the external channels with ECM [93]. As with the
fibrin bead assay, optimal stromal cell–EC pairs should be
validated to ensure assay reproducibility.
4.2 Limitations and challenges
With all EC assays, the source of EC is critical to assay success. Although commercial versions of HUVEC and ECFCEC are available, cells from these sources show limited utility in many 3D assays, likely due to a larger than optimal
number of cell doublings prior to shipment and use in the
laboratory. As a result, the use of freshly isolated EC will
provide the most consistent results and is strongly encouraged. Readers will note that isolation protocols for both cell
types are available [94, 95]. Additionally, patient-to-patient
variation between different EC isolations can lead to inconsistent assay results, an issue that may be avoided by pooling
several EC lines prior to use.
Given the many differences between the assays described
here, the useful length of these assays varies considerably.
Even in well-trained hands, the contraction of gels in the
collagen lumen assay effectively limits the useful time frame
of assays to 72 h or less. However, the other assays described
here can continue for much longer periods of time, with the
fibrin bead assay, retinal explant assay, and VMO platform
all suitable for time points up to 3 weeks under appropriate
conditions.
Lastly, the majority of these assays can be run in a relatively high-throughput manner, thereby accelerating the
speed with which genetic, molecular, or pharmacologic
screens [96] can be conducted to understand vascular morphogenesis. This is especially true with the fibrin bead and
collagen lumen assays, which utilize multiple beads or
multi-well culture plates to increase assay throughput. Similarly, while initial versions of the VMO platform were cumbersome to load in high-throughput numbers, this platform
is now used in an optimized configuration that incorporates
up to 16 individual VMO devices within a standard 96-well
plate [97]. This design simplifies translation to outside laboratories and interfacing with existing microscope and plate
reader infrastructure. Of all the assays described in this section, retinal explants are most adversely affected by delays
between initial dissection, mounting, and plating of tissue
samples. This inherently limits the number of animals that
can be dissected at once and, for now, limits the number of
retinas that can be screened simultaneously.
4.3 Concluding remarks
Vascular morphogenesis requires the interaction
between several cell types and their surrounding, 3D
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microenvironment. The 3D vascular morphogenesis models described here provide unique in vitro culture systems
that recapitulate this complex multi-step process. At the
same time, the simplicity and consistency of these assays
compared to in vivo models allow them to be conducted
in a high-throughput fashion. Finally, the use of human
cells in these assays improves the clinical relevance of such
models, making them readily amenable to drug discovery
applications.
5 Aortic ring assay
Explants of rat or mouse aorta have the capacity to sprout
and form branching microvessels ex vivo when embedded
in gels of ECM. Angiogenesis in this system is driven by
endogenous growth factors released by the aorta and its outgrowth in response to the injury of the dissection procedure.
This property of the aortic wall was first described in the
early 1980s [98] and led to the development of the aortic
ring assay [99], which is now widely used to study basic
mechanisms of angiogenesis and test the efficacy of proangiogenic or anti-angiogenic compounds [100].
5.1 Benefits and strengths of the aortic ring assay
The aortic ring assay offers many advantages over existing models of angiogenesis. Unlike isolated EC, the native
endothelium of the aortic explants has not been modified by
repeated passages in culture and retains its original properties. The angiogenic response can be inhibited or stimulated
with angiogenic regulators and analyzed by molecular or
immunochemical methods without the confounding effects
of serum (Fig. 3a, b). Angiogenic sprouting occurs in the
presence of pericytes, macrophages, and fibroblasts, as seen
during wound healing in vivo [100]. The ultrastructure of
neovessels at different stages of development can be evaluated by electron microscopy (Fig. 3c). The different cell
types can be identified with specific cell markers by immunostaining whole-mount preparations [101] of the aortic
cultures (Fig. 3d–g). Many assays can be prepared from the
thoracic aorta of a single animal (approx. 20–25 cultures/rat
aorta; approx. 10–15 cultures/mouse aorta). The angiogenic
response can be quantitated over time, generating curves of
microvascular growth. Aortic cultures can be used to study
mechanisms of vascular regression, which typically follows
the aortic angiogenic response as seen during reactive angiogenesis in vivo. Aortic rings transduced with viral constructs
or obtained from genetically modified mice can be used to
study the role of specific gene products in the regulation of
the angiogenic response [100].
Recently, rat aortic ring assay was adapted to human
arteries by using matrigel as 3D hydrogel [102].
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5.2 Assay overview
A detailed description of the aortic ring assay protocol is
available in previous reports [103, 104]. We provide here a
summary of key steps for the preparation of the assay. Aortic
rings are prepared from the thoracic aorta of 1–2-month-old
rats or mice. After excision from the animal, the aorta is
transferred to a Felsen dish containing serum-free Endothelial Basal Medium (EBM). Under a dissecting microscope,
the aorta is cleaned of blood and fibroadipose tissue using
Noyes scissors and microdissection forceps. Care is taken
not to stretch, cut, or crush the aortic wall during the isolation and dissection procedures. As the dissection progresses,
the aorta is rinsed in the four compartments of the Felsen
dish. Using a scalpel blade, the aortic tube is then crosssectioned into 0.5–1-mm-long rings. The most proximal
and distal rings which may have been damaged during the
dissection procedure are discarded. The remaining rings
are washed through sequential transfers into eight consecutive baths of serum-free medium, using compartmentalized
Felsen dishes. Aortic rings are then embedded individually
into thin collagen, fibrin, or basement membrane gels as
described. Once the gel has set, 500 μl of serum-free EBM
is added to each culture. Each experimental group comprises
quadruplicate cultures in four-well NUNC dishes. Aortic
ring cultures are incubated in a humidified CO2 incubator at
37 °C for 7–21 days.
5.3 Quantitative analysis of angiogenesis in aortic
cultures
The angiogenic response of the rat (or mouse) aorta can be
quantitated by visual counts or by computer-assisted imaging. For visual counts, cultures are examined every 2–3 days
and scored for angiogenic sprouting by using an inverted
microscope with bright-field optics equipped with 4× to 10×
objectives and a 10× eyepiece. Angiogenesis is scored by
counting microvessel sprouts, branches, and loops according to previously published criteria [99]. Aortic outgrowths
can also be quantified by image analysis using low power
images of the cultures thresholded to highlight the vascular
outgrowths [105–108]. Standard statistical methods are used
to analyze data and determine levels of significance between
control and treated cultures. An internal control group with
untreated aortic rings must be included in each experiment
to mitigate the effect of possible interassay variability.
5.4 Critical points
For this assay, we recommend using the thoracic aorta
because of its uniform size and intercostal artery branching
pattern. The abdominal artery can also be used, but its variable pattern of collaterals and tapering lumen may introduce
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437
Fig. 3 Aortic ring assay of
angiogenesis. a Serum-free
collagen gel culture of rat aorta
(asterisk) photographed at day
6 (microvessels marked by
arrowheads). b Aortic culture
treated with VEGF (5 ng/ml)
shows increased number of
microvessels (day 6). c Electron
micrograph of aorta-derived
microvessel with polarized
endothelium (E), patent lumen
(L), and surrounding pericytes
(P); endothelial tight junctions are marked by arrows. d
Phase-contrast micrograph of
microvessel composed of an
inner core of endothelial cells
and surrounding pericytes
(white arrowheads). e Pericytes
highlighted with immunoperoxidase stain for NG2. f Immunofluorescent image of aortaderived macrophages stained
for CD45; an isolectin B4
(IB4)-stained endothelial sprout
is visible in the background. g
Confocal image of microvessel
double stained for endothelial
cells (IB4) and pericytes (alpha
smooth muscle actin, αSMA).
Magnification bars = 500 μm (a,
b), 5 μm (c), 50 μm (d–g)
variability in the angiogenic response. Injury to the aortic
endothelium may be an additional cause of uneven sprouting
from different rings. Therefore, special care must be taken
not to damage the aorta by stretching or letting it dry during the isolation and dissection procedures. Dissection of
the aorta and preparation of the aortic ring cultures are best
performed in a tissue culture room with HEPA-filtered air to
avoid microbial contamination. Best results with this assay
are obtained using interstitial collagen or fibrin gels. Collagen can be produced in-house, as described [103, 104],
or purchased from commercial sources [109]. Fibrinogen
and thrombin for the fibrin gel are commercially available.
Matrigel, a basement membrane-like matrix of tumor origin,
can also be used [110]. Matrigel cultures, however, require
growth factor supplements due to the limited ability of the
aortic rings to sprout spontaneously in this dense matrix. The
growth medium used for the assay should be optimized for
the growth of EC in the absence of serum. Optimal results
in collagen and fibrin cultures can be obtained with EBM.
When preparing individual collagen gel cultures, given the
small volume of gel (20–30 µl), it is important to remove
excess growth medium from the aortic rings when they are
transferred into the collagen, fibrinogen, or Matrigel solution. This is accomplished by gently streaking the aortic ring
onto the bottom of the culture dish while holding it from the
adventitial side with microdissection forceps. When working with fibrin gels, which set rapidly, no more than four
cultures at a time should be prepared, to avoid disrupting
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the developing gel while positioning each ring. In addition,
for fibrin cultures, the culture medium should include a plasmin inhibitor such as epsilon aminocaproic acid to inhibit
fibrinolysis by the aortic rings, which would rapidly destroy
the matrix needed for EC to sprout.
The rat aortic ring assay is robust and very reproducible
when performed by an experienced operator. The mouse aortic ring assay is more variable than the rat aortic ring assay,
likely because of the small size of the rings. For this reason,
at least twice as many aortic rings should be used for this
assay. Miniaturization of the assay using IBIDI microchambers [104] and a smaller volume of growth medium (50 µl)
are recommended for the mouse aortic ring assay to ensure
spontaneous sprouting under serum-free conditions [104]. In
all cases, experiments should be repeated 2–3 times to obtain
sufficient number of data points for statistical analysis. The
growth medium can be replaced on a regular basis (3 times/
week) or left unchanged for the duration of the experiment.
If the medium is not replaced with fresh medium, the angiogenesis response and the stability of neovessels are enhanced
due to accumulation of endogenous growth factors in the
system. For immunohistochemical evaluation of the aortic
cultures, biomatrix gels should not exceed 20–30 µl and
should be well spread as a thin wafer around each ring. Formalin fixation should be limited to 10 min to avoid excessive
cross-linking of proteins. In addition, overnight incubation
may be needed for optimal penetration of the primary antibody into the gel.
5.5 Limitations and challenges
The main limitation of the aortic ring assay is the lack of
blood flow, particularly for angiogenesis-related genes that
are regulated by mechanochemical mechanisms. An additional potential limitation is the source of angiogenic ECs,
which are arterial and not venous, as neovessels in vivo
primarily sprout from postcapillary venules. Many studies
performed with this assay, however, have shown good correlation of results obtained with the aortic ring assay and
in vivo models of angiogenesis. If needed, the aortic ring
assay methodology can be applied to veins as reported [89].
Some investigators have described variability of the angiogenic response in different aortic cultures. This is due to the
delicate nature of the endothelium, which can be damaged
because of inadequate handling of the aorta or the aortic
rings, drying of the explants, or their excessive exposure to
alkaline pH. Suboptimal preparation of the gels resulting in
a defective matrix scaffold can also result in a poor angiogenic response. In addition, the age and genetic background
of the animal significantly affect the capacity of the aortic
rings to sprout spontaneously or in response to angiogenic
factors. Aortic outgrowths in Matrigel are much denser than
in collagen and fibrin and more difficult to quantitate by
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visual counts due to the intricate branching pattern of the
endothelial sprouts and the tendency of mesenchymal cells
to arrange in confounding networks, which mimic angiogenic sprouts. Immunostaining of the aortic outgrowths
with endothelial markers followed by image analysis may
overcome this limitation. For quantitative analysis of the
angiogenic response, the visual count method (described in
detail in Ref. [104]), becomes challenging when cultures
stimulated by growth factors produce 250–300 or more vessels. Since the outgrowths of rings oriented with the luminal axis parallel to the bottom of the culture dish (recommended orientation) are typically symmetrical, angiogenesis
in these cases can be quantified by counting the number
of microvessels in half of each culture and then doubling
the score. Alternatively, these cultures can be measured by
image analysis [105–108]. Finally, for the whole-mount
immunohistochemical stain, gel thinness is critical for optimal antibody penetration.
5.6 Concluding Remarks
Many of the molecular mechanisms orchestrating angiogenesis have been discovered, but many others remain to be
identified, studied, and evaluated as targets for the development of new therapies. The aortic ring assay reproduces
ex vivo cellular and molecular mechanisms that are essential
for the regulation of the angiogenic process. As such, this
assay provides an invaluable platform to test new hypotheses
and analyze the efficacy of the next generation of angiogenesis-targeting drugs.
6 Tumor microvessel density
and histopathological growth patterns
in tumors
Microvessel density (MVD) is often regarded as a surrogate marker of angiogenesis in tumors. Angiogenic tumors
contain areas with a high concentration of new but inefficient blood vessels, which have sprouted from existing
vessels and are often arranged in a tortuous, glomeruloid
tangle. In addition, to these “hot spots,” angiogenic tumors
also contain areas of low blood vessel numbers. Weidner
developed a method to assess MVD in vascular hot spots
using pan-endothelial immunohistochemical (IHC) labeling assays, using lectins such as WGA, Ulex Europaeus, or
antibodies against CD31, CD34, and von Willebrand factor and less often VE-cadherin, αvβ3 integrin, CD105, or
type IV collagen [111]. Vascular hot spots are identified by
light microscopy at low power magnification by scanning
the entire stained tumor section. Individual microvessels are
then counted at high power magnification in these regions.
As an alternative, microvessels can be counted in regions
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defined by systematic uniform random sampling (SURS)
[112]. These IHC staining assays are robust, fast, and easy
to perform and to automate. Although results in individual
studies are highly variable, meta-analyses have confirmed
the prognostic value of MVD, for example in breast cancer
[113] and isolated reports support the predictive value of
MVD for anti-VEGF treatment [114, 115]. However, the
inter-observer variability of the vessel counting algorithm
presents a significant barrier to the use of MVD as a biomarker [116]. Inter-observer variability can be reduced by
applying strict counting rules and by consensus training of
individual observers [112].
Although the importance of angiogenesis to tumor biology is well-established, multiple observations that primary
and metastatic tumors develop and progress in the absence of
angiogenesis suggest that other mechanisms are frequently
involved in tumor vascularization [117]. This represents
a challenge to Folkman’s hypothesis that the growth of a
tumor is only possible when accompanied by angiogenesis
[118]. Moreover, MVD cannot distinguish angiogenic and
non-angiogenic tumors. In addition to the high numbers of
blood vessels in angiogenic tumor hot spots, high MVD is
indeed also observed in non-angiogenic tumors which have
co-opted vessels in organs with extensive microvasculature,
such as the liver and the lungs [117]. The differentiation
of angiogenic and non-angiogenic tumors is, therefore, not
related to the absolute number of vessels per surface area but
to the growth pattern of a tumor.
6.1 Histopathological growth patterns
Histopathological growth patterns (HGPs) are defined
according to the morphological characteristics of the tumor
at the interface with the surrounding normal tissue. HGPs
are identified by light microscopy in standard hematoxylinand-eosin stained tissue sections, and distinct HGPs have
been described for tumors that grow in the lung, liver, skin,
brain, and lymph node [117]. Recently, international consensus guidelines for scoring these HGPs have been described
[119, 120]. One of the important differences in the biology
of tumors with specific HGPs is their means of vascularization. Liver metastases may present with one of two common HGPs, replacement or desmoplastic. In the replacement
HGP, cancer cells “replace” the hepatocytes while co-opting
the sinusoidal blood vessels at the tumor–liver interface.
Patients with colorectal cancer (CRC) liver metastases with a
replacement HGP respond poorly to the anti-VEGF-A treatment, bevacizumab, likely because these tumors utilize vessel co-option instead of angiogenesis [119]. By contrast, in
desmoplastic liver metastases, the cancer cells are separated
from the liver by a rim of desmoplastic tissue in which new
blood vessels are formed by sprouting angiogenesis. Desmoplastic CRC liver metastases showed a better response
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to bevacizumab [119]. Taken together, these observations
strongly suggest that HGPs can be used to guide the choice
of treatment for individual patients with liver metastases.
Across studies, approximately 50% of patients with CRC
liver metastases present with a predominant replacement
growth pattern and this proportion extend to 95% when
patients with breast cancer liver metastases are considered
[119, 120]. This clearly demonstrates that non-angiogenic
tumor growth is not a rare phenotype. The same applies to
primary lung carcinomas and lung metastases for which
angiogenic and non-angiogenic HGPs have also been
described [121, 122]. In the non-angiogenic, alveolar HGP,
cancer cells fill the alveolar spaces and incorporate the capillary blood vessels of the alveolar walls. Approximately, 40%
of the lung metastases from clear cell renal cell carcinoma
(ccRCC) present with a non-angiogenic HGP despite the fact
that nearly all primary ccRCC relies on sprouting angiogenesis, driven by loss of VHL protein function [122].
The prevalence of non-angiogenic tumors and their
resistance to anti-VEGF treatment require the identification
of a biomarker that accurately reflects this type of tumor
growth. HGPs constitute a good candidate biomarker. The
vascular pattern in a tumor section immunostained with panendothelial antibodies and the use of antibodies that mark
EC participating in sprouting angiogenesis are other potential histopathological methods to distinguish non-angiogenic
from angiogenic tumors. Indeed, when the number of hot
spots is determined by nearest neighbor analysis in wholeslide digital images of liver metastasis tissue sections stained
for CD31, the non-angiogenic, replacement-type metastases clearly resemble normal liver tissue. This contrasts with
angiogenic, desmoplastic liver metastases, which show a significantly higher number of vascular hot spots than normal
liver tissue and their non-angiogenic counterparts (Fig. 4).
Several groups have identified “tumor endothelial markers” or TEMs [123–125]. These are proteins that are selectively upregulated on tumor endothelium compared to normal endothelium. Tumor angiogenesis meta-signatures have
been generated using gene expression data of several tumor
types by using “seeds” of transcripts, which were known to
participate in angiogenesis and to be expressed by EC. The
resulting set of EC/angiogenesis-related transcripts that correlated with these seeds was then purified by selecting genes
that were modulated in response to anti-angiogenic treatment
[126]. Interestingly, several of the top ranking genes in this
signature have been confirmed as TEM by other independent
teams [127]. Although we do not currently know whether
TEMs are expressed equally on both angiogenic tumor vessels and co-opted vessels in tumors, it will be intriguing to
assess the expression of these genes, for example ELTD1,
CLEC14a, and ROBO4 in the vasculature of replacement
versus desmoplastic liver metastases. In primary and secondary non-angiogenic human lung tumors, only preexisting
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◂Fig. 4 Microvessel density and histopathological growth patterns. a
Unsupervised spatial modeling of the blood vessel pattern in normal
liver shows a low number of clusters per number of vessel profiles.
A selected region of interest (ROI) at the tumor–liver interface of
normal liver in CD31-stained tissue is shown (left). The Blood Vessel Analysis algorithm of Definiens™ segments and classifies blood
vessel objects (orange) and nuclei (blue) (mid). The Cartesian coordinates (x, y) of the centroids of all vessel objects in one ROI were used
in a simplified “SeedLink” clustering method [646] (right). Centroids
with the same color (e.g., red) belong to the same cluster. b Unsupervised spatial modeling of the blood vessel pattern in a colorectal cancer liver metastasis with a replacement growth pattern shows a low
number of clusters per number of vessel profiles. A selected region
of interest (ROI) at the tumor–liver interface of replacement growth
pattern in CD31-stained tissue is shown (left). The Blood Vessel
Analysis algorithm of Definiens™ segments and classifies blood vessel objects (red) and nuclei (blue) (mid). The Cartesian coordinates
(x, y) of the centroids of all vessel objects in one ROI were used in
a simplified “SeedLink” clustering method [646] (right). Centroids
with the same color (e.g. red) belong to the same cluster. c Unsupervised spatial modeling of the blood vessel pattern in a colorectal cancer liver metastasis with a desmoplastic growth pattern shows a high
number of clusters per number of vessel profiles. A selected region
of interest (ROI) at the tumor–liver interface of desmoplastic growth
pattern in CD31-stained tissue is shown (left). The Blood Vessel
Analysis algorithm of Definiens™ segments and classifies blood vessel objects (red) and nuclei (blue) (mid). The Cartesian coordinates
(x, y) of the centroids of all vessel objects in one ROI were used in
a simplified “SeedLink” clustering method [646] (right). Centroids
with the same color (e.g., red) belong to the same cluster. d Tukey
boxplots of the normalized number of clusters of blood vessel objects
for the desmoplastic growth pattern, the replacement growth pattern, and normal liver. There was a statistically significant difference
between the growth patterns as determined by one-way ANOVA
[F(2,22) = 10.8, p < 0.001]. A post hoc Tukey test showed that the
number of clusters divided by number of vessel objects was significantly different between the desmoplastic growth pattern and the
replacement growth pattern (p < 0.05, ‡), but also between the desmoplastic growth pattern and normal liver (p < 0.001,‡). However,
no difference was found between the replacement growth pattern and
normal liver (p > 0.05). Outliers are plotted as points (·) and extreme
values are plotted as asterisks (*)
vessels, with an LH39-positive basal membrane and weak
or absent αvβ3 integrin, and arising from the alveolar septa
entrapped by the neoplastic cells, were observed [128].
The HGP is not a static phenotype: Systemic treatment of
patients with CRC liver metastases can alter the HGP [119].
The fact that the HGPs can change in response to treatment
reflects the dynamic nature of tumor vascularization driven
by either angiogenesis or vessel co-option. This illustrates
the necessity for longitudinal assessment of vascularization
mechanisms in tumors during treatment and follow-up of
individual patients. In order to achieve this, the histopathological evaluations of the tumor vasculature as described
here must be supplemented by medical imaging techniques
and/or assays, which utilize small biopsies, circulating
tumor cells, endothelial cells, or circulating tumor derivatives. Moreover, the importance of an accurate biomarker
of ongoing sprouting angiogenesis or non-angiogenic vessel
co-option is corroborated by the extensively documented
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and strong link between angiogenesis and immune suppression [129–131] and the surge of clinical studies that
combine anti-VEGF and immunotherapy strategies [132].
In this regard, studies suggested that interactions between
endogenous galectins and glycans may link tumor immunosuppression and angiogenesis, conferring resistance to
anti-VEGF treatment [133].
6.2 Concluding remarks
Although the assessment of MVD has a prognostic value
in many different tumor types, this parameter cannot distinguish angiogenic and non-angiogenic processes of vascularization. However, the histopathological growth patterns
of, for example, lung and liver tumors accurately reflect the
means of vascularization, being sprouting angiogenesis or
non-angiogenic co-option of existing blood vessels. We
therefore propose to determine both the HGP and MVD
when studying the vascularization of tumors. In addition,
noninvasive surrogate markers of the HGP should be developed, for example medical imaging parameters, and these
markers should be integrated in clinical oncology trials.
7 Assessment of intussusceptive
angiogenesis
Angiogenesis is the de novo formation of blood vessels and
can follow either a sprouting or a non-sprouting course.
One important non-sprouting mechanism is intussusception (growth within itself; known also as vascular splitting).
Although both sprouting and intussusception lead to an
expansion of the capillary network, the processes involve
different cellular mechanisms, which are presumably regulated by different molecules. Sprouting angiogenesis (SA)
is characterized mainly by local vasodilatation, increased
vascular permeability, and cell proliferation. It is initiated by
proteolytic degradation of the basement membrane, which
is followed by migration of proliferating EC into the ECM.
The sprouts reorganize internally to form a vascular lumen
and to finally connect to other capillary segments [134].
Intussusceptive angiogenesis proceeds through transluminal tissue pillar formation and subsequent vascular splitting.
The direction taken by the pillars delineates intussusceptive
angiogenesis into its overt variants, namely: (1) intussusceptive microvascular growth (expansion of capillary surface
area), (2) intussusceptive arborization (remodeling of the
disorganized vascular meshwork into the typical tree-like
arrangement), and (3) intussusceptive branching remodeling
(optimization of local vascular branching geometry), including intussusceptive vascular pruning. It has been shown
that intussusceptive angiogenesis takes place not only during physiological processes, but also under pathological
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conditions including tumor growth, colitis, and in neurodegenerative disease, among others [135]. Although the morphological facets of intussusceptive angiogenesis as well as
molecular mechanisms of sprouting are well described, the
exact cellular and molecular mechanism of intussusceptive
angiogenesis is not yet characterized.
7.1 The concept of intussusception
and the methodological challenge
The concept of intussusceptive angiogenesis was first
described within the rapidly expanding pulmonary capillary bed of neonatal rats [136]. Morphological investigations during the last three decades have indicated that the
transluminal pillars arise into the vessel lumen according to
the four consecutive steps in pillar formation postulated by
Burri et al. in 1990: (1) creation of a zone of contact between
the opposing sides of the capillary wall; (2) reorganization of
the junctions between endothelial cells in the contact zone,
resulting in central perforation of the bilayer; (3) formation of an interstitial pillar core, which is invaded first by
the cytoplasmic processes of myofibroblasts and pericytes,
and then by collagen fibrils; and (4) the pillars expand in
girth (≥ 2.5 μm) and transform into a capillary mesh. The
described model with the four consecutive steps is based
mainly on transmission and scanning electron microscopy
methods (scanning or transmission electron microscopy),
and these techniques do not allow dynamical observation of
the pillar formation, fusion, and vascular splitting over time.
7.2 Limitations and challenges
Intussusceptive angiogenesis is spatially quite a complex
process, and it is not possible to document it on isolated sections. To overcome this limitation and maintain high resolution, a 3D reconstruction based on serial ultrathin sections
was reported for the first time in year 2000 [137]. This is a
very laborious approach feasible only in specialized laboratories. Even with significant expertise, only limited numbers
of pillars as a qualitative and descriptive illustration can be
visualized. A new technique is now available that makes
such task much more feasible—serial block face 3D imaging. With this new technique, exact morphological information on relatively big sample volumes (1 × 1 × 1 mm) within
a short time can be acquired: For example, the scanning of
a block the size of one zebrafish embryo would take around
1 day and would provide a 3D scan of the complete embryo
at ultrastructural level. Using the standard approaches, it
would have taken weeks, if not months. Moreover, the outcome from 3View is of much better quality and without loss
of information (slices). This technique enables a quantum
leap forward in 3D morphological studies [138].
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Vascular casting has been extensively used to demonstrate intussusceptive angiogenesis in 3D. The methodology
is based on creating a vascular replica of the microvasculature. As casting media Methylmethacrylate (Mercox, Japan
Vilene) or PU 4ii (VasQtec, Switzerland) is used [137]. A
few hours after perfusion, the organs are excised and transferred to 15% KOH for 2–4 weeks for dissolving the tissue.
After washing, the casts are dehydrated in a graded series of
increasing ethanol concentrations and dried in a vacuum desiccator. They are then examined using SEM. A new powerful methodology employing polymer-based contrast agent in
combination with micro-computed tomography (micro-CT)
has been reported recently [139]. Both casting and micro-CT
are very useful for evaluation of large samples, for example
entire mouse organs and in a subsequent step to “zoom” in
on the area of interest. The vascular casting combined with
scanning electron microscope has the potential to demonstrate greatest morphological details (Fig. 5a). The microCT does not provide a high level of resolution, but is very
rapid and the epitopes are not destroyed and can be used for
further morphological investigations [139].
Dynamic in vivo observation of the pillar formation and
subsequent vascular splitting over time is feasible employing conventional fluorescence microscopy, stereomicroscopy, and laser scanning microscopy (Fig. 5b). The latter
methodologies are widespread, but often cause partially
severe, phototoxic artefacts. Zebrafish lines with uniform
cytosolic expression of the respective fluorescent proteins
such as Tg(Fli1a:eGFP)//Tg(Gata1:dsRed) represent a useful model for dynamic observation of vascular splitting
and blood flow (Circulating erythrocytes). Zebrafish lines
with uniform cytosolic expression of the respective fluorescent proteins can be used, for example Tg(Fli1a:eGFP)//
Tg(Gata1:dsRed). The embryos are mounted in low-melting
agarose in the presence of phenylthiourea and can be monitored for 16–20 h. To reduce phototoxicity, imaging should
be performed with laser scanning microscopes with a long
working distance, using a heated chamber, fast scanning
(less than 1 µs pixel time), lowest possible laser power (in
a range of few percentages depending on the microscope),
and time steps of 10–15 min. Such a dense time pattern is
necessary to record the dynamic cellular alterations of pillar formation and splitting. Using these settings, no signs of
photobleaching or phototoxicity (membrane blebbing, cell
apoptosis) have been observed.
7.3 Concluding remarks
Due to its 3D complexity, intussusceptive angiogenesis
can only be properly investigated by a combined methodological approach. Matching the dynamic cellular changes
in capillaries obtained with in vivo time-lapse studies with
morphological data from light microscopy level up to the
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Fig. 5 Intussusceptive angiogenesis—the methodological chal- ▸
lenge. a Scanning electron microscopy image of transluminar pillars. Early stage is characterized by tinny pillar (arrow), formed
mainly by endothelial protrusions coming from the opposing EC. At
later stages, the pillar (arrowhead) is increasing in girth and its core
is invaded by perivascular cells, fibroblasts, and fibers (visible in the
lower disrupted part of the pillar). Lumen of the vessel is marked
with asterisks. Adapted from [647]. b Dynamic in vivo observation
of the regenerating zebrafish fin vasculature demonstrated a newly
formed pillar (rectangle). c Three-dimensional reconstruction based
on serial semi-thin sections from the same area depicted in b, d transmission electron micrograph demonstrates the transluminal tissue pillars (rectangle in b, c) at ultrastructural level. Black asterisk indicates
the core of the pillar, while arrowhead pointed to cell–cell contacts
between the endothelial cells (EC). Er erythrocyte, Col collagen fibers. Adapted from [648]
ultrastructural level by transmission electron microscope
can provide complex and detailed information (Fig. 5b–d).
The sites of interest should be documented first by intravital microscopy as described above. After the in vivo documentation, the area of interest can be harvested, fixed, and
processed either for paraffin sectioning and LM with immunohistochemistry or alternatively for serial semi-thin sectioning and 3D analyses using, for example, Imaris Software.
To obtain a deeper insight into the morphological substrate
and tissue components involved in the process, the same
tissue should be processed for transmission electron microscope (Fig. 5).
8 In vivo sprouting lymphangiogenic
assay and AAV‑mediated gene transfer
of vascular endothelial growth factor c
(VEGFC)
The lymphatic system plays a key role in the body in maintaining tissue fluid homeostasis, in lipid absorption, and in
immune cell trafficking. In some life-threatening diseases,
such as cancer, lymphangiogenesis, the formation of new
lymphatic vessels, contributes to disease progression. By
contrast, inherited or acquired insufficiency of lymphatic
vessel development results in various forms of lymphedema
[140, 141]. The most important receptor signal transduction
system guiding lymphatic growth is the VEGF-C/VEGFR-3
system. Increasing tissue concentration of VEGF-C by transgenic overexpression or viral gene delivery system leads to
lymphatic overgrowth [142, 143]. By contrast, overexpression of soluble VEGFR-3 extracellular domain leads to
suppression of lymphatic vessel growth in cancer models
[144]. Thus, stimulation or suppression of lymphatic vessel growth in experimental animals and in human patients
can be the ultimate way to improve or inhibit lymphatic
function. In this section, a description is provided of two
in vivo techniques that allow studying the lymphangiogenic
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◂Fig. 6 Stimulation of lymphatic and blood vessel growth in vivo.
a–c Preparation of gelatin sponges for lymphangiogenic assay. a
Small pieces of gelatin sponge are detailed with a puncher before
to be soaked with tumor cells or a compound. Prepared sponges
are inserted in mouse ear between the two skin layers. b, c Immunohistochemical analysis of sponges (b) and sentinel lymph node (c)
resected from mouse with control sponge (PBS or medium without
growth factor) or from mouse with sponge imbedded with growth
factor. Lyve-1 (lymphatic endothelial cell marker) is stained in green,
and CD-31 (blood endothelial cell marker) is stained in red. Sponges
soaked with growth factor showed higher lymphangiogenesis and
angiogenesis compared to control sponge. Scale bar in b, c—250 µm.
d–f Preparation of AAV and transduction of skeletal muscle. d Schematic representation of different VEGF-C and VEGF-D isoforms,
produced by step-by-step proteolysis, and general AAV production
and usage protocol. e Immunohistochemical analysis of t.a. muscle transduced with AAV8 encoding VEGF-C-ΔNΔC or VEGF-DΔNΔC. Tibialis anterior muscles of C57BL/6 J male mice (8 weeks
old) were injected with 109 AAV8 particles in 30 µl of PBS, and the
mice were euthanized 2 weeks later. T.a. muscle samples were isolated and analyzed immunohistochemically for the indicated markers.
HSA human serum albumin. f Analysis of the functionality of lymphatic vessels. Lectin (from Lycopersicon esculentum), conjugated
with FITC (FITC-lectin) was injected to the distal part of t.a. muscle.
After 45 min, the mice were euthanized and t.a. muscle was isolated,
fixed, and stained for Prox1. Lectin is visualized by FITC. Scale bar
in e, f—50 µm
process. One is called the ear sponge assay, and the other
one describes viral gene delivery techniques to achieve the
same goals. Several viral gene delivery vectors are known
to date, such as adenovirus, adeno-associated virus (AAV),
lenti/retrovirus, and baculovirus. Here, we describe as an
example, the use of AAV for the transfer and analysis of
lymphatic vascular growth factors.
8.1 Description of the ear sponge assay and AAV
generation/tissue transduction
In this model, sprouting lymphangiogenesis is reproduced
in a complex in vivo microenvironment. This application
is based on the use of a biomaterial, gelatin sponge that is
inserted in the mouse ear to develop a large lymphatic network [145]. The gelatin sponge is cut into pieces of about
3 mm3, which are transferred in a 96-well plate (one sponge
piece per well) for pre-treatment (Fig. 6a). The sponge can
be soaked with tumor cells, cell-conditioned medium, lymphangiogenic growth factors or inhibitors. Cell suspension
or compound solution (20 µl) is seeded on the top of the
sponge. The plate is incubated at 37 °C, for 30 min, to allow
the diffusion of the solution into the biomaterial. Sponges
are next embedded in a type I collagen solution (7.5 volumes of interstitial type I collagen solution, 1 volume of
10 × Hanks’ balanced salt solution, 1.5 volume of 186 mM
NaHCO3, pH adjusted at 7.4 with a 1 M NaOH solution) and
placed in a new well for 30 min to allow collagen polymerization. For sponge implantation into mice, a small incision
is made in the basal, external, and central parts of the mouse
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ear. Ear skin layers are slowly detached with forceps to make
a hole for sponge insertion, which is then suture shut. Mice
are killed at different time points (2–4 weeks post-sponge
implantation) to evaluate lymphangiogenesis in the sponge
and lymph nodes (Fig. 6a–c).
Preparation of the recombinant AAV encoding VEGF-C
or VEGF-D of mouse origin starts from the cloning of the
ORF into the special AAV plasmid vector, which possesses
flanking sequences 5′ and 3′ Inverted Terminal Repeats
(ITRs) (Fig. 6d). Such vectors can be obtained from commercial sources, for example from Addgene (www.addge
ne.com). Next, the AAV plasmid vector is used to generate
AAV particles by transfecting the plasmid into the host cell
lines derived from embryonic human kidney 293 cells. We
suggest using three-plasmid transfection system, in which
AAV and adenovirus helper genes are administered into
239T cells together with the VEGF-C/VEGF-D encoding
AAV vector plasmid by simultaneous transfection. 293T
cells are seeded at an initial density of 7.7–106 cells/145
cm2plate. When reaching cell confluency level of 70–80%,
cells are transfected with three plasmids (gene encoding one
and the two are the helper plasmids) (1:1:1 molar ratio). The
AAV accumulating inside the cells are released from the
cells by three consecutive freeze/thaw cycles using liquid
nitrogen and + 37 °C water bath, respectively. AAV particles
are then purified and concentrated by ultracentrifugation. It
is important to run a functional test to assess the ability of
the AAV particles to transduce mammalian cells and drive
expression of transgenic proteins. This can be done by transfecting cultured 293T cells and determining the expression
of a transgene by any suitable method, such as Western blotting, ELISA, or fluorescent microscopy in case of fluorescent
proteins.
For the treatment of skeletal muscle (tibialis anterior, t.a.,
in mice), the volume of the AAV solution should be kept to a
minimum and should not exceed 50 µl/t.a. for the adult mice
of ≥ 7–8 weeks old. The AAV dose (concentration of AAV
particles in the injected volume) for a single injection is subject to thorough consideration, taking into account various
factors, such as susceptibility for AAV transduction in a certain tissue, biological activity and toxicity of the transgene,
half-life of the transgenic protein in free circulation or in a
tissue. Gelatin sponge transplantation and injection of AAV
solutions are performed under either isoflurane or liquid
(ketamine/xylazin) anesthesia. Transgene expression starts
soon after the treatment and becomes detectable in 1–2 days,
increases until it reaches the plateau (in 5–7 days), and can
slightly decline later on, but never shuts down completely.
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8.2 Applications, benefits, and strengths
of the assay: analysis of lymphangiogenesis
and angiogenesis
Lymphatic vessels are visualized by immunohistochemical staining using antibodies against, for example LYVE1,
Prox1, or VEGFR3, which are relatively specific for lymphatic EC (Fig. 6b, c, e, f). As the mouse ear displays also
a high blood vessel density, especially after angiogenic
stimulation, angiogenesis can be analyzed by using antiCD-31 antibody. In addition, lymphatic function prior to
and after the AAV treatment can be studied in mice under
anesthesia by injecting a lymphatic tracer [such as FITClectin (FL-1171; Vector Laboratories)] to the distal part of
the muscle as the tracer is taken up by the lymphatic vessels
and carried by lymph flow. The moving front of the tracer
can be observed and recorded and serves as a measurable
parameter of the functional lymphatics. The tracer can be
also visualized by immunohistochemical staining, combined
with simultaneous co-staining for lymphatic endothelial
marker(s) (Fig. 6e, f).
Lymphatic remodeling in sentinel lymph node The ear
sponge assay applied with tumor cells mimics the early steps
of cancer progression and the establishment of lymph node
pre-metastatic niche [146, 147]. The collagen capsule provides a barrier that retains cells in the sponge. Lymphatic
vessels present in the ear drain factors to lymph nodes leading to a remodeling of the lymphatic vasculature in the sentinel lymph node prior to tumor cell arrival (Fig. 1c). Lymph
node metastases are detected between 3 and 4 weeks postimplantation (depending on tumor cell types and the number
of cells used).
AAV gene delivery provides stable transduction Because
of low immunogenicity, AAV itself does not provoke a
strong immune response, in sharp contrast to in vivo transduction with adenovirus, which activates the NF-KB pathway. Unlike AAV, adenovirus gene delivery is accompanied
by a significant immune response to the adenovirus-encoded
proteins, resulting in the shutdown of transgene expression
due to host immune attack. This feature of AAV allows longlasting tissue transduction in mice for up to 2 years. If the
transgene delivered to the mice via an AAV vector encodes
an isogenic murine protein, it does not provoke an immune
response either. Although the purified transgenic proteins
can also be used for in vivo treatment, their administration
into the tissues usually produces an inferior effect due to the
instability and fast dilution of the injected protein when it
diffuses out from the site of injection. It should be noted that
lymphangiogenic and angiogenic proteins must be present
for a prolonged period of time to allow vessel formation
and maturation of the newly formed vessels. Immature vessels often formed after adenovirus-mediated gene delivery
are unstable and prone to regress soon after the transgene
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expression ceases [148]. Multiple injections of the protein
to the same tissue are not desirable due to the trauma caused
by the injections.
Analysis of various forms of lymphangiogenic factors
Various forms of VEGF-C or VEGF-D can be used including immature (full-length) pro-VEGF-C/VEGF-D and
mature VEGF-C/VEGF-D-ΔNΔC forms, which are naturally produced during step-by-step proteolysis (Fig. 6d).
Only mature forms VEGF-C/VEGF-D-ΔNΔC demonstrate
full lymphangiogenic activity in vitro and in vivo, primarily
due to higher affinity interactions with VEGFR-3. In nature,
they are all expressed as immature pro-VEGF-C/VEGF-D,
which then undergo step-by-step proteolysis producing fully
active mature ΔNΔC forms. Thus, AAV-mediated expression of ΔNΔC form of VEGF-C or VEGF-D provides somewhat artificial stimulatory conditions for lymphatic vessel
growth.
Lymphangiogenic factors can also stimulate blood vessel
growth The mature forms of both VEGF-C and VEGF-D
are able to bind to and stimulate not only VEGFR-3, but
also VEGFR-2, although the affinity toward VEGFR-2 is
significantly lower than VEGF-A (by a factor of 20–100).
Blood vessel growth is induced when AAV-VEGF-C or
AAV-VEGF-D is administered to the murine skeletal muscle
[143], which has also been demonstrated in rabbit skeletal
muscle transduced by recombinant adenoviruses [149]. The
angiogenic effect of VEGF-C, however, can be decreased,
while preserving full lymphangiogenic activity, by using
a VEGF-C156S mutant form of the growth factor [150];
VEGF-C156S has a reduced binding affinity for VEGFR-2,
while binding affinity to VEGFR-3 is largely unaltered.
8.3 Limitations and challenges
Sponge preparation Steps related to sponge preparation
are critical for the reproducibility of the assay. The addition of a drop of 20 µl with cells or factors on the top of
the sponge is better than an immersion of the sponge in the
solution. Moreover, it is important to turn the sponge halfway through the incubation period to allow homogenous
diffusion in the biomaterial. Finally, the collagen capsule
formation is a key step to allow sponge insertion between
ear skin layers without cell or solution loss.
Control of transgene expression To date, broad usage
of AAV-mediated overexpression of VEGF-C/VEGF-D
(or VEGFR-3-Fc) in mice or in human patients in clinical
practice is hindered by the inability to efficiently regulate
transgene expression. When a lymphangiogenic effect is
achieved, it would be desirable to shut off the expression
of the transgene. Long-term overexpression of VEGF-C/
VEGF-D would lead to a heavily overgrown and unstructured lymphatic vasculature, which would be poorly
functional. Blood vessels may also respond to long-term,
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uncontrolled VEGF-C/VEGF-D overexpression by overgrowth and increased leakiness. Currently existing ON/OFF
switch systems, such as under tetracycline regulation, are
either inefficient, or toxic for humans, although some have
been adapted for in vitro use. The versatility of the CRISPR/
Cas9 delivery system may provide solutions to these problems in the near future.
Unequal transduction of target cells in the transduced
tissue Unlike in transgenic animals, where transgene expression can be induced in virtually every cell in a target tissue
such as muscle, viral gene delivery fails in some of the target
cells and this can lead to tissue mosaicism and non-homogenous protein delivery. Cell types vary in their sensitivities
to AAV transduction due to differential expression of AAVbinding receptors. For example, tail vein injection of AAV9
results in a significantly higher transduction of heart than of
the liver or pancreas [151].
Local trauma associated with ear sponge implantation
and viral vector delivery During surgery, the integrity of
the ear must be assessed as perforations can result in sponge
loss. Of note, it is not possible to identify mice using ear
clips since both ears are used for the assay. Depending on
the pathological context of the study, a limitation of the ear
sponge model is local inflammation induced by the sponge
insertion. Regarding viral vector delivery, when AAV is
administered locally, the injection itself produces some
trauma in the target tissue, which can lead to altered sensitivity of cells along the vector injection path.
8.4 Concluding remarks
The ear sponge assay is a suitable model to study lymphatic
vessel remodeling in the both primary tumor and sentinel
lymph node prior (pre-metastatic state) and after tumor cell
arrival (metastatic state). Gene delivery via AAV represents
a reliable and relatively inexpensive approach for gene therapy in inherited or acquired vascular disease models. AAV
provides a convenient way to study biological properties of
bioactive proteins, such as vascular growth factors in vivo
and under conditions mimicking the natural expression of
the endogenous genes.
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(Fig. 7). In either system, pericyte recruitment to EC tubes
can be readily observed over a 72–120 h period and a consequence of this recruitment is deposition of the vascular basement membrane between the abluminally recruited pericytes
and the ECs that line the branched networks of tubes [153].
Interestingly, EC tube formation results in the creation of
networks of ECM spaces, which are termed vascular guidance tunnels [158], where the EC tubes reside, and are also
a physical space into which pericytes are recruited along the
basal and abluminal EC tube surface [153, 158]. Both EC
and pericytes migrate along each other in a polarized manner within these tunnels, resulting in basement membrane
deposition underneath the abluminal surface of the EC-lined
tubes [152, 153, 159]. Basement membrane deposition leads
to much narrower and elongated EC tubes in EC-pericyte
co-cultures compared to EC-only tubes which progressively
get wider and less elongated with time [153, 159].
9.1 Benefits and strengths of the assay
The major benefit and strength of this assay model is its high
reproducibility and high replicate number in that it is performed in half-area 96-well plates. The assay is typically set
up in 60 wells per plate, and this can be readily established
using one T75 flask of human EC (e.g., HUVEC) and one
T25 flask of human pericytes (brain-derived vascular pericytes) (labeled with GFP). The assay allows performance
of siRNA suppression experiments using either cell type
or lysates that can be prepared to assess signaling transduction cascades by Western blot. Also, the recruitment of
pericytes can be examined in time with or without blocking
antibodies to various targets (e.g., growth factors, receptors, integrins, ECM molecules, etc.) or signal transduction
inhibitors. The consequence of pericyte recruitment to EC
tubes can be revealed by examining basement membrane
deposition (by immunofluorescence staining or transmission
electron microscopy) (Fig. 7), or by measuring capillary tube
widths and lengths. Finally, real-time video analysis can be
used to monitor pericyte and endothelial motility as they coassemble to form tube networks with abluminally recruited
pericytes in 3D matrices.
9.2 Assay overview
9 Assay for pericyte recruitment
to endothelial cell‑lined tubes, capillary
assembly, and maturation
A fundamental question concerns the growth factor and signaling requirements governing how human EC and pericytes
co-assemble to form capillary tubes in a 3D matrix or tissue environment [80, 152–155]. To this end, bioassays have
been developed in 3D collagen or fibrin matrices to address
such issues using serum-free defined conditions [156, 157]
HUVEC or human artery endothelial cells (HUAECs)
and human brain-derived vascular pericytes or bovine
retinal pericytes are trypsinized and seeded within 3D collagen matrices at a density of 2 × 106 cells/ml for EC and
4 × 105 cells/ml for pericytes. Both types of EC or pericytes
work very well in this assay system. FGF-2 and stromalderived factor-1α (SDF-1α) are each added at 200 ng/ml
into the collagen gel mix (2.5 mg/ml of rat collagen type
I). Twenty-five μl of gel is added into each individual 96
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Fig. 7 Serum-free defined model of human endothelial cell-pericyte
tube co-assembly in 3D collagen matrices. a Human EC and GFPlabeled pericytes (Peri) were seeded together at the indicated cell
densities and after 120 h, were fixed and stained with anti-CD31
antibodies. The immunostained cultures were imaged using confocal microscopy. The serum-free defined culture system contains SCF,
IL-3, SDF-1α, FGF-2 and insulin (a component of the RS II supple-
ment), which are required to be added in combination. b Cultures
fixed at 120 h were immunostained with antibodies to CD31, laminin
(LM), and fibronectin (FN) and were imaged using confocal microscopy or were examined by transmission electron microscopy. Arrows
indicate the capillary basement membrane. L indicates lumen while
Nuc indicates nuclear labeling. Bar equals 25 μm
well (A/2 plates from Costar). After 30 min of equilibration
in a CO2 incubator, culture medium is added (100 μl per
well) which is Medium 199 (100 μl per well supplemented
with 1:250 dilution of Reduced Serum Supplement II) [160],
and 40 ng/ml each of FGF-2, stem cell factor (SCF), interleukin-3 (IL-3), and 50 μg/ml of ascorbic acid are added.
Cultures are then placed in a CO2 incubator for 72–120 h.
After this time, cultures are fixed with 3% paraformaldehyde
in PBS (120 μl per well). Cultures can then be imaged under
fluorescence to quantify EC tube area and pericyte recruitment under the various conditions of treatment. Individual
cultures can be immunostained with various antibodies
directed to EC or pericyte cell surfaces and/or ECM antigens (i.e., basement membrane proteins such as fibronectin,
laminin, and collagen type IV) [153]. Alternatively, cultures
can be used to make lysates using SDS sample buffer or
used to isolate total RNA to perform gene expression experiments [153]. Real-time movies can be performed to examine
pericyte recruitment and measure pericyte motility during
capillary assembly and maturation [154]. In the latter case,
nuclear GFP-labeled pericytes can be utilized to track pericyte motility [154].
50 mg of heparin, and 100 ml of fetal calf serum (then sterile filter complete media). Tissue culture flasks are coated
with 1 mg/ml of gelatin solution for at least 30 min. prior to
seeding the trypsinized EC. Pericytes are grown using the
same culture media and gelatin-coated flasks. Bovine hypothalamic extracts as well as rat tail collagen preps are used
at 7.1 mg/ml protein in 0.1% acetic acid. The assay system
is highly robust and reproducible [153, 156, 157].
Especially for longer-term experiments (i.e., multiple
days), it is important to completely surround each well with
100 μl of water. This reduces dehydration of media in the
microwells and maintains the health of the cultures. For
cultures extending longer than 72 h, the cultures are replenished by removing 90 μl of media and replacing with 100 μl
of media per well. This medium is slightly different than
that used above to start the assay: Medium 199 containing
a 1:250 dilution of RS II, ascorbic acid at 50 μg/ml, as well
as 80 ng/ml of FGF-2, and 40 ng/ml each of SCF, IL-3, and
SDF-1α.
9.3 Critical steps in the assay
The assay system above is highly dependent on the health
of the growing EC. To each 500 ml bottle of Medium 199,
we add 200 mg of lyophilized bovine hypothalamic extract,
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9.4 Limitations and challenges
The major limitations of this in vitro assay relate to how
reflective the assay model is to the in vivo state. Overall,
the data and conclusions obtained over the years have
been repeatedly demonstrated to be observed using mouse,
zebrafish, or quail in vivo models [155]. One of the interesting issues is the state of differentiation of the EC that are
Angiogenesis (2018) 21:425–532
used in these in vitro assays, and how similar are they to EC
in vivo that are undergoing vasculogenesis or angiogenic
sprouting. This is a very difficult question to answer and
needs to be addressed in future studies. Another potential
weakness of the current in vitro systems is that flow conditions have to be adapted or the gels have to be transplanted
into a mouse model to interface the human vessels with an
in vivo vascular system.
9.5 Concluding remarks
Here, a defined assay system is described which allows for
the assembly of capillary tube networks, with EC-lined tubes
and abluminally recruited pericytes in 3D collagen or fibrin
matrices. This system has been previously utilized (and can
be employed in future studies) for detailed investigations of
the molecular and signaling requirements for this process,
by independently assessing the functional role of ECs vs.
pericytes during these tube co-assembly events.
10 EC co‑culture spheroids
Angiogenesis, the sprouting of endothelial cell capillaries
from preexisting blood vessels, is multifactorial including processes such as proliferation and directed migration.
These processes in turn are influenced not only by autocrine but also by paracrine stimuli mediated by, for example,
pericytes, fibroblasts, immune cells, and tumor cells in the
case of cancer. This intercellular communication by stimuli
includes growth factors, chemo- and cytokines as well as
other secreted factors, like enzymes, extracellular basement membrane components, and the exchange of genetic
material (miRNA, mRNA, DNA) via extracellular vesicles
[161–163]. Anatomically, these various cell types are finetuned and orchestrated to become organized 3D structures
to ultimately allow and control the flow of blood. However,
the field of angiogenesis has typically focused on studying
angiogenesis-associated processes in vitro in pure EC cultures. Although studying EC in isolation has some advantages in investigating certain EC-specific elements and to
certain defined processes within angiogenesis, 3D co-culture
spheroids allow one to study the direct intercellular cues
and cross talk between EC and their surrounding cells. Cocultures with tumor cells create a simplified dynamic tumor
microenvironment due to the changes in shape and size in
time. As a result, the 3D spheroid assay sheds light on the
biological conduct and communication of EC with a multitude of different cell types under various stresses or treatment conditions in a controlled manner. Importantly, it offers
the possibility to investigate multiple different interventions
(e.g., chemotherapies, radiation, experimental drugs) in a
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medium-throughput manner in an environment more relevant to the in vivo situation.
10.1 Strengths and benefits
of the three‑dimensional co‑culture EC
spheroids
Studies on in vitro 2D monolayer cell cultures and their
translation/extension to clinical settings have their limitations, as they are not capable of mimicking the nutrient
and oxygen gradient and an environment reminiscent of
the in vivo setting. The preclinical 3D co-culture spheroid
assay facilitates intra-/intercellular cross talk mimicking
aspects of in vivo vascular architecture, for example lumen
formation and polarization, and is particularly valuable for
interrogating these interactions in the presence of two or
more cell types growing together with natural adhesion and
layering properties. Additionally, and at least as important,
it lends itself for controlled experimental manipulation and
replication thereof as it can be applied in the developmental/regeneration field as well as the cancer biology and the
cancer treatment fields. Particularly in the cancer field, the
3D co-culture tumor cell/EC spheroid assay is a valuable
instrument for interrogating interactions between the tumor
stroma and parenchyma to better understand the mechanisms
of radiation and cancer therapeutics and how cells support or
damage each other in their response to these interventions.
Ultimately, the ability to screen a relatively high number
of conditions against these mini-tumor microenvironments
has potential to promote the establishment of improved drug
pharmacokinetics, efficacy, and safety profiles [164].
Inherent in many in vitro assays, one of the strengths of
the spheroid assay is the relative ease of focused manipulation to aid in answering specific hypotheses. Namely, prior
to the paired co-culture one of the cell types can be genetically altered by retro- or lenti-viral constructs, including
fluorescence (gain-of-function) or by shRNA/siRNA knockdown or CRISPR/Cas9 (loss-of-function). Additionally, the
sensitivity or resistance to a particular pharmacological or
radiation treatment approach can be tested. We and others
have observed marked differences in treatment response and
viability of tumor cells growing in the presence/in contact
with EC and vice versa [164].
10.2 Assay overview and types of assays
Multicellular 3D spheroids can be generated by various
methods, including the (1) spontaneous spheroid formation using (ultra) low binding plates; (2) “hanging drop”
technique; (3) suspension cultures (e.g., by spinner flasks
or bioreactors); (4) scaffold-based models (e.g., hydrogels),
and (5) magnetic levitation [165]. Since spontaneous spheroid formation and the hanging drop method are arguably
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the most cost-effective and likely take the least amount of
specialized skill set, we will focus on these two iterations
of the model (Fig. 8). One of the pioneering approaches to
generate self-assembling multicellular 3D tumor endothelial
spheroids in hanging drops was by Timmins et al. [166].
Subsequently, this assay was validated by various groups
and expanded on in terms of cell types tested and it evolved
as an instrument to be transplanted into animals, devoid of
Matrigel, to study cancer progression and ultimately metastasis [164].
The initial spheroid structure is produced by pipetting a
single-cell suspension of 1000 tumor cells in a 20-μl droplet
on the inside of the lid of a, for example, 48-well cell culture
plate (Greiner Cellstar, BioExpress, Kaysville, UT). Gravityenforced self-aggregation is facilitated by inverting the lid
and placing it on its receptacle (i.e., the 48-well plate). The
culture medium and incubation conditions do not need to be
adjusted and thus can stay as preferred. After 72 h, the lids
are set inside up and the addition of 2000 EC in a volume
of 5 μl is added to the existing culture. The lid is then reinverted, and the hanging drops can be incubated and monitored for growth, cell type incorporation characteristics, and
other aspects for an additional 14 days depending on the cell
types used (Fig. 8).
Additionally, the spheroids can be implanted in vivo and
used in animal models or within specialized techniques
such as the dorsal skin fold window chamber, creating an
in vitro/in vivo assay. For decades, it has been postulated that
growing tumor cell-only spheroids and implanting them in
animals lead to a much higher tumor take with far fewer total
cells inoculated. This was hypothesized to be due to the creation of a microenvironment in the spheroid as well as possibly engaging or enriching for cancer stem cell phenotype.
Although the optimal conditions vary when co-culturing different cell types with EC, when using tumor-EC spheroids,
as few as two spheroids per implantation are sufficient to
generate tumor growth and metastasis [164]. This approach
may be valuable in studying various treatments or aspects of
tumor progression thought to be dependent on vascularization and/or stimulation of metastasis or aggressiveness via
tumor cells engaging EC.
The spheroids can be assessed by multiple readouts
depending on the laboratories preferences, techniques
available, and skill set. Namely, quantitative and qualitative
evaluations include imaging techniques such as bright field
and fluorescence microscopy, molecular marker assessment
by, for example, immunohistochemistry or immunofluorescence, or functional assays such as cell viability and survival assessment with or without growth factors, biologics,
therapeutics, or radiotherapy [164]. The functional assays
are generally performed after transferring the spheroids into
low-binding 96-well plates to better control and standardize
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the additions of, for example, growth factors, therapeutics,
or reporters (e.g., viability or proliferation stains; Fig. 8).
10.3 Limitations and challenges
Although in theory this assay can be used for multiple different co-cultures with a number of cell types, some restrictions
arise in the use of particular cell types as well as the combinations thereof. Namely, primary isolates have a limited
duplication potential and life span, but this can be bypassed
with immortalized cells. Relatedly, cell density needs to be
carefully controlled based on the cell type (short vs. long
doubling times) and the planned length of the assay. A major
challenge in pairing cells in co-culture growth is to find a
growth medium that will supply each cell type with enough
nutrients to survive, but that also does not overstimulate
proliferation. Allowing cell types to interact and support
themselves by the mere process of adhesion and exchange
of various factors is one of the more exciting aspects of this
model and frequently has led to new insight in the absence
of the high serum, high nutrient mediums used in isolated
2D cultures of the past. Depending on the chosen setup of
cells (e.g., primary vs. immortalized) and pairing of the cell
types (e.g., endothelial and pericytes, stroma and (tumor)
parenchyma, same species or mouse/rat vs. human) interpretation and generalization of the results must take these
outsets in consideration.
10.4 Concluding remarks
In summary, the EC spheroid assay is an in vitro and possibly an in vitro/in vivo co-culture method assisting in the
comprehension of initial stages of angiogenesis and angiogenesis-associated processes in a continuum—from initiation to development and progression. The system allows for
monitoring tissue development as well as testing pro- and
anti-angiogenesis targeted treatments with and without the
combination of chemo/radiation therapy in a more relevant
microenvironment than traditional 2D cultures. Thus, this
in vitro/in vivo preclinical cell and animal model holds
promise to not only enable the identification of authentic
and novel biomarkers but also provide enhanced predictive
utility for drug development and discovery.
11 Endothelial cell metabolism
Metabolism is a biochemical reaction network that catabolizes nutrients into metabolic products such as energy and
biomass that are needed to survive, proliferate, and adapt to
environmental challenges. As such, they provide a functional
readout of the observed phenotypic changes. Metabolomics
is the systematic study of the aforementioned metabolic
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Fig. 8 EC co-culture spheroid assay. a Overview of the procedure
of the hanging drop spheroid growth assay over time. Prior to initiating the co-culture, tumor cells (green) are allowed to establish initial micro-spheroids by themselves. Typically, after 72 h EC can be
introduced (red) and their incorporation monitored over time up to
14 days. b After 7 days, the tumor-EC spheroid can be quantitatively
and qualitatively evaluated by various techniques, such as c (fluorescence) microscopy, histology, functional assays or implanted in vivo.
Breast tumor cells (green; GFP-4T1) are combined with 2H11 EC
(red) stained with MitoTracker Red. Scale bar = 100 µm
products and is currently enjoying a revival (“Metabolomics,
the apogee of the omics trilogy” [167]). The ultimate goal of
metabolomics is to understand: (1) how cells under different
circumstances apply and reroute their metabolic pathways,
and (2) what are the required start and end products in relation to the observed phenotype.
The majority of EC remains quiescent for years [168], but
upon exposure to pro-angiogenic stimuli (such as VEGF),
they can rapidly switch to a proliferative and migratory state
in health and disease. At the base of this switch lies a tightly
regulated metabolic network, providing the necessary energy
and building blocks for EC to respond accordingly [169,
170]. ECs rewire their metabolism in a highly specific manner to optimally execute these functions [171]. Indeed, ECs
are highly glycolytic, producing large amounts of ATP and
lactate even in the presence of ample oxygen [170].
In the following section, we will discuss three approaches
that have been established to monitor metabolic activities of
EC: (1) tracer metabolomics, (2) radioactive tracer-based
metabolic assays, and (3) measurements of extracellular
acidification rate and mitochondrial respiration by the Seahorse XF analyzer. These approaches are complementary,
and it is recommended to perform these assays in parallel
for a more complete metabolic phenotyping.
be due to increased uptake but also by de novo synthesis
using 3-phosphoglycerate from glycolysis as a substrate. As
such, the majority of metabolites can be linked to multiple
sources and by merely measuring the (relative) abundance
of metabolites, one cannot readily deduct the responsible
pathways and their active interplay. This knowledge is paramount to understanding cellular biology. A specialized field
in metabolomics, Tracer Metabolomics [172], tackles this
challenge and uses labeled non-radioactive substrates (carrying 13C, 15N, 18O, 2H, etc.) to monitor the distribution of
the labeled isotopes throughout the metabolic network and
as such, provides the actual contribution of nutrients to specific metabolites and consequently the relative activity of
metabolic pathways.
11.1 Tracer metabolomics
The interpretation of metabolic activities based on the
(relative) abundances of metabolites under specific conditions remains a formidable challenge, largely because the
network of biochemical reactions is highly interconnected.
For instance, an increase of intracellular serine levels can
11.2 Experimental tracer metabolomics setup of EC
A possible cell culture format for EC is performed using
a 6-well plate, in which at least 150,000 ECs are plated.
ECs need to be grown in customized medium of which the
substrate of choice is withdrawn and replaced by its nonradioactive isotopically labeled counterpart (tracer). The
majority of tracers are nutrients that are taken up by the
cell (for instance, glucose, glutamine, palmitic acid, etc.),
of which one of the constituent atom species (12C, 14N, 1H,
16
O, etc.) is replaced by its non-radioactive isotopologue
(respectively, 13C, 15N, 2H, 18O, etc.). ECs take these labeled
nutrients up and process them similarly compared to their
unlabeled form. Consequently, all of the downstream metabolites and metabolic pathways linked to the specific tracer
will incorporate the labeled atoms. It is critical that ECs
reach isotopic steady state, a condition that is accomplished
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when the labeled isotope enrichment in a given metabolite
is stable over time relative to the experimental error [172].
Practically, steady state in EC is reached in 24–48 h using,
respectively, glucose or glutamine tracers [173, 174], for
palmitic acid up to 72 h of incubation is required [169]. The
difference in labeling time necessary to reach isotopic steady
state depends on both the fluxes (i.e., rate of conversion)
from the nutrient to the metabolite of interest, and the concentration of that metabolite and all intermediate metabolites
[172]. The fractional contribution, which is the fraction of a
metabolite’s carbon (or other atom) produced from a certain
nutrient, in the downstream metabolites as well as their isotopic envelopes (isotopologues) can be quantified using mass
spectrometry (MS). Prior to MS analysis, ECs are washed
using a physiological (0.9%) NaCl solution and metabolites
are extracted in an organic (50% methanol–30% acetonitrile–20% H2O) extraction buffer. It is of utmost importance
that the extraction is carried out swiftly because a cell’s
metabolism changes within the order of seconds to minutes
[175]. Extracts are centrifuged to remove the precipitated
proteins and insolubilities. The supernatant, containing the
polar metabolites, is loaded onto the MS platform.
11.3 Mass spectrometry
The instrument of choice for carrying out (tracer) metabolomics is mass spectrometry (MS), mainly due to its superior
sensitivity and speed of analysis in comparison with other
technologies such as nuclear magnetic resonance (NMR)
spectroscopy, as well as its flexibility in applying different
kinds of chromatography (liquid or gas chromatography,
LC, or GC) to study different chemical classes of metabolites [176]. We will only focus on the analysis of polar
metabolites (amino acids, organic acids, hexose phosphates,
etc.). Apolar species such as lipids are beyond the scope
of this section. A crucial requirement for the MS in tracer
metabolomics relates to its resolution, the ability to accurately quantify the isotopologue profiles of the metabolites
of interest. As such, the current benchtop LC–MS OrbiTRAPs are among the instruments of choice to conduct tracer
metabolomics. Alternatively, GC-linked (triple) quadrupole
MS can be used as well. For the separation of polar metabolites prior to the MS, two main types of chromatography can
be applied: (1) hydrophilic liquid chromatography (HILIC)
[177] and (2) ion-pairing chromatography [178]. The latter
has superior binding and separation capacities compared to
HILIC, but it restricts any further usage of the chromatograph to ion-pairing mode. In this perspective, HILIC provides a better solution, despite the lesser separation capacity
compared to the ion-pairing setup as it comes without any
restrictions to apply different chromatography settings on
the same platform.
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11.4 Data processing
Data extraction of isotopic profiles occurs via vendor-specific software or in-house developed software platforms
(e.g., BIOMEX™) [179]. In general, the workflow starts
with in-house libraries containing information on the exact
mass of the target molecule and its retention time during
the chromatographic separation to identify the metabolite
of interest. Next, the abundances of the isotopes of different
metabolites of interest are quantified. Following a correction for natural abundances [172, 180] to facilitate proper
interpretation of the labeling data, the fractional contribution
is calculated [172].
11.5 Radioactive tracer‑based assays
As introduced above, unlabeled or labeled metabolites can
be quantified by MS or NMR. However, these techniques
display some limitations, as they do not quantify the absolute flux, which reflects the rate of turnover of molecules
through a metabolic pathway. Radioactive isotope labeling
is a widely used experimental approach to analyze metabolic
flux. This technique is based on the principle that nutrients
labeled with radioactive tracers (e.g., 3H and 14C) at specific
positions within the molecule release the radioactive label
as 3H2O or 14CO2, respectively, upon specific enzymatic
activities. Consequently, the generated 3H2O and 14CO2
diffuse into the medium and can be measured by trapping
its release on soaked filter paper and subsequently quantified by scintillation counting [170]. As an example, the use
of [5-3H]-glucose enables to quantify glycolytic flux [181]
because the single tritium at the C5 of glucose is removed
by a condensation reaction catalyzed by enolase (Fig. 9a),
hence releasing radioactive 3H2O in the medium. Similarly,
by using [9,10–3H]-palmitic acid, one can measure the radioactive 3H2O formed during fatty acid oxidation [169].
In order to map the different metabolic fates of glucose
and other crucial substrates, a set of differently radioactivelabeled tracers are needed to identify and quantify the ongoing biochemistry. As an example, in parallel with the glycolytic flux measured by [5–3H]-glucose, the downstream
oxidation of glucose can be monitored by quantifying the
release of 14CO2 by [6–14C] glucose or uniformly labeled
[14C6] glucose. Notably, by using [6–14C] glucose, information on the oxidation of glucose in the tricarboxylic acid
(TCA) cycle is obtained, whereas the [14C6] glucose tracer
generates 14CO2 via two different metabolic pathways: the
oxidative pentose phosphate pathway (oxPPP) and the TCA.
Also, the carbon at position 1 in glucose is liberated in the
oxPPP. Hence, using [1–14C] glucose in parallel with [6–14C]
glucose, the oxPPP flux can be estimated by subtracting the
rate of [6–14C] glucose from the one of [1–14C] glucose. In
summary, assessing the choice of the tracer and its specific
Angiogenesis (2018) 21:425–532
labeling positions is essential in terms of the biological
question that needs to be addressed. Similar approaches are
available to investigate other “non-glucose” related fluxes,
for example by using [14C5] labeled glutamine to determine
glutamine oxidation.
11.6 Measurements of extracellular acidification
rate and mitochondrial respiration using
a Seahorse XF analyzer
The Seahorse extracellular flux (XF) analyzer enables to
assess different aspects of cell metabolism. Apart from
providing an indirect readout of anaerobic glycolysis
through measurements of the extracellular acidification
rate (ECAR) [182], it can also assess oxidative phosphorylation (OXPHOS) (through oxygen consumption rate
(OCR)). A commonly used method to determine glycolysis
by the Seahorse XF analyzer is the measurement of acidification of the medium (pH changes) in a glycolysis stress
test [183]. In this technique, glucose, oligomycin (inhibitor of ATP synthase in the electron transport chain (ETC)),
and 2-deoxy-glucose (2-DG, inhibitor of hexokinase) are
sequentially administered while measurements of ECAR
are being performed. In the first phase of the experiment,
ECs are cultured in medium deprived of glucose or pyruvate
(the glycolysis stress medium). Supplementation of glucose
feeds glycolysis, which allows estimation of glycolytic rate
by calculating the difference between ECAR before and
after addition of glucose. ECAR, prior to glucose injection,
is referred to as non-glycolytic acidification resulting from
other sources of extracellular acidification not attributed to
glycolysis (Fig. 9b). Next, oligomycin, which inhibits mitochondrial ATP production, lowers the ATP/ADP ratio, and
shifts energy production to glycolysis, is supplemented to
measure the maximum glycolytic capacity. The difference
between ECAR before and after oligomycin supplementation is a measure of the glycolytic reserve capacity. The last
injection of 2-deoxy-glucose (2-DG) inhibits glycolysis, and
the resulting decrease in ECAR confirms that the ECAR
produced in the experiment is due to glycolysis. Moreover,
inhibition of glycolysis by 2-DG also provides information
on non-glycolytic ECAR of cells (Fig. 9b).
Mitochondrial respiration can be assessed by using a
modified Seahorse Cell Mito Stress Test [184]. This test
sequentially uses modulators of respiration that target different components of the ETC. Oligomycin, carbonyl cyanide 4-(trifluoromethoxy) phenyl-hydrazone (maximizing
OCR by uncoupling the OXPHOS), and antimycin (blocking OXPHOS by inhibiting complex III) are sequentially
injected to determine basal mitochondrial OCR (OCRBAS),
ATP-dependent OCR (OCR ATP), maximal respiration
(OCRMAX) and non-mitochondrial respiration (OCRnon-mito),
proton leak, and spare respiration (Fig. 9c) [184].
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11.7 Limitations and challenges, problems
and pitfalls in probing EC metabolism
Tracer Metabolomics provides information on relative pathway activities, qualitative changes in pathway contributions
via alternative metabolic routes, and nutrient contribution to
the production of different metabolites. In this perspective,
a limitation of tracer metabolomics is the lack of quantitatively estimating metabolic fluxes, which provide additional
quantitative and complementary information such as pathway directionality. Nonetheless, resolving metabolic fluxes
is time and data intensive [185] and often requires applying
computational modeling of the data. A challenge for tracer
analyses is the requirement of expert knowledge at the level
of mass spectrometry and biochemistry. This relates to the
choice of specific tracers needed to answer biological questions as well as the tools (chromatography, mass spectrometry, software) available to extract and process the required
data. The optimal cell culture experiments apply customized medium of which the unlabeled form of the tracer has
been withdrawn. This setup maximizes the amount of label
entering the cell and increases the likelihood to identify
and quantify the labeled “downstream” targets. A pitfall
occurs when the unlabeled form cannot be withdrawn; in
this case, the labeled compound can be added in equimolar
quantities to the medium as the unlabeled compound. This
poses potential problems: (1) The amount of label incorporated in downstream metabolites is reduced and pathways
using low amounts of the tracer might be overlooked; and
(2) doubling the concentration of a specific substrate can
induce metabolic changes into the cellular biochemistry, for
instance doubling the glucose levels from the physiological 5.5–11 mM places EC in a diabetic-like “high glucose”
environment, triggering additional metabolic changes or, in
worst case, inducing toxicity.
Not all of the metabolites present in the cell are detectable by MS, the major causes for this relate to the abundance
and/or intrinsic technical challenges, such as the ionization
potential of the metabolite of interest. To tackle this challenge and still gain information on this missing piece of
the puzzle, one needs to look at the fractional contribution
and isotopologue profiles of neighboring metabolites that
reflect the labeling pattern of the missing component, for
instance the lack of oxaloacetate detection can be accounted
for by looking at the isotopologue profile of aspartic acid.
Consequently, engaging on tracer analyses implies that sufficient metabolites need to be taken into account ensuring
full coverage of the pathway(s) of interest.
General limitations of (radioactive) tracer-based assays
Cell metabolism is a very dynamic process, and the metabolic needs as well as metabolic activity of EC have been
shown to correlate with the cell cycle phases [171, 186].
Therefore, one should carefully control cell density while
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Fig. 9 Methods to measure EC metabolism using radioactive tracers
and Seahorse XF analyzer. a Schematic representation of glycolytic
flux measurements with [5–3H]-glucose. A single tritium present on
5C glucose is released as water in the ninth step of glycolysis catalyzed by enolase. b Schematic representation of the modified Glycolysis Stress Test. The first measurements of ECAR are performed
while ECs are incubated in glycolysis stress test medium (without
glucose and pyruvate). The injection of glucose leads to the saturation of glucose concentration and allows measuring glycolytic rate
(blue). Second, injection of oligomycin blocks oxidative ATP production and shifts the energy production to glycolysis, with the subsequent increase in ECAR revealing the cellular maximum glycolytic
capacity (green). The difference between glycolytic capacity and glycolysis rate defines glycolytic reserve. The final injection of 2-deoxyl-
glucose (2-DG) inhibits glycolysis, and the resulting decrease in
ECAR confirms that the ECAR produced in the experiment is due
to glycolysis. ECAR, prior to glucose injection or after 2-DG injection, is referred to as non-glycolytic acidification (pink). c Schematic
representation of the modified Seahorse Cell Mito Stress Test. First
injection of oligomycin blocks ATP synthase and allows the calculation of the ATP coupled oxygen consumption rate (OCRATP; red).
Second, injection of FCCP maximizes the OCR by uncoupling the
OXPHOS, enabling to calculate the spare respiration (reserve capacity). Third, antimycin-A treatment blocks complex III of ETC enables the calculation of the basal mitochondrial respiration (OCRBAS;
green), the maximal mitochondrial OCR (OCRMAX; orange), the proton leakage (blue), and the non-mitochondrial OCR (pink)
performing these experiments. Moreover, the results and
interpretation of metabolic studies using radioactive tracers
depend on appropriate normalization of the data, as ineffective or poorly chosen normalization methods can lead
to erroneous conclusions. Therefore, normalization to total
protein concentration, cell number, or DNA content is considered as the golden standard in metabolomics.
Seahorse XF analyzer. Although assays performed on a
Seahorse XF analyzer requires only small number of cells
(30,000 cells/well in 24-well assay plate format), they still
present several limitations that one should take into consideration while performing these experiments. Additional
metabolic processes, like CO2 generated during tricarboxylic acid (TCA) cycle activity, can change the pH of the
media and interfere with the readout and interpretation of
the results. Since ECAR is essentially a measurement of pH,
buffering agents (e.g., sodium bicarbonate) are not included
in the assay medium. Moreover, as bicarbonate and media
pH play a role in regulating glycolysis, they can influence
and confound measurements of ECAR. In order to increase
the accuracy of the performed experiments, application
or pre-treatment with additional chemical inhibitors (e.g.,
AR-15585; lactate export blockers) can be of use to determine whether changes in pH come from lactate excretion or
other sources of media acidification. Optionally, glutamine
can be also withdrawn from the media, in order to assess
ECAR coming from glycolytic pathway and not from glutaminolysis [187] albeit the removal of specific nutrients
can also induce off-target metabolic changes. In addition,
depending on the used cell type, the concentration of each
chemical compound must be carefully determined, in order
to obtain the optimal result for each of these measurements.
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For instance, it has been observed that an overdose of oligomycin can lead to maximal inhibition of OCR, which can
result in a progressive increase of OCR over time [184].
Unfortunately, the molecular mechanism underlying this
observation remains elusive.
12 Endothelial cell precursors
Stem and progenitor cells provide a necessary homeostatic
source of tissue-specific mature cellular elements that
permit appropriate cell functions through replacement of
injured, diseased, and senescent cells in many organ systems
throughout the life span [188]. While the identity, biology,
and molecular regulation of hematopoietic-, intestinal-,
skin-, and skeletal muscle stem and progenitor cells are well
recognized, little is known of the organization of the reparative cells that comprise the vascular system. In 1997, Asahara and colleagues [189] reported on the identification of
circulating progenitor cells for the endothelial lineage. Subsequent studies have clarified that those putative “endothelial progenitor cells” (EPC) did not possess the capacity to
undergo a stable lineage switch to the endothelium, but were
comprised of numerous hematopoietic cells that can serve
paracrine pro-angiogenic functions to promote vascular
repair and replacement but are incapable of integrating as a
bona fide EC in the injured vasculature [190]. In fact, these
pro-angiogenic cells can upregulate “endothelial cell” markers and thus give the impression they were becoming EC at
sites of injury, but failed to persist as functional vasculature
long term. A study by the Mayr lab later showed that these
are in fact mononuclear cells and that the “endothelial” surface markers were acquired by a process of membrane fusion
with platelet microparticles [191]. Only endothelial colonyforming cells (ECFC), also called late outgrowth or blood
outgrowth EC (BOEC), are direct EC precursors that form
vessels in vivo [190]. This paragraph will briefly discuss
available evidence for and remaining controversies regarding the evaluation of endothelial stem and progenitor cells
in mouse and man.
12.1 Assays to identify endothelial stem
and progenitor cells
A simple working definition of a stem cell is a clonal, selfrenewing cell, which gives rise to differentiated cell types
[188]. A somatic stem cell may be multi-potent (giving
rise to multiple types of differentiated cells) or unipotent
(differentiating into a single lineage of mature cells). A
progenitor cell displays clonal proliferative potential, but
progenitor cells lack self-renewal potential. However, progenitor cells can expand into a single lineage of differentiated cells. Multiple assays have been developed to define
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stem and progenitor cells that reside in different organs and
tissues [188]. Stem cells in any tissue must be interrogated
for clonal self-renewal (most stringently proven by longterm in vivo persistence with retained capacity for contribution to differentiated progeny [fate mapping approach] and/
or use of transplantation assays into primary and secondary
recipient hosts). Progenitor cells are often measured with
colony-forming assays in vitro (to prove a precursor product
relationship). Stem and progenitor cells may also display
distinguishing cell surface markers that can be used to prospectively isolate the cells through the use of monoclonal
antibodies and flow cytometry or magnetic bead separation. Most recently, stem and progenitor cells from some
tissues have become definable by single-cell gene expression
technologies to permit identification by a unique molecular
signature [192, 193]. Presentation of some selected publications (as examples) identifying putative stem and progenitor cells for the vascular system (using many of the above
criteria) follows below.
Progenitor cell assays To identify the progenitor cells
for the endothelial lineage, we need to decide what criteria
should be used to define cells belonging to the endothelial
lineage. Other paragraphs in this paper have made arguments
as to the “best” assays used to define EC characteristics and
their functions based on published evidence. Obviously,
there are numerous phenotypic, morphological, physiological, genomic, proteomic and functional parameters that
constitute unique and characteristic behaviors of EC. However, one cannot identify cells of the endothelial lineage by
expression of a few cell surface markers or of a few RNA
transcripts. Several selected works that use comprehensive
definitions for identifying progenitor cells for the endothelial
lineage follow.
Patel et al. [194] recently proposed several features that
can be used to define resident vascular endothelial progenitor cells for the murine vasculature. Based upon the level of
expression of vascular endothelial cadherin (VE-cadherin/
CD144), platelet EC adhesion molecule (PECAM-1/CD31),
VEGFR-2, and leukocyte common antigen (CD45), ECs present in blood vessels in induced healing wounds or growing
in response to implanted tumors were identified as endovascular progenitor cells (EVP), transit amplifying cells (TA),
and definitive differentiated cells (D). Evidence for these
distinct populations displaying different functional states
was presented using clonogenic in vitro analysis, fate mapping studies in vivo, transplantation analysis, variations in
gene expression, differences in phenotypic analysis (flow
cytometry and immunofluorescence), and the requirement
of certain transcription factors for the differentiation of EVP
to transient amplifying and definitive differentiated states.
Evidence for the identification of human endothelial progenitor cells (circulating or resident) has been published by
several groups. Analysis of the circulating blood of patients
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following a sex-mismatched bone marrow transplant indicated that some of the BOEC were derived from the host, but
the most proliferative cells were donor bone marrow derived
[195]. Umbilical cord blood was reported to be enriched for
clonogenic ECFC that displayed a hierarchy of proliferative
potential and in vivo vessel formation compared to adult
peripheral blood [196]. Furthermore, resident vascular EC
derived from umbilical veins or human abdominal aorta contained clonogenic ECFC with vasculogenic potential. Very
recently, human-induced pluripotent stem cell (hiPSC)derived ECFC have been reported with properties that are
similar to umbilical cord blood ECFC but with some distinct
differences in gene expression [197].
Vascular endothelial stem cells Fang et al. [198]
reported that stem cells for the endothelial lineage
can be isolated from lung blood vasculature. Vascular
endothelial stem cells (VESC) expressing the phenotype
CD31+CD105+Sca-1+CD117+ (devoid of any mature lineage markers for the hematopoietic system) can be isolated
from collagenase-digested lung tissue. The VESC displayed
clonogenic colony-forming activity in vitro and when
implanted in Matrigel subcutaneously in mice, gave rise to
donor-derived blood vessels. Indeed, when 15 VESC (green
fluorescent protein tagged: GFP) were implanted with B16
tumor tissue, GFP-labeled blood vessels were identified in
primary, secondary, tertiary, and quaternary tumor implants.
Even a single VESC displayed the capacity to form donor
blood vessels in vivo in a Matrigel implant. While some
flow cytometric estimations of the frequency of the CD117
expressing VESC in different tissues were reported, no
detailed visualization of the location of the VESC in arteries, veins, and capillaries in different tissues was presented.
In addition, no fate mapping studies to identify the contributions of the putative VESC within various vascular beds in
homeostatic endothelial turnover were reported.
Naito et al. [199] used the Hoechst staining method to
identify resident vascular EC in the side population that were
dormant in steady state, but possessed clonal colony-forming
activity, produced large numbers of mature endothelial progeny, and when transplanted into ischemic lesions, restored
blood flow and reconstituted de novo blood vessels at the
site of injection. Although the surface markers of the SP
cells were similar to primary capillary EC, the gene expression pattern of the SP and main population that fail to retain
the Hoechst dye was significantly different. Several unique
cell surface markers were identified in the SP cells that may
permit prospective isolation of these putative VESC. Use of
Hoechst method fails to permit identification of the putative
VESC in the organ and tissue vasculature during development or after injury and will rely upon further studies using
novel cell surface markers.
More recently, Yu et al. [200] identified protein C receptor-expressing EC as VESC in the mammary fat pad, skin,
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and retina. These VESC exhibited robust clonal expansion
in tissue culture, high vessel reconstituting ability upon multiple rounds of transplantation, and long-term clonal expansion in lineage tracing experiments. Indeed, the VESC were
determined to be bipotent, with contributions not only to the
endothelium but also to pericytes throughout vessels in multiple tissues. The authors suggested that VESC underwent
endothelial to mesenchymal transition to become pericytes
in the examined vascular beds [200].
12.2 Limitations and challenges
Most of the > 9564 publications identified by the term EPC
in Pubmed (endothelial progenitor cell, Pubmed, July 30,
2017) have failed to provide sufficient evidence that the
putative EPC under investigation display clonal proliferative potential and/or the capacity to directly form EC that
undergo vasculogenesis in vivo to form new vessels or
integrate long term into injured vessels in vivo as EC. If
the EPC in question cannot directly give rise to cells of the
endothelial lineage at a clonal level (in vitro or in vivo) or
function as a bona fide EC in vivo, then the term EPC should
not be applied to those cells [190]. The work of Patel et al.
[201] has shown that endothelial progenitors can be identified by applying stringent criteria. However, several issues
remain to be addressed such as the specific sites, localizations and contributions of EVP, transient amplifying and
definitive differentiated cells localization in organs and
tissues at homeostasis (artery, vein, or capillary bed), the
contributions of EVP to transient amplifying and definitive differentiated cells during homeostasis, differences in
the EVP among different organs across the life span of the
mouse, and, last but not least, determination of whether the
EVP represents an endothelial stem cell. Human EPC have
been identified [195–197]; however, no unique identifying
markers have permitted prospective isolation of ECFC from
circulating blood or blood vascular endothelium to permit
identification of the site of origin of ECFC in human subjects and determination of whether these cells display stem
cell activity for the endothelial lineage. Several papers have
published evidence for the presence of resident VESC in
mice; however, the relationship between the unipotent VESC
[198, 199] and the bipotent VESC identified by Yu and colleagues [200] remains unclear. Identification of unique and
distinguishing characteristics of the VESC that discriminate
these stem cells from progenitor and mature endothelial elements awaits further analysis. The difficulty in identifying
proper VESC begs the question what the homeostatic function of these stem cells would be in tissues composed of
differentiated endothelial and other vascular cells that retain
a highly regenerative capacity.
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12.3 Concluding remarks
Altogether these data suggest that progress in identifying
stem and progenitor cells for the vascular endothelium is
possible when using multi-parameter evidence that the putative precursor cells are irreversibly differentiating into cells
of the endothelial lineage. The use of multi-parameter analytics will be required to permit novel studies on the biology
of vascular endothelial stem and progenitor cells and likely
may alter our understanding of vascular development and
the pathophysiology of vascular diseases.
13 Microfluidic assays
The use of microfluidic cell culture systems has changed
the way in which we study and manipulate living cells,
and numerous researchers have leveraged the capabilities
of these systems to advance and refine our understanding
of angiogenesis and microvascular function. Cutting-edge
microfluidic cell culture models for studying angiogenesis
have successfully incorporated principles from quantitative analyses of vascular function, in vitro flow chambers,
microfabrication techniques, and 3D tissue scaffolds. Consequently, microfluidic approaches have enabled unprecedented levels of control of chemical gradients, fluid flow,
matrix composition, and cell–cell interactions, all of which
can be integrated to provide a physiologically relevant context for studying angiogenesis.
13.1 Development and capabilities
Recent advances in microfabrication and biological integration have helped to propel the design and implementation of
microfluidic systems, which comprise an emerging class of
highly modular in vitro culture models of the microcirculation [202, 203]. Microfabricated devices can provide cultured cells with a microenvironment similar to that in vivo,
including the correct ECM (ECM) composition, chemical
species, associations with other cells, and mechanical signals
to mimic tissue- and organ-level function [204]. Interestingly, microfluidics allows generating chemical gradients
and enables the study of EC chemotaxis in 3D geometries.
These systems also contain networks of micron-scale fluidfilled channels that are similar in size and architecture to
microvessels in vivo [205, 206] to faithfully reproduce
certain microvascular phenomena in vitro. Consequently,
the application of microfluidics for studying angiogenesis
has emerged as a major research thrust at the interface of
microsystems engineering and biomedicine. At present, the
majority of microfluidic systems are constructed by soft
lithography or replica molding of an elastomeric and biocompatible polymer (polydimethylsiloxane or PDMS) [207].
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This process is well suited to create structures with defined
shapes on the micrometer scale that can be used to position
cells and tissues, control cell shape and function, and create
highly structured 3D culture microenvironments [208–210].
Researchers have used this capability to assess the influence of the host tissue environment and heterotypic cell–cell
interactions on angiogenesis [211] by co-culturing vascular
EC organized as an intact vessel structure with other cell
types such as cancer cells [212–215], stromal fibroblasts
[216, 217], vascular smooth muscle cells [218], and bone
marrow cells [219]. Applications for these microfluidic coculture studies include fundamental studies of cell population behaviors, high-throughput drug screening, and tissue
engineering [211]. Application of microfluidic systems has
also contributed significantly to our understanding of flowmediated angiogenesis (Fig. 10). For instance, multiple
studies have shown that interstitial flow potently induces
angiogenesis [220–222] and more recently lymphangiogenesis [223, 224]. These studies have also shown that the vessel sprouts triggered by interstitial flow preferentially form
against the direction of flow [220–223]. In other words, vessel sprouts tend to originate from a local pressure minimum
and seek the higher pressure vessel or source [221]. Interestingly, this same sprouting behavior was recently observed
in vivo in avian embryos [225]. In addition to interstitial
flow, intravascular shear stress has been shown to control
sprouting angiogenesis [221, 226] and anastomosis of vessel sprouts [227]. Collectively, these findings suggest that
the local fluid mechanical environment can both determine
whether sprouting occurs and specify where these sprouts
originate from the parent vessel. Moreover, the ability of
microfluidic system to apply specific levels of fluid mechanical stimuli in a highly controlled environment will be invaluable toward advancing the mechanobiology of angiogenesis
[228].
One limitation to PDMS replica molding is that this process produces microchannels and subsequent endotheliallined vessel structures with rectangular cross sections. To
address this limitation, alternative fabrication techniques
have been used to form engineered microvessels with circular cross sections. The most widely used technique to fabricate this type of microchannel is to cast a 3D scaffold housed
within a PDMS chamber around a cylindrical needle or rod
of approx. 100 μm in diameter [229]. Once the scaffold has
polymerized, the needle or rod cast is removed leaving an
open cylindrical microchannel embedded within the 3D
scaffold. Scaffold-embedded circular microchannels have
been used to monitor focal leaks in the endothelial-lined
microchannels, by adopting imaging techniques of fluorescent tracer dyes conjugated to macromolecules (e.g., albumin or high-molecular weight dextran) that are widely used
in intravital microscopy techniques for measuring vascular
permeability in vivo [230, 231]. In addition to quantifying
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vascular barrier function, circular microchannels have been
used to measure vessel sprouting and anastomosis [232].
Since the fabrication of circular microchannels employs
a non-lithographic technique, it loses some of the desirable
attributes achieved with soft lithography such as rapid prototyping and scalability for high throughput [205]. In addition, circular microchannels require casting and removal of
a cylindrical needle or rod inside a hydrogel; thus, they are
constrained to simple linear structures [233]. To address
the aforementioned limitations of the cylindrically casted
microchannels, another circular lumen fabrication technique
was recently developed that uses the principle of viscous finger patterning of pre-polymerized hydrogel specified inside
PDMS microchannels of different geometries, including
branched networks that mimic the topology of bifurcating
microvessels [234]. Another hydrogel-embedded microfluidic model that is of significance uses micropatterned stencils to produce microvascular network inside natural 3D
hydrogels such as collagen and alginate [235, 236]. Finally,
the recent advent of 3D printed microfluidic vessel networks
[237] have enabled the patterning of more complex branching networks that mimic in vivo physiology [238].
13.2 Advantages of microfluidics for angiogenesis
studies
The use of a chip supported by a microfluidic system for EC
culture represents specific advantages. Compared to in vivo
studies, these systems are relatively straightforward to apply
controlled perturbations of extrinsic cues, such as fluid
shear stress and biomolecular gradients. Furthermore, in
contrast to conventional in vitro angiogenesis assays, vessel
Fig. 10 PDMS microfluidic device for analyzing angiogenesis.
Fluid flow can be controlled connecting syringe pumps to the ports,
or by imposing hydrostatic gradients. Flow can be directed through
the endothelial lumens (green), or across the endothelial junctions,
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structures formed in microfluidic systems can be readily perfused to evaluate the effects of fluid flow on vessel maturation. The feature of these systems enables the visualization
and the precise quantification of vessel function (e.g., MVD
and vascular permeability) in response to various extrinsic
cues and allows the construction of distinct tissue compartments (e.g., vascular and perivascular). Moreover, the cells
that comprise each tissue compartment can be independently
genetically modified to reproduce conditional knockout studies in vitro. Interestingly, the material that is typically used
for microfluidic devices (PDMS) is fairly inexpensive and
enables rapid prototyping and the total volume of biological reagents (e.g., cells, ECM, and culture media) that is
required for microfluidic experiments is very low and on
the order of microliters. Finally, these devices can be further
implemented with optical sensors for analytical measurements. Altogether these features enable in vitro screening
platforms for the efficacy of candidate drug compounds in a
physiologically relevant setting [239, 240].
13.3 Limitations and challenges
Microfluidic technologies typically require specialized fabrication techniques and make use of challenging complex 3D
geometries that require multilayer construction. These technologies are beyond the research expertise of most biology
laboratories. Moreover, despite the beneficial properties of
PDMS (e.g., biocompatible, elastomeric, optically transparent, gas and water permeable), there are also several important limitations such as leaking of non-crosslinked oligomers
and non-specific absorption of hydrophobic molecules that
can critically affect cell-signaling dynamics [241, 242].
through the central matrix gel. Sprouting occurs through the apertures
that flank the central 3D matrix and is easily visualized and quantified
(Adapted from [221])
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13.4 Concluding remarks
Existing and emerging technologies for casting, molding,
or 3D printing are changing the way we study cells in vitro.
These methods bridge the gap between tissue culture dishes
and animal models, allowing more informative and representative co-culture systems. Future development may
allow recapitulation of additional aspects of vascular biology in vitro such as intussusceptive splitting or network
remodeling.
14 Flow cytometry and cell sorting assays
Cancer cells modify the microenvironment at the primary
site to favor tumor growth and dissemination [243, 244].
Similarly, tumor cells induce systemic alterations and modify the microenvironment at the (prospective) metastatic site
to create a (pre) metastatic niche favorable for tumor cell
seeding, survival, and growth [245]. The tumor microenvironment (TME) comprises many different cell types, in
particular blood and lymphatic endothelial cells, carcinomaassociated fibroblasts (CAF), myeloid and lymphoid inflammatory and immune cells [243, 244]. Tumor angiogenesis
promotes local tumor growth and invasion by providing
oxygen and nutrients, cytokines and growth factors (angiocrine effects), as well as an escape route toward metastasis [246]. In response to tumor-produced factors, recruited
CD11b+ myeloid cells polarize toward an alternative (M2)
activation state, in contrast to the classical activation state
(M1) [247, 248]. M2-polarized CD11b+ cells promote tumor
growth, invasion, metastasis, and angiogenesis through the
release of growth and motility factors, including VEGFs,
FGFs, tumor necrosis factor (TNF), platelet-derived growth
factor (PDGF), and chemokines [247, 249, 250]. Diverse
tumor-promoting pro-angiogenic CD11b + cell types
have been reported, including VEGFR-1+CD11b+ [251],
Gr1+CD11b+ myeloid-derived suppressor cells (MDSC)
[252], Tie-2-expressing CD11b+ monocytes (TEM) [253],
and cKit+CD11b+ cells [254].
Traditionally, the TME is analyzed by immunohistological staining of tissue sections. While histological techniques have the advantage to render tissue morphology, they
nevertheless bear some relevant limitations: They require
antibodies that work on fixed tissues; they can only detect
one or a few markers simultaneously; stained cells cannot
be recovered for further analysis; they are laborious, timeconsuming, and unpractical for large series of samples; and
they do not allow precise quantification of the number of
cells and signal intensity. To circumvent these limitations,
alternative techniques were adopted [255]. Flow cytometry
(often also referred to as FACS—fluorescence activated cell
sorting) originally developed to analyze blood-borne cells
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[256, 257] is such a technique, now widely used to study the
cellular composition of the TME. In addition, “true” FACS
allows the isolation of cells for further analysis [258].
14.1 Types of assays
Analytical flow cytometry The critical step to analyze the
cellular composition of the tumor microenvironment is to
obtain a suspension of viable, single cells from the removed
tumor tissue (Fig. 11). To avoid collecting cells circulating
in the blood, mice are terminally perfused with physiological
solution prior to tumor take. Also, care should be taken to
remove the surrounding normal tissue from the tumor lesion.
Harvested tumors are dissociated by collagenase and DNase
treatment, and red blood cells are lysed with a lysis buffer.
The cell suspension is passed through a 70-µm mesh filter
to remove tissue debris and cell aggregates that can clog the
FACS, followed by an incubation step with an anti-Fc receptor blocking antibody to avoid non-specific binding of relevant antibodies to immune cells. There are many commercial
kits that perform well tumor dissociation and FcR blocking
(e.g., from Miltenyi Biotec). Finally, cells are incubated with
different antibodies for markers of cells of interest, for example, for vascular endothelial cells (CD45−/CD31+/GP38−),
pericytes (CD45−/CD146+/CD31−/CD34−), Type 2 proangiogenic TAM (CD45+/CD11b+/EGR2 high/CD38 low),
or MDSC (CD11b+, Ly6C+/G+). By using GP38 staining,
it is possible to separate vascular endothelial cells (GP38-)
to lymphatic endothelial cells (GP38 +). A DNA dye can be
added to exclude dead cells. Analysis of blood-circulating
cells only requires a red blood cell lysis prior to incubation with antibodies of interest. Cells can also be stained for
apoptosis [259], DNA content (to determine cell cycle state),
for the expression of intracellular proteins such as kinases
(including their phosphorylation state) [260], cytokines, and
pro-angiogenic factors [261] by adding a permeabilization
step during the staining procedure.
The advantages of FACS-based studies are manifold:
They allow the analysis of multiple markers, routinely 6–10,
and up to 12 or 16, depending on the machine, laser, and
configurations. This permits monitoring multiple cell populations at once with limited numbers of cells (e.g., 105–106).
FACS experiments are easy and rapid to do and can be effectively used as a screening platform to determine the effect
of drugs on endothelial cells or on tumor-associated proangiogenic cells. By using the right strategy and markers,
one can determine the percentage of most cells of the TME.
For example, we used FACS to characterize the effect of
VEGFR/VEGFR-2 inhibition on angiogenesis and immune/
inflammatory cells [262, 263], to identify a novel pro-metastatic/angiogenic CD11b+cKit+ cell [254, 263] and to study
the contribution of CD11b+ cells to angiogenesis and lymphangiogenesis [247, 264, 265] in mice and human. Many
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factors regulating angiogenesis, including soluble proteins
and microvesicles, are released from the primary tumor site
and can enter the systemic circulation. FACS can be used to
quantify multiple circulating cytokines at once using bead
arrays (e.g., BD™ Cytometric Bead Array) [266]. The assay
takes advantage of the broad dynamic range of fluorescence
detection offered by flow cytometry and the simplicity of
antibody-coated beads resulting in the efficient detection of
multiple cytokines at once. This assay allows analysis of
smaller sample volumes and is faster compared to traditional
ELISA. Cancer-released microvesicles, encompassing exosome, apoptotic bodies, and microparticles, are small cell
membrane-derived vesicles expressing cellular proteins at
their surface and containing DNA, RNA, miRNA and proteins, and small molecules of the cell of origin. They have
gained considerable attention recently as they allow to study
tumor features through a so-called liquid biopsy approach
[267, 268]. FACS can be used to study microvesicles to
determine their level and the surface composition [269].
Fluorescent and Magnetic Activated Cell sorting (FACS
and MACS) Virtually, all stainable cells can also be isolated
by FACS, including endothelial cells, immune/inflammatory cells, CAF, and tumor cells. Isolated cells can be used
for functional experiments or lysed immediately to measure
mRNA/DNA content by RNA/DNA sequencing or PCR or
detect proteins by proteomics or antibody arrays, during
natural tumor progression or in response to treatments, for
example to identify pathways and their alteration by drugs
[262, 264, 270]. A critical parameter for successful cell isolation is the relative density of the to-be-sorted cells in the
starting population: The lower the frequency (e.g., below
1%) the longer the sorting time (which may affect viability) and the lower the purity (which can pose problems for
genetic, genomic, or proteomics studies). Microvesicles can
also be isolated by FACS [269]. An alternative to cell isolation by FACS is magnetic cell sorting (MACS). Cells are
stained with antibodies coupled to nanosized superparamagnetic particles and isolated positively or negatively with the
aid of a column and a magnet [271]. This technique allows
gentle separation of a larger number of cells. While negative
selection can be performed with multiple antibodies (i.e., to
eliminate many different cells), positive selection can only
be done with one marker at a time. MACS is often used as
a first enrichment step for rare cells of interest, followed by
multi-parametric FACS sorting.
14.2 Limitations and challenges
Despite the power of FACS to analyze multiple cellular
or molecular parameters in a single experiment, there are
important limitations. A main one is the absence of morphological information and position of cells. For example,
the pericyte coverage of blood vessels and the organization
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of the vessels (size and diameter) as well as the relative
positioning of immune/inflammatory cells to vessels or
tumor cells cannot be obtained by FACS analysis. Having
this information is important as, for example, some treatments do not decrease the number of vessels but alter their
structure (“vascular normalization”) or modify the localization of immune/inflammatory cells but not their number.
Thus, FACS studies should be supported by immunohistological analyses. A second limitation is that the technique
is semiquantitative. Absolute cell numbers can be indirectly
calculated though, by relating the relative frequency of the
cell of interest obtained by flow cytometry analysis, to one
main population in the test sample (e.g., CD45 + cells),
whose cell number was previously counted manually. Cell
sorting has limitation for viability and contamination. For
example, endothelial cells obtained by FACS are difficult
to culture and use for ex vivo/in vitro (functional) experiments. As stated above, sorting of rare cells (< 1%) can be
hampered by contaminating cells (in some cases, up to 50%).
A pre-enrichment by MACS is highly recommended in such
situations. As these technologies are rapidly evolving, new
opportunities are emerging allowing circumventing some
of these limitations or opening new opportunities. These
include laser scanning cytometry for the multi-parametric
study of tumor microenvironment in situ [272] and assessment of cellular/subcellular elements in individual cells
[258], the use of label-free spectral measurements to analyze
native individual cells [273], or the detection and imaging
of circulating cells directly in blood or lymph [274], to cite
only a few.
14.3 Concluding remarks
Flow cytometry and FACS-based experiments and analyses offer important (semi)quantitative, high content, and
accurate information about tumor angiogenesis and associated cells of the TME and in the circulation. It is a remarkable powerful and robust technique, which can be used as a
screening platform to test the anti-angiogenic capability of
molecules and drugs for anticancer treatment or to in-depth
analysis of the composition of the TME during tumor progression or in response to therapeutic interventions. FACS
analysis should always be complemented with immunohistological studies to characterize the morphology of the vessels
and positioning of cells of interest, particularly when investigating anti-angiogenic effects of new molecules and drugs.
Angiogenesis (2018) 21:425–532
461
Fig. 11 Representative flowchart of flow cytometry and cell
sorting experiments. Overview
of samples collection (tumors,
organs bearing metastasis,
and blood) from mice and the
procedure to obtain a singlecell suspension after tissue
dissociation, erythrocytes lysis,
and cell labeling with fluorescent or paramagnetic-coupled
antibodies. Analysis of bloodcirculating cells only requires
a red blood cell lysis prior to
incubation with antibodies of
interest. Intracellular antigens
can be detected by adding a cell
permeabilization step. Fluorescent labeled cells are analyzed
by flow cytometry and sorted
by FACS while cells labeled
with paramagnetic-coupled
antibodies can be sorted by
MACS. MACS can be used to
pre-enrich cells for subsequent
FACS sorting. Cell populations
of interest are identified in by
dot plots analysis of collected
data
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15 Loss‑of‑function approaches
in the developing zebrafish
In recent years, the zebrafish (Danio rerio) has become a
highly utilized vertebrate model system for studying vascular development and angiogenesis in vivo. Vascular development in zebrafish is remarkably similar to that in mice and
humans in terms of both anatomy and function. Zebrafish
are genetically accessible with externally fertilized, optically clear embryos and larvae that permit high-resolution
imaging of developing tissues. Fish also have large clutch
sizes for experimentation, and their embryos can survive
and continue to develop for several days without a functional
circulation via passive diffusion of oxygen. Although the
beneficial experimental attributes of zebrafish have made
this an extremely appealing and approachable model system,
like any model organism, there are limitations to the studies that can be performed and their interpretation. Here, we
focus on loss-of-function approaches used to study vascular
development in the zebrafish and address some of the pitfalls in interpreting data generated using currently available
approaches.
15.1 Morpholinos
Prior to the advent of genome-editing technologies in the
zebrafish, other than mutations fortuitously isolated in
forward-genetic screens, morpholinos (MOs) were the
only available reasonably well-validated loss-of-function
method. MOs are specially modified stable antisense oligomers designed to block either gene translation or gene
splicing when injected into embryos [275]. In the absence of
reverse genetic tools, MO-based experiments rapidly gained
popularity in the fish community. Although the advent of
this technology was a huge step forward for the zebrafish
community, over time many concerns have been raised about
the specificity of phenotypes generated from MO injections,
particularly the potential for off-target and/or pleiotropic
effects from MO injection [275–278]. Indeed, while it is
possible to show that a particular MO has the desired inhibitory effect on the target gene, it is impossible to exclude
that there are (latent) unspecific effects [279]. These concerns have been greatly heightened by mounting evidence
that MOs and genetic mutants for the same genes frequently
do not yield comparable phenotypes [276]. In order to promote greater validity of experimental studies using MOs,
attempts have been made to develop standards for their use.
One recently promulgated set of “community guidelines”
suggests the use of multiple MO targets, RNA/DNA rescue
experiments, dose–response curves for titration of the MO,
and, importantly, validation with a genetic mutant where
possible [275]. It is generally accepted that proper MOs
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use requires rigorous confirmation of their specific blocking ability, either by assessing loss of protein production
for translation-blocking MOs (if specific antibodies are
available) or by assessing the generation of alternate splice
variants by RT-PCR when using splice blocking MOs. MObased experiments can also be “rescued” by co-administration of mRNA or transgenes expressing a wild-type copy of
the targeted gene. However, the best and generally accepted
validation for any MO phenotype is confirmation of the same
phenotype in a zebrafish genetic mutant. As discussed further below, many mutants are now available from forward
genetic ENU (N-ethyl-N-nitrosourea) mutagenesis screens,
“TILLING” approaches [280], or genome-editing methods.
CRISPR technologies have made it easy and straightforward
for any laboratory to carry out reverse genetic mutation of
virtually any gene of interest [278, 281–301]. In short, when
possible, all MO experiments should be confirmed and validated with a genetic mutant before they are used extensively
for experimental studies.
Common off-target effects noted in MOs-injected animals
include p53-mediated cell death, defective circulation and
“ballooning” (edema), and developmental delay/stunting
of embryos and larvae. Altered vascular development and
patterning should be interpreted with extreme caution in
the presence of any of these phenotypes, as vessel growth
and patterning in developing embryos and larvae are highly
dependent on the integrity of adjacent tissues and of the
animal as a whole (see below for further detailed discussion). The recently published community guidelines noted
above include recommendations on how to interpret MO
phenotypes, how to validate these phenotypes, and current
expectations in the field for MO usage [275].
Although MOs approaches, and especially their use without proper validation, have justifiably come under a great
deal of scrutiny (if not outright skepticism) in recent years
due to well-documented frequent off-target effects and lack
of correlation with mutant phenotypes, it is worth noting
a few positive/beneficial features of MOs that make their
continued use (with caution) worthwhile. First, translationblocking MOs can block both the zygotic and maternally
supplied activity of target genes, whereas maternal-zygotic
mutants can be difficult or impossible to obtain. Second,
MOs can be used immediately with any zebrafish line or
strain unlike mutants, which need to be crossed into the
appropriate genetic backgrounds (when using combinations
of mutants and transgenes, this can take many generations).
Third, multiple MOs can be injected simultaneously to target
two, three, or more members of a gene family with potentially overlapping functions. Fourth, recent work has shown
that upregulation of related compensating gene family members can sometimes occur in genetic mutants (by mechanisms that are not yet clear), while this does not appear to
take place in MOs-injected animals [277], arguably making
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MOs a better representation of targeted loss of gene function
in these cases.
15.2 Genetic mutants
The ability to carry out large-scale forward-genetic phenotype-based ENU mutagenesis screens for developmental mutants has been one of the major strengths of the
zebrafish. Large numbers of mutants have been isolated
affecting almost every conceivable developmental process, and these mutants have led to innumerable important
new discoveries [302–309]. Forward genetics has been an
extremely powerful approach for identifying genes necessary for developmental processes, but until recently reverse
genetic approaches for targeting specific genes of interest
were not available in the zebrafish. “TILLING” approaches
employing high-throughput next-generation sequencing of
libraries of mutagenized fish have been used to screen for
ENU-induced mutations in specific genes [310] but these
approaches are relatively laborious and expensive, and some
genes are not as easily mutagenized. More recently, genomeediting technologies and particularly CRISPR/Cas9-based
methods have made reverse genetic targeting of virtually
any gene easy and cost-effective for most laboratories
[281–283]. A number of resources now make identifying
“ideal” CRISPR cut sites, primer selection, and mutation
screening relatively simple [281, 311]. Generating specific
alleles by homologous integration is still relatively challenging in the zebrafish, but the technology and resources to generate “knock-in” mutants using are also rapidly improving
and are likely to soon become accessible to most zebrafish
laboratories [283].
Although CRISPR mutants offer a simple and effective
approach for loss-of-function analysis free of many of the
caveats regarding off-target effects associated with MOs use,
the resulting phenotypes must still be interpreted rigorously.
Vascular phenotypes observed in animals that display significant cell death, defective circulation, edema, developmental delay/stunting of embryos and larvae—while they result
from defects in the targeted gene—may represent secondary, indirect consequences of the genetic mutation and may
not reflect a direct requirement for the targeted gene during
angiogenesis. It is also worth noting that transient CRISPR
approaches involving, for example, injection of guide RNAs
into animals transgenically expressing Cas9 in specific tissues (“CRISPRi”) are subject to the same general concerns
regarding off-target effects of MOs, although the spectrum
of off-target effects observed with the two approaches will
likely be distinct. Phenotypes observed in either MO- or
CRISPR-injected animals must be interpreted with caution
and ideally should be verified using a stably transmitted germline genetic mutation.
463
The ease and efficiency with which CRISPR mutants can
be generated also makes it possible to simultaneously induce
and screen for mutations in multiple genes at the same time.
This is advantageous for studying the role of gene families in the zebrafish. Gene paralogues are more common in
zebrafish than in mammals due to a genome duplication that
occurred many millions of years ago during the evolution
of teleost fish, and genetic compensation by alternate gene
family members can lead to obscured phenotypes [277]. It
has become more common to see reports in which double
or even triple mutants have been generated in closely related
gene family members [312]. As noted above, compensatory
upregulation of related family members has been shown to
occur in at least some genetic mutants [277], and phenotypes comparable to those noted in MO-injected animals
have been unmasked by generating double or triple mutants.
15.3 Assessing vascular phenotypes
in loss‑of‑function models
Whether MOs or mutants are used, assessment of vascular
phenotypes involves a number of specialized considerations,
since proper vessel formation can be altered or disrupted by
developmental delay or by changes in circulatory flow and
local or generalized defects in non-vascular tissues. Below,
we discuss some general considerations in assessing vascular
phenotypes and point out some potential pitfalls in utilizing
and interpreting zebrafish vascular data.
15.4 Where is the gene expressed?
One of the first considerations in assessing the vascular function of a particular gene, whether using MOs or mutants,
is where it is experessed. This is generally assessed using
whole-mount in situ hybridization (WISH) of zebrafish
embryos and early larvae [313], although in older animals in situ hybridization of tissue sections can be used.
If a gene shows an exclusive vascular expression pattern
(Fig. 12a, b), the gene may have a vascular-specific function,
and vascular phenotypes observed likely reflect a vascularautonomous role for the gene. On the other hand, if the
gene shows an exclusively non-vascular expression pattern
(Fig. 12c, d), it is not reasonable to assume that the gene
has a vascular-autonomous function, and any vascular phenotypes that are observed are likely indirect. In many cases,
expression is observed in both vessels and in non-vascular
tissues (Fig. 12e), and some commonsense judgment must
be applied in assessing whether observed loss-of-function
phenotypes make sense in terms of the expression pattern of
the gene. If significant non-vascular expression is observed,
additional experimental approaches will be needed to assess
the cell autonomy of gene function, including transplantation
13
464
experiments or tissue-specific transgenic expression to “rescue” the phenotype (Fig. 12f–h).
15.5 Assessing general morphology
and development in morphants or mutants
Apparent vascular-specific phenotypes often occur as secondary, non-specific consequences of general developmental
delay and other localized or general changes in non-vascular tissues or organs. Staging tables are available for the
zebrafish [294], and it is important to determine whether
MO injections or genetic mutants result in either overall
delays to development or gross embryonic/larval patterning,
or localized defects in the development and morphology of
specific tissues or organs (Fig. 13a, b). For morphants, this
should be done by comparing animals injected with specific
MOs to siblings injected with a control MOs and uninjected
siblings. For mutants, they should be compared to phenotypically wild-type siblings from the same clutch of eggs. In
Fig. 12 Assessing EC autonomous gene function in zebrafish. a–e
Whole mount in situ hybridization of 24–48 hpf zebrafish embryos.
a–b ve-cadherin labeling of the vasculature in the trunk (a) and
head (b) of a zebrafish embryo showing vascular-specific labeling. c–d tagln/sm22 (c) and vegfa [308] (d) showing labeling
of non-vascular (somitic) tissues in the trunk. e cds2 [309] labeling of both vascular (arrows) and non-vascular tissues. f Confocal micrograph of a growing trunk intersegmental vessel in a 32 hpf
Tg(kdrl:mRFP-F)y286 embryo (red vessels), mosaically expressing
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either case, if the overall development of the animal is significantly delayed or readily apparent, gross morphological
changes are noted compared to the control sibling animals
and vascular phenotypes noted in these animals should be
interpreted with caution. For example, reduced head size
and/or extensive cell death in the central nervous system are
common effects noted in both mutant and morphant models,
and this secondarily results in vessel defects in the heads of
affected animals (Fig. 13g, h). The timing and extent of trunk
intersegmental vessel growth is frequently used as a convenient and quantitatively robust assay for assessing angiogenesis defects in zebrafish embryos, but delayed development or
abnormal formation of the adjacent somites results in defects
in vessel sprouting and growth that, again, are not directly
linked to vascular-specific functions (Fig. 13c–f, i–j). Similarly, the formation of the thoracic duct as a readout for lymphatic development is sensitive to developmental delay and
general effects on morphology; hence, the absence of the
thoracic duct in itself is insufficient to demonstrate a defect
Tol2(fli1a:H2B-TagBFP-p2A-egfp-F) transgene (blue EC nuclei,
green EC cytoplasm), showing blue, green, and red fluorescent channels and all three merged. g–h Higher-magnification images of GFP
fluorescence (g) and merged GFP/BFP/RFP fluorescence (h) in a 48
hpf Tg(kdrl:mRFP-F)y286 transgenic animals mosaically expressing
a Tol2(fli1a:H2B-TagBFP-p2A-egfp-F) transgene in a single EC in
a trunk intersegmental vessel. Scale bars = 20 µm (f), 10 µm (g–h).
Images in panels f–h are from Ref. [649]
Angiogenesis (2018) 21:425–532
in lymphangiogenesis. Staging of embryos and excluding
developmental delay are critical, as is monitoring the distance between dorsal aorta and PCV in phenotypic embryos.
Importantly, lack of a lymphatic system as found in ccbe1 or
vegfc mutant embryos [314–316] only leads to formation of
edema after 120 dpf—any edema formation prior to 120dpf
in experimental embryos is therefore very unlikely to result
from a lymphatic defect.
15.6 Assessing circulatory flow and cardiac function
Even in animals with apparently normal overall development, defective blood flow (due, for example, to heartspecific defects) can affect the timing and extent of vessel
growth. Assessing blood flow and cardiac function in the
zebrafish is easily accomplished by direct imaging using a
dissecting light microscope. For higher resolution, tracers
injected into the blood stream such as quantum dots, fluorescent microspheres, lectins, and varying molecular weight
fluorescent dextrans can be used to assess blood flow rates,
blood flow directionality, vascular leak, vessel drainage
sites, and solute uptake [317–320]. Although zebrafish can
survive a number of days in the absence of blood flow, animals lacking circulation do eventually become sick, edemic,
stunted in growth, and die. If a mutant or MO experiment
generates zebrafish with no blood flow or decreased cardiac
output, assessment of vascular phenotypes should take place
as early as possible during development, during a time frame
in which the animal is as healthy as possible. Although
embryos and early larvae can develop reasonably normal
for one or 2 days in the absence of circulation (Fig. 13k, l),
overall vascular development becomes increasingly affected
as time goes by (Fig. 13k, p). As a general rule, phenotypes in animals deficient in flow should be assessed prior
to 6 days post-fertilization (dpf), and ideally before 3 dpf,
particularly when flow is entirely absent. As noted for animals with general developmental delay or morphological
defects, caution should also be taken in interpreting vascular
phenotypes observed in animals that lack circulatory flow.
Quantitative assessment of blood flow and hemodynamics can be accomplished using Particle Velocimetry (PV),
in which tracer particles injected into the blood stream are
tracked through different vessels [321] and flow speed is
determined by calculating the displacement of the particles
over time [322]. Due to depth and/or the decreased transparency at later stages, PV has mainly been used in embryos or
early larvae, although recent advances in confocal microscopy, ultrasound, and tomography tools [323] and methods
have facilitated deeper imaging in more opaque tissues.
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15.7 Assessing vascular patterning
Visualization and assessment of vascular patterning in
the zebrafish is easily accomplished using widely available transgenic fluorescent reporter lines labeling vessels.
Numerous transgenic zebrafish lines are available with
cytoplasmic, nuclear, or membrane-localized fluorescent
proteins expressed specifically in cardiovascular cells and
tissues [324–326]. These include lines marking the heart
endocardium and myocardium, blood and lymphatic vessel
endothelium, hematopoietic derived cells, and perivascular cells like pericytes and fluorescent granular perithelial
cells. Long-term time-lapse imaging methods have also been
developed that permit continuous, real-time imaging of heart
and vascular development in these transgenic lines [327].
Together, the availability of fluorescent transgenic reporter
lines and methods for high-resolution, dynamic imaging of
these lines has revolutionized the study of cardiovascular
development.
Alternatively, or in addition, intravascular injection of
fluorescent tracers such as quantum dots, fluorescent microspheres, or fluorescent dextrans, or non-fluorescent tracers
such as India ink, Berlin blue, or Evans blue dyes can be
used to visualize zebrafish blood vessels and their patterning [317, 319, 327–332]. Although the wide availability of
vascular-specific transgenic lines has reduced the need for
these traditional methods for examining the patterning of
developing vessels, microangiography is still the method of
choice for assessing vascular integrity [333], vessel lumenization, and flow (as noted above) in blood vessels, as well as
permeability, drainage, and solute/fluid uptake in lymphatic
vessels. The microangiographic techniques are fast, robust,
and cost-effective methods for visualizing the vasculature
and can be used to visualize vessels in non-transgenic animals or at more mature developmental stages.
The patterning of vessels in genetically or experimentally
manipulated animals can be examined for alterations including reduced/absent vessel growth, excessive vessel growth
or branching, altered patterning of vessels, changes in flow
patterns/vascular connections, etc. Changes can occur either
broadly throughout the animal or may be localized to particular vascular beds. Up to approximately 7 dpf, the pattern
of blood vessels observed should be compared to the published staged atlas of vascular anatomy [328]. Although the
positioning of major vessels (e.g., dorsal aorta, cardinal vein,
and intersegmental vessels in the trunk, or basilar artery in
the head) is relatively invariant in normal animals at given
stages, the precise location and paths taken by many smaller
and/or later forming vessels (e.g., the central arteries of the
hindbrain) are less stereotypic, and minor changes in the
patterning of these vessels may not be significant. As noted
above, it is important to assess vessel defects in genetically
or experimentally manipulated animals in the context of
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Angiogenesis (2018) 21:425–532
Fig. 13 Assessing embryonic morphology and effects on vascular patterning. a–f Transmitted light (a, b), epifluorescence (c, d),
and confocal (e, f) images of three dpf developmentally normal (a,
c, e) or developmentally abnormal (b, d, f) embryos from the same
Tg(fli1:egfp)y1 transgenic “wild-type” zebrafish population. The normal animal has a normal morphology (a) and normal vessel patterning (c, e), while the developmentally abnormal animal has a stunted,
somewhat malformed trunk (B) and also displays trunk intersegmental vessel patterning defects (d, f). g, h Confocal images of the cranial vasculature in 48 hpf wild-type sibling (g) and y284 mutant (h)
Tg(kdrl:mRFP-F)y286 animals. The y284 mutants have small heads
heads and eyes, accompanied by reduced and abnormal formation
of cranial vessels and aortic arches (brackets). The vascular defects
should be interpreted with caution, as they can be primary or solely
a consequence of the smaller head (or possibly both). Transplantation and/or mosaic transgenic expression can be used to assess the
vascular cell autonomy of phenotypes. i, j Confocal (top) and corresponding transmitted light (bottom) images of 48 hpf Tg(fli1:egfp)y1
wild-type sibling (i) and fused somites (fssy66) mutant (j) animals.
Improper somite formation in fssy66 mutants (lack of chevron shaped
somite segments, seen in the bright-field images) indirectly results
in altered intersegmental vessel patterning (j, top). k–p Confocal
images of the hindbrain vasculature (k, l) or trunk intersegmental vessels (m–p) in Tg(fli1:egfp)y1 control (k) or cardiac troponin T type 2a
(tnnt2a) (l–p) deficient animals at 1.5 (m), 2 (k, l, n), 2.5 (o), or 3.5
(p) days post-fertilization (dpf). Control animals have normal blood
flow, but tnnt2a-deficient animals have no heart beat and lack all
blood flow. Although formation of the vasculature is largely normal
for several days in the absence of blood flow (l, n, o), abnormalities in
vessel growth and patterning begin to appear at later stages, such as
enlargement of the dorsal intersegmental vessels at 3.5 dpf (p). This
illustrates the need to analyze vascular phenotypes as early in development as possible in zebrafish with absent or defective blood flow.
All images are lateral views, rostral to the left, except panels k and l,
which show dorsal views, rostral to the left. Images in panels i and j
are from Ref. [306], images in panels k and l are from Ref. [650], and
images in panels m–p are from Ref. [651]. Scale bars = 50 μm
the animal as a whole, including whether any developmental delay or gross morphological defects are present. The
timing and extent of intersegmental vessel sprouting is frequently used to assess angiogenic phenotypes in developing
zebrafish, but their patterning can be strongly affected by
developmental delay, non-vascular morphological defects,
and reduced or absent circulatory flow (Fig. 13).
15.8 Concluding remarks
13
A variety of approaches for reverse genetic targeting of specific genes for loss-of-function analysis are now available
in the zebrafish, most notably MOs and CRISPR-generated
mutants. Although MOs have been somewhat discredited in
recent years as a “first line” tool for assessment of loss-offunction phenotypes, they remain valuable when rigorously
Angiogenesis (2018) 21:425–532
validated by a corresponding mutant with the same phenotype. Furthermore, in some cases, mutants are susceptible
to genetic compensation by upregulation of related genes,
while morphants are not. Detailed “community guidelines”
for the use of MOs were recently published and should be
referred to [275].
Any researcher generating loss-of-function vascular phenotypes in the zebrafish (or in any other model organism,
for that matter) needs to critically assess their data in light
of some key questions. Where is the gene expressed? Do
mutant or morphant phenotypes make sense when compared
to the expression pattern of the gene? The more “vascular-specific” expression is, the more likely the associated
vascular phenotypes are specific. Conversely, if a gene is
ubiquitously expressed throughout the animal, the burden
of proof is on the researcher to determine the vascular cellautonomous function of the gene, using transplantation,
tissue-specific transgene “rescue” experiments, or other
methods. Do mutants or morphants show significant developmental delay or gross morphological phenotypes, and do
the vascular phenotypes most likely reflect developmentally
“younger” animals or problems in non-vascular tissues? Do
the mutants or morphants have absent or strongly reduced
circulatory flow? Are vessel phenotypes being assessed in
these animals at a time point prior to lack of flow causing significant general defects in development? Taking all
of these criteria into consideration when assessing vascular phenotypes resulting from gene knockdown or genetic
mutants is important to ensure that reliable, valid conclusions are made when studying regulators of angiogenic
development utilizing the zebrafish.
16 Chorioallantoic membrane assays
Although the chicken chorioallantoic membrane (CAM)
assay (Fig. 14a) appears to have been first reported by Rous
and Murphy in 1911 [334], more than a century ago, to
study xenoplastic growth of mammalian tumors, the chick
embryo itself has been a target of scientific study beginning
with Aristotle. Due to the fact that fertilized chicken eggs
are readily available and the embryos are easily visualized,
chicken embryos became a favorite target for experimental
biologists and, not surprisingly, the CAM assay became one
of the classic procedures for students, being included in one
form or another in virtually every college laboratory manual.
Due to these advantageous properties, the CAM has been
widely used not only for direct vascular biology studies, but
also in bioengineering, cosmetics testing, transplant biology,
drug and vaccines development, vaso-occlusive therapies,
or cancer research [335–343]. There are certainly several
hundreds of variations in the experimental protocol, all of
them involving placing a test material on the CAM or an
467
intraveneous injection. The variables, however, often largely
unbeknownst to the user, lead to highly significant differences in results and a striking diversity in observations and
conclusions. Several aspects of the diversity are described
below.
16.1 In ovo CAM assays
The “standard” (i.e., “classic”) assay consists of exposing
the CAM in the incubated egg by making a window through
which the CAM can be accessed and placing a test graft,
implant, or substance on, in, or through it. After resealing
the window, periodical examination of the test area can be
performed, ultimately measuring the result, frequently by
harvesting the area for chemical or histological analysis.
Incubation temperature. The standard Hamburger and
Hamilton stages of chick embryo development for stages
14–35 were based on incubation at 39.4 °C using White
Leghorn chickens [344]. The other stages were based on
incubation temperature of 37.5 °C. Stage 33 (7 days) is when
many investigators place a graft or sample on the CAM. This
may sound trivial, but at stage 33 the CAM is slowing its
growth. If the CAM is accessed on day 7 of an embryo incubated at 37 °C (common in most laboratories), CAM cells
are still rapidly proliferating [56]. Moreover, depending on
the source and strain of eggs used, there may be a 12–18 h
variation in developmental age.
Making a window. The original method used a handheld
saw blade, which worked well but frequently led to debris
falling on the developing CAM. Such debris can provide
artifacts difficult to distinguish from results due to planned
experimental protocols. Ed Zwilling (in 1952) had a Dremel
drill sanding disk in his garage which worked well and is
now used nearly universally, but even so, debris, often too
fine to see, can materially influence the results [56].
Placement of test material/grafts. The CAM itself is, as
indicated by its name, multilayered. Placing a filter or graft
on the membrane is quite different from inserting it part way
(preferred by many) or penetrating the CAM completely so
that it projects through the entire membrane. Results may
be completely different depending on the mode of insertion
as well as on the physical forces used [345]. A second variable is the choice of location on the membrane. The CAM
at stage 30–35 is highly vascularized (hence, a good choice
for transplants), but there are vessels of all sizes. Choosing
a site near a large vessel sets up conditions quite different
from choosing a less vascular site. Sprouting will depend
on placement as well rate of dispersion of test substances
and the elution from disks and filter membranes. The reader
is referred to several excellent comprehensive reviews that
describe a wide range of modifications and adaptations of
the basic CAM assay [335, 346–351].
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Angiogenesis (2018) 21:425–532
Fig. 14 Chorioallantoic membrane of the chicken embryo (CAM). a
Bright-field image of the CAM vasculature presenting the functional
vascular network from capillaries to big vessels imaged at embryo
development day 10 (E10). Bar corresponds to 500 μm; b Vascular
network of the CAM presented on the fluorescent angiography with
FITC-dextran and an intravenous contrast agent. Bar corresponds to
500 μm; c Scanning electron microscopic picture showing the chorionic epithelium (CH) and two large vessels (VE) in the intermediate mesenchyme of a CAM at day 5 of incubation (reproduced from
[652]); d A 12-day CAM incubated on day 8 for 4 days with bioptic
specimen of ACN/neuroblastoma cell line tumor xenograft, showing
numerous blood vessels around the graft (reproduced from [653]).
Bar corresponds to 500 μm; e Immuno-histology of a glioma (U87
tumor) implanted on the CAM with tumor cells in red (vimentin
staining and vessels in green(SNA isolectin staining), magnification
x4; with permission from [362], copyright (2005) National Academy of Sciences, USA; (f) a pancreatic adenocarcinoma (BxPC3)
nodule (blue, Hoechst) inside of the CAM surrounded by blood vessels (green, SNA isolectin staining). bar corresponds to 100 μm;
with permission from [363], copyright, Elsevier (License number
4307171269037); g MCF7-derived tumor implanted on CAM induces
angiogenic response. Tip cell (red arrow) and incompletely attached
pericytes (yellow arrow)
Day 7 of the CAM incubation represents the proper time
to place grafts on the CAM surface for investigation of angiogenic responses. There are several reasons why this time
for grafting is preferential: (1) Areas between big vessels are
less vascularized and the assessment of angiogenic events is
more objective, (2) the immune system is not yet developed,
and (3) interaction between immature developing mesoderm
of the CAM and a graft gives a proper microenvironment
for the survival, development, and vessels acquisition by
angiogenic grafts.
of a curved rather than flat-bottomed dish, and modifications
have ranged from specially designed watch glass-type vessels to plastic wrap loosely fastened over emptied food cans.
Major advantages over in ovo methods are better visibility,
potential to use transmitted light, ready application of several test sites, time-lapse monitoring, and multiple manipulations. Disadvantages include significant mortality, greater
risk of infection, the need for more rigorous control of the
incubation environment, and materially greater expense.
16.2 Ex ovo CAM assays
16.3 Angiogenesis platform using a cubic artificial
eggshell with patterned blood vessels
Once it was demonstrated that one can carry out CAM
assays in shell-less egg cultures [352, 353], this modification became a staple variant of the assay. Again, there have
been numerous improvements and variants, but all involve
transferring the egg content at an early stage (day 2–3) into
a container, allowing the CAM to develop and then carry
out procedures as in the standard in ovo protocol. The major
modification from the original petri dish method is the use
One of the newest models using the chick embryo chorioallantoic membrane as a tool for blood vessels assessment and
dynamics was recently described by Huang et al. [354]. The
authors provided a methodology to direct blood vessel formation on the surface of a three-dimensional egg yolk using
a cubic artificial eggshell with six functionalized membranes. By using this method, blood vessel assessment of
the CAM is improved by controlling their growth directions
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in special microfluidic chambers with a width of 70, 250,
and 500 μm. Since the blood vessels with blood flow in
microchannels are still alive, several culture chambers for
implanted cells/tissues can be fabricated in the patterned
surface during one experiment. Because the cubic eggshell
with patterned surfaces is transparent, reagent injection and
screening is possible in each of the chambers separately,
and the development and spreading of blood vessels into
the cultured tissues in the chamber can be observed using a
microscope [354].
16.4 Assessing vascular patterning
Visualization and assessment of vascular patterning in the
CAM model is one of the most challenging aspects of the
assay. Quantification of angiogenesis by appearance (number
of vessels and their hierarchy, degree of branching, etc.),
whether by simple inspection or quantitative image analysis, is highly variable. To obtain statistically significant data
requires multiple samples. Monitoring during the course of
the experiment is often foiled by the spreading and hence
moving of the test site from the observation window. Most
importantly, many of the visible measurements cannot distinguish well between neovascularization and vasodilation or
between inhibition and vasoconstriction. Several semiquantitative or quantitative imaging [355, 356] or immunohistochemistry-based [357] methods for vascular and capillary
pattering assessment have been developed. The descriptors
include, for example, branching points/area, vessel or capillary density, or mean mesh size. Apart from intravenous
injection of fluorescent dextrans (Fig. 14b) [358, 359] or
non-fluorescent tracers such as ink [360, 361], MRI and PET
imaging, scanning electron microscopy (Fig. 14c), or optical Doppler tomography are now available to facilitate the
visualization of the CAM vasculature (see a comprehensive
review [335]). For some of these techniques, however, the
resolution might be an important limitation and should be
considered prior to imaging technique selection. Another
point to take into account in imaging technique selection
is the vascular development stage, vascular permeability,
and lymphatic uptake. Recently, imaging was facilitated by
the replacement of the original eggshell by an artificial one
[354].
16.5 The experimental chicken embryo tumor
model
Tumor cells can easily be grafted on the chicken CAM and
develop vascularized tumor (Fig. 14d). The way tumor
implantation is done is dependent on the tumor type that
is studied, see Table 2. In the case of glioblastomas, when
implanted onto the CAM [362] (Fig. 14e), a plastic ring
is placed on the surface of the chorioallantoic membrane
469
at embryo development day 10 (E10). Gentle lacerations
of the CAM surface are carried out with scalpel blades in
order to facilitate and promote tumor engraftment. Implantation of four million tumor cells inside of the plastic ring
in serum-free culture medium is usually used with daily
monitoring over seven to 8 days. At E17 or E18, remove the
plastic ring by cutting out the CAM area and proceed for
analysis (OCT inclusion, snap-frozen, etc.). Tumors such as
the highly angiogenic glioblastomas (U87 etc.) are growing
very well with an angiogenic response that starts at day 3–4
post-implantation.
In case of highly metastatic tumors such as pancreatic
ductal adenocarcinomas (PDAC), the procedure is a little different [363]. In this case, tumor cells are deposited
directly inside the plastic ring without previous laceration
of the CAM. Tumor nodules are invading deeply the CAM
tissue and are surrounded by blood vessels [363] (Fig. 14f).
CAM tumor samples can then be analyzed using various
methods (qPCR, Western blot, immunolabeling, etc.).
Another variant is the CAM metastasis assay [364]. This
assay can be used either to determine spontaneous metastasis
of cells deposited on the CAM or to determine experimental
metastasis after intravenous injection of tumor cells in the
chorioallantoic vein. If the metastasis assay is not strictly
speaking an angiogenesis assay, it allows, however, the study
of tumor cell dissemination through the vasculature.
16.6 Functional genomics analysis using
the chicken CAM model
Transcriptomic analysis of the CAM Transcriptomic analysis
can be performed on the CAM tissue during development,
after stimulation by exogenous factor or after tumor implantation [362, 363]. CAM tissue can either be processed as a
whole, or vessels can be micro-dissected and analyzed by
qPCR, micro-array, or RNAseq. In case of tumor implantation, the signals coming from the tumor and the stroma can
de differentiated because there is only 5% overlap. Bioinformatics analysis may help to identify the vascular cell signature of genes identified by determining vascular-specific
ESTs and an angioscore [365].
Proteomic analysis of the CAM Proteomic studies can
be done on the whole CAM tissue (snap-frozen after dissection) or can be carried out by in vivo biotinylation of
the chick embryo using shell-less embryo cultures [366].
The experimental procedures can be divided into a number
of independent steps: (1) embryo cultivation; (2) perfusion
and biotinylation of chicken embryo and its extra-embryonic vasculature; (3) lysis of selected tissue or organs and
purification of proteins; (4) deglycosylation of proteins; (5)
proteolytic (trypsinization) digestion of proteins; (5) mass
spectroscopy of peptides obtained from proteolytic digestion
of proteins and measurement of peptide masses and their
13
470
Angiogenesis (2018) 21:425–532
Table 2 Variety of applications
with the CAM
References
Developmental angiogenesis
Differentiation of vascular endothelium
Membrane proteome associated with the vasculature
Gene transfer/global gene expression
Metabolic profiling
Transcriptome analysis in the “wound model”
Vascular and endothelial cell targets from isolated chicken membranes
Lymphangiogenesis
Prox-1 in the lymphatic endothelial cells
Ingrowth of lymphatics into the tumors
Embryonic lymphangiogenesis
Vasomodulating therapies
Radiosensitizing activity
Microbeam radiation therapy
Photodynamic therapy and diagnosis
Tumor angiogenesis
Tumor growth in the CAM
Experimental metastasis
Interstitial pO2 gradients in solid tumors
Accessing molecules activity
Growth factor (receptors) inhibitors; endothelium-targeting molecules
Metal-based compounds
Inflammatory and tumor cells or purified effector molecules
Pro-angiogenic molecules
Stem cells
Human mesenchymal stem cells
Human skin-derived stem cells
Drug delivery, nanoparticles
Drug delivery for cancer treatment
Visible laser irradiation + gold nanoparticles
Screening of nanocarrier vehicles
Engineering
Tissue engineering and biomaterials
relative representation within the sample; and (6) statistical
identification of original proteins from known genetic database after calculation of nucleotide sequence from acquired
peptide masses. Also, in this case, bioinformatics analysis
may help to identify the vascular cell signature of genes
identified by determining vascular-specific ESTs and an
angioscore. A detailed protocol can be found in [367].
16.7 Limitations and challenges
The CAM assay is a model of developmental angiogenesis.
Exponential growth of new blood vessels and capillaries
occurs between embryo development days 5 and 9. For testing drugs or transplants, it should always be realized that
effects are observed in the context of this developmental
background. After Day 9, the CAM vasculature is fully
13
[352, 655]
[366]
[654–658]
[659]
[660]
[367]
[357, 661]
[662]
[663]
[662–666]
[667]
[666–676]
[348, 362, 363, 677, 678]
[677–682]
[341, 683]
[341, 684, 685]
[684–691]
[692]
[693]
[692–696]
[697]
[698, 699]
[700]
[701, 702]
[701–706]
established and allows investigation under conditions without this background angiogenesis. By evolutionary point
of view, chickens and humans are far from each other, but
a paper published by the International Chicken Genome
Sequencing Consortium [368] highlighted a high similarity
between human and chicken genome, proving that 60% of
chicken genes have counterparts in humans. Although this
might be a an argument in favor of the use of the chicken
CAM, we need to be conscious for species differences and
consider other models to validate experimental results. It
should also be realized that the chicken model is genetically
further remote from the human system, which in some cases
induces difficulties using the CAM. Although the model is
very attractive for many researchers, the chicken embryos
hatch around development Day 21. It is therefore recommended not to exceed day 18 for experimental testing. In
Angiogenesis (2018) 21:425–532
some countries, regulatory rules apply after development
day 14–15, resulting in limitations of experimental testing
for only 10–11 days. Nonetheless, there is little doubt that
the classic CAM assay has produced a remarkable array
of highly valuable information about neovascularization,
inhibition of angiogenesis, normal xenograft behavior, and
tumor growth and development. Its relative simplicity, its
avoidance of many of the regulatory rules placed on live
animal research, as well as its low cost have made it a reasonable assay for preliminary screening. In the case of proteomic analysis when using the biotinylation technique, there
may be problems with biotin penetration since biotin may
diffuse in the surrounding tissue in excess. This may lead
in picking-up proteins not strictly linked to the vasculature.
Thus, the duration of the biotinylation procedure is critical.
Harsh conditions of protein elution, which can be a problem, which may lead to contamination of the protein pool
by non-specific proteins. Despite the lack of its immunity,
chick embryo CAM does not support implants from tissues
rich in lymphocytes as colon and lymph nodes, nor implants
derived from bone marrow. This is considered a limitation
of the model, but also a challenge to find alternative ways to
use such tissues in the CAM model.
471
appears at E8.0. Vasculogenesis starts in the distal allantois
and forms a plexus that connects with the dorsal aorta of
the embryo (E8.25) before the chorioallantoic fusion (E8.5),
which is instrumental in placenta formation (E9.5). This
plexus undergoes a deep remodeling resulting in the formation of umbilical artery and vein, which invade the chorion
by sprouting angiogenesis. These processes recapitulate
the molecular and genetic distinct features of developing
vasculature in embryo and in adult life, including the role
exerted by signaling and transcriptional pathways triggered
by established angiogenic inducers and modulators [369,
370], such as VEGFs, PDGFs, ANGPTs, ephrins, NOTCH
ligands, and WNTs.
Even if less known than other angiogenesis assays, the
murine allantois explant assay represents a powerful tool to
investigate general mechanisms of blood vessel formation,
including the remodeling of a primitive vascular plexus, the
angioblast differentiation, the arterial and venous fate, the
sprouting angiogenesis, and the maturation of the capillary
network by mural cell recruitment. In particular, this assay
allowed reaching seminal contributions to vascular biology.
For example, it was exploited to investigate the role of VEcadherin, vascular endothelial protein tyrosine phosphatase,
and sphingosine-1-phosphate in angiogenesis [371–374].
16.8 The CAM as a screening platform
17.1 Overview assay
The CAM model has been used for various applications
in various fields of research. Many excellent papers and
reviews summarize these applications, showing that each
application demands the use of specific protocols (Table 2).
16.9 Concluding remarks
Even in the last 3 years, close to a thousand publications
have employed CAM assays. Whereas classically trained
embryologists are well aware of the complexity of embryogenesis and are almost overly cognizant of the problems
encountered when working with a rapidly changing target,
researchers less acquainted with the vagaries associated with
developing embryos may need to develop a grudging respect
for the intricacies underlying the CAM assay. This being
said, it is deservedly a most important technique, major contributions have resulted from its use, and no doubt it will
continue to be an important tool in angiogenesis research.
17 Murine allantois assay
The allantois is an extra-embryonic structure, which undergoes vasculogenesis, vascular remodeling, and angiogenesis
and is pivotal in establishing the chorioallantoic placenta
and umbilical circulation. The allantois is characterized by
a mesenchymal core and by an enveloping mesothelium and
The allantois is dissected from E8.5 mouse embryos using
tungsten needles and is placed individually on collagen- or
fibronectin-coated coverslips in eight-well culture dishes
(BD Biocoat). Explants are cultured in 0.5 ml of culture
medium (DMEM 4.5 g/l glucose, 10 mM L-glutamine, PenStrep), containing 15% fetal calf serum for 18 h. Explants
are then washed and fixed in 4% paraformaldehyde or
methanol:DMSO (4:1) for 20 min at room temperature and
processed for immunohistochemistry or TUNEL assay.
Pregnant female mice are killed on day 8 of gestation
(approximately E7.75) or the following day (approximately
E8.75). It is important to synchronize mice mating to dissect
the allantois at the proper stage [375, 376]. Uteri are placed
in Dulbecco’s A phosphate-buffered saline for isolation of
decidua that is placed in Hepes-buffered DME medium
with 7.5% fetal calf serum. With the aid of fine forceps and
a scalpel, the embryo is isolated from the uterine tissues
under a stereomicroscope. In the embryo, two structures
can be recognized: embryonic tissues (clear and cylindrical
shaped,) and the ectoplacental cone (reddish, cone shaped,
and located at the end of embryonic tissue in the uterine
mesometrial region). Allantoises are mouth-aspirated into a
hand-pulled glass microcapillary (60–120 μm diameter). The
ectoplacental cone/chorion should cleanly break off, leaving
an opening to the exocoelomic cavity for the easy aspiration
of the allantois. Immediately after inflating the exocoelom,
13
472
which allows better visualization of the allantois through
the yolk sac, the tip of the microcapillary is aimed toward
the distal tip of the allantois and, while gently aspirating,
“sheathe” the entire length of the allantois with the tip of
the microcapillary. While maintaining minimal aspiration,
lift the allantois, still in the exocoelom, toward the meniscus. At the meniscus, gently suction the allantois into the
microcapillary, thus leaving behind the embryo, which will
drop to the bottom of the dish. The allantois can now be
manipulated [375].
Three different methods can be used to maintain the allantois in culture. It can be kept in suspension in rolling cultures
allowing the development of a 3D spheroid characterized by
a vascularize core enveloped by a mesothelial layer. Explants
are cultured in 0.5 ml of DMEM medium containing 50%
rat serum or 5% (“low serum”) or 50% fetal calf serum.
To culture longer than 1 day, explants are given completely
fresh gas-equilibrated medium at 24 h intervals [375, 376].
A second system is based on the hanging drop system.
The allantois is suspended in 0.03 ml of DMEM supplemented with 10% FCS and located onto the underside of a
lid of a plastic bacterial dish. The hanging drop cultures are
placed into a 5% CO2 incubator, and after 18 h, the originated 3D spheroids can be resuspended in the above medium
containing angiogenic inducers (e.g., VEGF) and the hanging drop procedure is repeated [377]. Finally, allantois can
be easily maintained in adherent conditions (50% rat serum
in DMEM medium) up to 72 h on glass surface or on plastic
surface coated with fibronectin or poly-lysine and differently stimulated [370, 374, 378]. Allantois isolated at E8.0
adheres within 12 h, and after 18–20 h of culture, the explant
adopts a circular shape with a vascular plexus, which covers
the central area.
17.2 Limitations and challenges
The main positive feature of this assay is that it can analyze
vasculogenesis or angiogenesis dependent on the explant
isolation. Actually, when the allantois is isolated early (E8.0,
headfold stage), explants undergo vasculogenesis, and when
isolated later (E9.5, 22–26 somites), undergo angiogenesis
[370, 371]. Second, allantois explants can be imaged with
the use of time-lapse microscopy to follow the sequence
of events that occur during vessel formation in vitro [372].
Finally, cultured allantois explants can be immunostained for
markers of vessel formation and can be sectioned for histological analysis. It is further evident that allantois isolated
from genetically manipulated mouse models can be useful
to understand the role of specific genes in vascular development. Finally, because the allantois is implicated in placenta
vascularization, this model can be specifically exploited to
investigate pathogenetic mechanisms of placental diseases.
Negative aspects of the murine allantois assay are the short
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Angiogenesis (2018) 21:425–532
time of culture viability (72 h) and a solid experience in
manipulating embryo mice.
17.3 Concluding remarks
The allantois angiogenesis assay is not so popular and widely
used in the vascular biology community but it is useful to
discriminate if a vascular phenotype is primitive or secondary to other embryonic genetic defects, in particular when
the genetic manipulation is early lethal. The robustness of
the assay also validated by computer-assisted analysis [379]
can be exploited in drug evaluation as recently reported
[380]. Thus, the allantois is an important tool at infancy to
address relevant issues of vasculogenesis and angiogenesis
in development and disease.
18 In vivo angiogenesis plug assay
A widely used in vivo assay for the evaluation of pro- or antiangiogenic factors is the in vivo angiogenesis plug assay,
which makes use of basement membrane extracts (BME) or
Matrigel. This assay is reliable, easy to perform without the
need for special equipment, reproducible, quantitative, and
quick [381–383]. The assay is performed by injecting, for
example, 0.1 ml BME/Matrigel in the subcutaneous space of
an animal. The liquid solidifies to form a plug at body temperature. Over time, blood vessels will sprout into the plug.
With two plug sites per animal, the setup is quick and allows
for multiple test compounds or concentrations to be tested.
Thus, drug screening can also be evaluated for effects on the
activity of angiogenic or anti-angiogenic factors [384–387].
The drug can either be placed in the plug together with the
test factor by mixing with the Matrigel matrix or be given to
the host animal. Cells or even exosomes can also be tested
when mixed into the gel for their production of angiogenic
factors. Furthermore, the assay is highly versatile. For example, the role of certain genes can be evaluated using genetically modified mice (overexpressing or ablating a protein
gene), or animal models of diseases, such as aged or diabetic
animals. Also, the effect of certain drug treatments can be
evaluated. Matrigel is a mixture of basement membrane
components derived from an animal tumor [388, 389]. It
contains predominantly laminin-111, collagen IV, heparan
sulfate proteoglycans, and various growth factors. Matrigel
is a liquid at 4 °C and gels at higher temperatures. It is used
in various angiogenesis assays, including the tube formation,
aortic ring, and plug assays.
18.1 The in vivo plug assay method
In this assay, the test compound and/or growth factor is
mixed with a basement membrane extract at 4 °C, known
Angiogenesis (2018) 21:425–532
as Matrigel, and then injected, while still cold, subcutaneously into mice where it will gel. After approximately
1 week, the plug is excised and assayed for angiogenesis
by various methods, including histology, the Drabkin assay
(hemoglobin content), immunofluorescence, quantification
of fluorescent Dextran injected intravenously just before
plug harvest, etc. [381, 382, 384–387]. Complete protocols
have been published [382, 387].
The assay requires the use of BME/Matrigel without phenol and with low content of growth factors. BME/Matrigel
is thawed overnight on ice in the refrigerator, because it
gels at a temperature above 4 °C and is not usable. BME/
Matrigel can be supplemented with an angiogenic inducer,
and it is mandatory to mix it well (do not vortex as bubbles
will form) with the BME/Matrigel at 4 °C without diluting the matrix as this reduces the gelling properties. The
test mixture or the control without test materials should be
cold when injected as it will gel quickly in the subcutaneous
location due to the warmth of the animal. Injection is done
with 1-inch 21–25 g needles, which are changed after 2–4
injections as they may become dull. Mice (n = 3) are slowly
injected with 0.1 ml into both groin areas (Fig. 15) with the
tip of the needle as far as possible from the injection site to
prevent leakage. A bump should appear at the site where
the test material is released from the tip of the needle. It
is preferable to hold the syringe in place for about 30 s to
allow the test material to gel. The syringe is gently rotated to
remove the needle to help further seal the injection site hole.
At the end of the experiments, mice are killed. For the
harvest, a square segment of skin containing the plug is
excised. Alternatively, the skin can be cut around the plug
leaving plenty of space so as not to damage the plug. Then,
the underside is exposed. The plug is small and yellow in
color but it may also be pink or red depending on the degree
of angiogenesis. The control plug depending on the assay
method will be colorless. The plug is safely and gently
removed with scissors, embedded in HistoGel, and fixed for
histology, followed by sectioning and staining with Masson’s trichrome. Most of the angiogenesis will be from the
edges of the plug, and the quantitation can be based on the
density of the vessels and/or extent of in growth toward the
center of the plug. At least three fields per plug located at
approximately the same distance from the edge of the plug
are analyzed. Alternatively, Drabkin reagent can be used to
assess to the amount of blood in the plug or fluorescein isothiocyanate dextran (Mw 150,000–200,000) can be injected
into the tail vein of the animals and extracted from the plug
for quantitation. However, these approaches can be flawed by
either the presence of compressed non-perfused vessels or
by the presence of leaky blood vessel or hemorrhagic areas.
Some researchers isolate RNA from the plugs and use qPCR
of EC genes [390] as a quantitation assay.
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18.2 Key steps in 3D imaging and analyses
Matrigel is very amiable to a variety of fluorescence labeling
techniques and generation of 3D images, due to being optically translucent and highly permeable. Intravenous injection
of large (200,000 Mw) fluorescein isothiocyanate (FITC)labeled dextran identifies functional vessels within Matrigel,
while smaller (4400 Mw) tetramethylrhodamine (TRITC)labeled dextran can be used to highlight the permeability of
these vessels. Figure 14 shows an example of differences in
vascular permeability detected with this method. Stable vascular networks with low permeability are induced by FGF-2
(Fig. 14b), whilst highly permeable vessels are induced by
VEGF (Fig. 14c). To achieve these images, labeled dextrans
at 10 mg/ml each are mixed in a 1:1 ratio in 100 µl, and
injected i.v. 15 min prior to euthanasia. Excised Matrigel
plugs can be fixed in 4% PFA for 2 hours and then stored in
70% ethanol. Using a scalpel, all skin or connective tissue is
removed and then Matrigel plug slices of ~ 300–500µm are
prepared for whole-mount imaging using confocal microscopy. Antibody labeling, such as VE-cadherin (clone BV13,
eBioscience) and endomucin (clone V7C7, Santa Cruz) can
be performed over-night using a permeabilization buffer
containing 1 % BSA, 0.5 % Triton X-100. High resolution
imaging is carried out using confocal microscopy, to acquire
a series of Z stacks, which can be used to generate 3D projections using software such as Volocity. Permeability can
be identified by the presence of extravasated TRITC-dextran,
which is quantified and normalized against FITC-dextran
[391].
18.3 Limitations and challenges
As the age and gender of mice can influence the results of
this assay, it is important to standardize these parameters.
Young C57BL/6 mice are recommended as Matrigel is
derived from this mouse strain. Immunodeficient mice can
also be used (when heterologous cells are mixed in the gel),
but are more costly. It is important to note that there can
be considerable variability in the plug responses under the
same treatment; therefore, the use of 3–6 mice per data point
is recommended, with two injections in each mouse. For
pro- and anti-angiogenic agents that are tested as a local
application in the gel, it is important to establish their residence time in the gel. Some known pro-angiogenic factors
are known to leak out too fast to be effective.
The Matrigel should be handled according to the instructions. If the Matrigel has solidified or contains particulate
matter after thawing, it should not be used, because this may
interfere with the even distribution of the test material and
can yield artefacts. When mixing the test material with the
Matrigel, it is not recommended to vortex, as bubbles may be
formed that will distort the gel in vivo. Letting the mixture
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Angiogenesis (2018) 21:425–532
Fig. 15 In vivo BME/Matrigel
plug assay in mice. Injection
of BME/Matrigel in the groin/
abdomen areas of a mouse. The
left image is a plug without
growth factors; the right image
represents a plug with an
angiogenic growth factor. b, c
Confocal microscopy images
of whole-mount Matrigel
plugs, with labeled dextran
tracers. Matrigel plug angiogenesis induced by (b) FGF-2
resulted in stable vessels, with
co-localization of large FITCdextran and small TRITCdextran within the vessels,
which appear as yellow (XYZ
planes). Matrigel plug angiogenesis induced by (c) VEGF
resulted in highly permeable
vessels, with most TRITCdextran extravasated outside the
FITC-dextran positive vessels.
Images were captured using a
Carl Zeiss LSM510 META or
LSM 780 confocal microscope.
Volocity® software (Perkin
Elmer) was used to reconstruct
3D images of the vessels from
serial Z-sections
stand on ice for 10 min will reduce bubbles. Diluting the
Matrigel matrix with too much test substance volume will
reduce the gelling capacity and therefore the ingrowth of
blood vessels. Purchasing Matrigel that is at least 12–14 mg/
ml will reduce this problem. A common problem may be that
in certain cases the plug may be hard to see when harvesting.
If this happens it may be best to take the larger area of skin
around the injection site for histology. It should be noted that
13
fixation can affect some antibody epitopes therefore, frozen
sections may also be considered. HistoGel (Thermo Scientific) is recommended for preservation of the fragile plug
before fixation. Cracks or bubbles in the matrix may require
selection of areas in the plug that are clear of these artifacts.
An adaptation of the assay is an easy to use kit: DIVAA,
Directed In Vivo Angiogenesis Assay, Trevigen Inc., that
avoids many of the injection and artifact problems [392].
Angiogenesis (2018) 21:425–532
18.4 Concluding remarks
Among the various in vivo angiogenesis assays, the plug
assay is the most widely used because it is the most versatile,
least costly, and easiest to perform. Furthermore, results are
available within a week. Care must be taken at all steps in
the assay, including in the selection of the recipient mice,
handling and injecting the Matrigel, harvesting the plug, and
counting the vessels.
19 In vivo vascular network forming assay
In this assay, human endothelial cells (ECs) and mesenchymal cells are combined to form perfused vascular networks as rapidly as 4 days after subcutaneous injection into
immune-deficient mice. Thus, this assay provides an in vivo
system to study human blood vessel formation and function with the option to manipulate the human cells in vitro
prior to implantation. Briefly, human ECs are suspended in a
liquid extracellular matrix (ECM) or hydrogel with supporting mesenchymal cells and the mixture is then subcutaneously injected into immune-deficient mice (Fig. 16a). The
selected ECM or hydrogel should be compatible with vascular morphogenesis and should ideally polymerize soon after
injection to confine the cells. The vascular networks formed
within the implant are lined with the human ECs and surrounded on the abluminal side by human perivascular cells
[393–396] (Fig. 16b). This assay is simple and essentially
relies on three key components: (1) human ECs, (2) supporting mesenchymal cells, and (3) hydrogel. Nevertheless, the
nature of these three elements can vary considerably, conferring wide versatility to the assay. For example, the human
ECs can be derived from progenitor cells (e.g., endothelial colony-forming cells (ECFCs) isolated from umbilical
cord blood or adult peripheral blood), from primary tissues
(e.g., human umbilical vein ECs (HUVECs), human dermal
microvascular ECs, or white adipose tissue-derived ECs), or
from differentiated human embryonic or induced pluripotent
stem cells [393, 394, 397, 398]. The supporting mesenchymal cells may originate from a plethora of sources as well,
including progenitor cells (e.g., mesenchymal progenitor or
stem cells (MPCs, MSCs) from bone marrow, cord blood
[394, 399], or adipose tissue [400]), and primary cells (e.g.,
saphenous vein smooth muscle cells (HSVSMC) [393] and
human dermal fibroblasts [401]). Moreover, murine sources
of mesenchymal cells can alternatively be used, including
MSCs from bone marrow, white adipose tissue, skeletal
muscle, and myocardium [402] as well as the murine embryonic mesenchymal precursor 10T1/2 cell line [403]. Lastly,
this assay is compatible with a variety of ECMs and hydrogels that support formation of human vascular networks
including Matrigel [393], Type I collagen [395], fibrin [395],
475
PuraMatrix [395], and photo-crosslinkable methacrylated
gelatin (GelMA) hydrogels [404]. In summary, this two-cell
model for building human vascular networks in the mouse is
versatile, rapid, and relatively simple to perform.
19.1 Background/history
Seminal studies by Schechner and colleagues [405, 406] set
the stage for the assay described above. They suspended
HUVECs in collagen/fibronectin gels wherein the HUVECs
formed tubular structures in vitro within 20 h. They then
placed the cell-laden gel pieces by incision into the abdominal wall of severe combined immune deficiency (SCID)
mice. Human EC-lined vessels were detected in the implants
by 30 days, although the investigators found that HUVECs
needed to be transduced with the anti-apoptotic gene Bcl-2
to achieve meaningful survival. After 60 days, basement
membrane deposition, EC-EC junctions, recruited host
α-smooth muscle actin + mural cells, and TNF-α induced
E-selectin and VCAM-1 were evident [406]. In summary,
HUVECs formed well-developed, functional vessels, albeit
slowly. The next logical step was to supply mural cells 1)
to avoid reliance on host mural cell recruitment and 2) as
a source of growth factors for the implanted ECs. In this
regard, including the murine mesenchymal precursor cell
line 10T1/2 was shown to significantly increase vessel density beginning at day 14 after implantation [407]. In summary, these early studies represented a “tissue-engineering”
proof-of-concept that collectively demonstrated that preassembled human vascular networks transplanted into mice
were able to connect with host vessels.
19.2 Assay overview
We adapted these concepts to develop a rapid, simple, and
non-surgical approach for the vascular network forming
assay. Our prototypical assay consists of 2 × 106 human
ECFCs + human bone marrow-derived MSCs combined
at a ratio of 2:3 and suspended in 200 microliters of icecold Phenol Red-free Matrigel for each implant. The cell/
Matrigel suspension is then injected subcutaneously using a
26 gauge needle; two implants are placed on each dorsal side
of 6–8-week-old athymic nu/nu mice. Soon after implantation, Matrigel forms a gel at 37 °C such that cells are confined at the site of injection. Perfused vessels are detected
as early as day 4, have a robust presence by day 7 (~ 100/
mm2), and are long lived [394, 396]. Inclusion of 1 µg/ml
basic fibroblast growth factor (bFGF) in the cell/Matrigel
suspension significantly increases the number of perfused
vessels detected on day 4 [408]. Negative controls usually
include injecting the ECM or hydrogel without cells or with
ECs alone. Detailed descriptions of the methods including
endothelial and mesenchymal cell isolation procedures are
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Fig. 16 In vivo vascular network formation assay. a Schematic of the two-cell model. b
Tail vein injection of rhodamine-UEA-I and FITC-GS-B4
labels perfused human (red) and
murine (green) vessels on day
7 after cell/Matrigel subcutaneous injection. c) Quantification of lectin-labeled human
and murine vessels shows that
perfused human vessels present
at day 5 and day 7
provided in these publications from our laboratories [95,
394, 408]. Also, a detailed video protocol of this assay is
available [409].
19.3 Visualizing and quantifying vessels
At desired time points, the cell/hydrogel implants are
removed by making a dorsal incision adjacent to the implantation site to expose the subcutaneous space. The implants
13
are 3–5 mm in diameter and easily distinguishable from surrounding subcutaneous adipose tissue by shape, consistency,
and pink/red (vascularized) color (Fig. 16). Digital photographs provide initial data on the macroscopical aspect of
the implants. The excised specimens are then fixed in 10%
neutral buffered formalin overnight, followed by paraffin
embedding and sectioning (5–7 µm sections) for histological analysis. Hematoxylin and eosin (H&E) staining is used
to identify red blood cell-containing lumens; empty lumens
Angiogenesis (2018) 21:425–532
can be counted separately to track non-perfused vascular
tubes [396]. This analysis will capture human vessels as
well as host vessels that have sprouted into the gel to form
connections with the human vessels [410]. Typically, vessels are counted in ten randomly selected areas from two
H&E-stained sections, ~ 150 µm apart, for a total of 20 areas
for each implant. Ideally, counters should be blinded to the
experimental groups to reduce potential bias. Image analysis
software such as ImageJ can also be used to facilitate the
counting. As noted above, this assay typically produces a
total vessel density between 50 and 150 vessels/mm2.
19.4 Distinguishing human from murine vessels
Three strategies have been used to visualize and quantify
human versus murine blood vessels in the implant. The first
is to use human-specific antibodies such as the monoclonal
antihuman CD31 antibody (clone JC70A, Dako, Agilent)
that has been shown by numerous groups to specifically label
human vessels in histological sections. The second approach
is to genetically label the human ECs prior to implantation
with green fluorescent protein or an equivalent fluorescent
marker using lentiviral or retroviral constructs. The supporting human mesenchymal cells can be labeled with a different
fluorescent marker for dual in vivo tracking [394]. The third
approach is to use plant lectins—Ulex europaeus agglutinin I (UEA-I) which is specific for human endothelium and
Griffonia simplicifolia isolectin B4 (GS-B4), which binds to
mouse endothelium but has no reactivity with human cells
because humans lack the α-linked galactose residue needed
for GS-B4 carbohydrate binding. For instance, tail vein
injection of rhodamine-labeled UEA-I and FITC-labeled
GS-B4 10 min before removing the implants has been shown
to effectively label perfused blood vessels and shows the
connections between human and mouse vessels within the
grafts [410] (Fig. 16b). Using this lectin approach, three
distinct patterns can be seen: UEA-I-positive human vessels, GS-IB4-positive murine vessels, and UEA-I/GS-IB4double-positive chimeric vessels; each can be quantified and
reported as vessels/mm2 (Fig. 16c) [410–412]. Alternatively,
biotinylated forms of UEA-I and GS-B4 can be used to stain
the vessels in histological sections.
19.5 Longitudinal analyses
Vascular volume within the implants can be measured longitudinally by contrast-enhanced ultrasonic imaging using
a high-resolution system such as the Vevo 2100 (VisualSonics Inc, Toronto, Canada) [396, 410]. This technique
requires imaging before and shortly after tail vein injection of an echogenic microbubble contrast agent, preferably
with a syringe pump to achieve consistent levels of contrast among the different animals. In addition, to assess cell
477
retention over time, the human ECs can be genetically modified for luciferase expression (e.g., Lenti-pUbiquitin-firefly
luciferase-GFP); this allows visualizing the location of the
human ECs over time in individual mice [394, 411, 412].
This procedure simply requires an intraperitoneal injection
of the substrate luciferin followed by live bioluminescence
imaging 30–40 min later using a system such as the Xenogen
IVIS 200.
19.6 Advantages
This in vivo vascular network model is ideally suited for
studies on the cellular and molecular mechanisms of human
vascular network formation. The assay is amendable to both
loss-of-function and gain-of-function experiments. For
example, specific genes of interest can be tested by carrying out shRNA/siRNA knockdown or CRISPR/Cas9 gene
editing in the ECs and/or the mesenchymal cells prior to
implantation. Conversely, specific genes can be expressed
using retroviral or lentiviral constructs, providing a gainof-function test strategy. Alternatively, the human cells can
be treated with drugs prior to implantation, which restricts
drug exposure to the cell of interest and avoids confounding systemic effects. These types of experiments have been
used to study human vascular tumors and vascular malformations [413–415]. Furthermore, Melero-Martin and colleagues showed that human ECs can be genetically engineered to provide controlled release of a protein of interest
from the newly formed human vascular networks. In their
study, human ECFCs that were genetically engineered to
overexpress erythropoietin (EPO) were shown to form vascular networks that increased erythropoiesis and corrected
anemia in the recipient mice [416], revealing the engineered
network’s potential to serve as a drug delivery device. An
additional advantage is that the human ECs and mesenchymal cells, as well as host cells, can be easily retrieved from
the implant at any desired time point and sorted into purified populations for molecular and biochemical analyses, or
for transplant into secondary, recipient mice. Finally, it is
worth noting that hydrogel suspensions of human ECs and
mesenchymal cells have also been shown to form vascular
networks in murine ischemic myocardium and ischemic hind
limbs, providing physiological benefit to those tissues [411,
412]. This demonstrates that the blood vessel-forming ability of the cells can be recapitulated in settings beyond the
subcutaneous implant site.
19.7 Limitations and challenges
Implanting human vascular cells necessitates the use of
immune-deficient recipient mice, which precludes the use
of many available transgenic murine models. The use of
murine cells would solve this, although obtaining robust and
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478
phenotype-stable murine ECs remains a challenge. Murine
mesenchymal cells are less problematic; murine MSCs from
four different tissue sources have already been shown to support vessel formation in vivo [402]. Therefore, it should be
feasible to switch to syngeneic murine cells for implantation into desired murine strains. Second, this assay is not
well suited for high-throughput studies because of the relatively large number of human cells required, the expense of
immune-deficient mice, and the time-consuming analytical
procedures. Alternatively, in vitro 3D microfluidic models in
which perfusable human vascular networks form can provide
a valuable intermediary between this in vivo assay and 2D
cell culture models [417].
19.8 Concluding remarks
This in vivo vascular network forming assay is technically
simple, reliable, and experimentally accessible; it provides
a functional test for assessing the blood vessel-forming
potential of human ECs in vivo. The assay allows wide versatility in the use of cell sources. The human cells can be
manipulated genetically and/or pharmacologically prior to
implantation to address a variety of questions. Human (and
murine) cells can be retrieved from the implants at desired
time points for cellular and molecular analyses. The assay
is also amendable to the use of human ECs from patients or
ECs genetically engineered to express a human mutation to
model specific vascular pathologies.
20 Developing mouse retinal vasculature—
tip cells
The developing mouse retina is an extensively used model
to study angiogenesis. Vessels originate around birth from
the optic nerve and grow radially to the peripheral retinal
margin during the first postnatal week as a flat and easily
imaged plexus. Subsequently, the superficial capillaries start
sprouting downwards to form the deep and intermediate vascular plexus. The network also undergoes some remodeling,
and in approximately the third postnatal week, all vascular
layers are completely mature. Several aspects of vascular
development can be studied in this system, such as endothelial tip/stalk behavior, sprout anastomosis, lumen formation,
vessel pruning, network remodeling, formation of arteries
and veins, and others. The model is particular useful for the
analysis of genetically altered mice.
20.1 Visualization of cellular markers and gene
expression
To appreciate spatial relationships and network topology,
the retinal vasculature is best studied in retinal whole-mount
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preparations. This, however, contains some challenges. Dissecting the retina in one piece requires practice. Furthermore, depending on fixation conditions and antibodies used,
immunohistochemistry can be difficult. The likely reason for
this is the formation of diffusion barriers at the retina–vitreous interface. Two fixation protocols have emerged minimizing these problems. The first approach uses fixation for just a
few minutes in paraformaldehyde followed by a longer incubation in ice-cold methanol (which can also act as storage
medium) [418]. Alternatively, paraformaldehyde fixation is
carried out for several hours on ice. Different antibody stains
may behave differently depending on the fixation method
used. Furthermore, in situ hybridization (ISH) can also be
used on retinal wholemounts. Here, care needs to be taken
to prevent the retina from curling up during the hybridization step [419]. By far the most simple and robust method of
staining the retinal vasculature is based on labeled isolectin
B4 (IB4), which binds a carbohydrate epitope on mouse
endothelial cells, but also on microglia and macrophages.
20.2 Analysis of tip cells
Endothelial tip cells are defined by their position at the
tips of vascular sprouts (Fig. 17). They also have long and
dynamic filopodia, migratory behavior, and low proliferative activity. On the other hand, stalk cells, following the tip
cells, proliferate and form the vessel lumen. Manipulations
that change the balance between the tip and stalk phenotype alter branching frequency of angiogenic sprouts. The
basis for this concept was set by Gerhardt et al. [420], who
used lectin labeling (Fig. 17a, b) and ISH against PDGFB
(Fig. 17c) or VEGFR2 to identify tip cells. The Notch ligand
DLL4 plays an important role for maintaining the tip cell
state and is also a tip cell marker. In the search for additional
tip cell-specific markers, whole genome genetic profiling
strategies were used [421, 422] and many other useful markers have emerged such as CD34, apelin (APLN), angiopoietin-2 (ANGPT2), chemokine receptor type 4 (CXCR4), and
endocan (ESM1) [419–423]. ESM1 is probably the most
specific tip cell marker in mouse retinal vasculature, labeling additionally only some arterial ECs, whereas labeling
of CXCR4 is high in tip cells, but also to a lesser extent
present in some EC of the vessel plexus, in arterial EC and
in perivascular cells [423].
The actin cytoskeleton provides a driving force for tip
cell movement during angiogenesis, and labeling F-actin
with phalloidin [420] is another way of assessing endothelial tip cells, especially by highlighting and counting their
filopodia. As an indirect method to distinguish tip cells from
other EC phenotypes, labeling with antibodies against the
adherens junction protein VE-cadherin, the adhesion protein PECAM1/CD31 or against proteins in the basal lamina (fibronectin, collagen, laminin) is used in combination
Angiogenesis (2018) 21:425–532
with IB4, in which all ECs are labeled, but tip cell filopodia
exclusively with IB4 [420, 424]. The use of markers of the
basal lamina is based on the concept that formation of new
sprouts requires degradation of extracellular matrix to allow
migration of tip cells, thus showing reduced staining in proximity of tip cells. Indeed, it was recently shown by triple
labeling of tip cells with F-actin, cortactin, and collagen
type IV that tip cells use so-called podosomes to degrade the
extracellular matrix [425]. Labeling with anti-Ki67 or BrdU
[420, 424] is used as marker of proliferation, a property that
is greatly reduced in tip cells.
Several transgenic mouse models are also available today
to specifically study tip cells in the mouse retina. LifeactEGFP labels actin associated with cell–cell junctions, and
apical and basal membranes, and highlights actin-based
structures such as filopodia and stress fiber-like cytoplasmic bundles [426]. Also, a Cre transgenic strain was developed (Esm1-CreERT2) [423], mimicking tip cell-specific
expression of ESM1 in the retina [421]. Combining this
transgenic strain with the R26-mTmGT/+ Cre reporter generated, a conditional knockout strain that can be cross-bred
with global knockout strains for tip cell genes as Cxcr4, or
stalk cell-specific genes as Notch1, and Jag1 to study tip
cell fate [423]. Furthermore, mosaic mutant mice can be
generated by injecting morulas or blastocysts of mice with
DsRed or eGFP expressing embryonic stem cells that have
undergone gene targeting. The ability of the mutant embryonic stem cells to contribute to EC in the tip position is a
direct readout of the importance of the mutated gene for EC
tip cell function. Mosaicism can also be achieved by using
very low amounts of tamoxifen in tamoxifen-inducible,
endothelial specific Cre transgenic strains. Interestingly,
479
even heterozygous gene deletions, with no phenotype in
globally targeted mice, can result in a competitive disadvantage in a mosaic background, resulting, for example, in a
reduced proportion of mutant EC in tip position [427, 428].
Tip cell behavior can also be studied in ex vivo and
in vitro models, which have the advantage over in vivo
models that they are suitable for an array of interventions
and readouts and allow a rapid and low-cost screening of
potential anti-angiogenic molecules. These models, such as
retinal or dorsal aorta explants, and EC-coated beads embedded in fibrin gels are well suited to study tip cells and are
described in detail in other chapters of this article. It has
recently also been demonstrated that tip cells can be identified in EC monolayer cultures such as primary HUVECs,
HMVECs and in immortalized endothelial cultures by staining for CD34, an endothelial and hematopoietic stem cell
marker, which stains the tips of vascular sprouts but also
the luminal side of all EC in vivo [429]. In contrast, in vitro
CD34 reliably and specifically stains endothelial cells with
almost all characteristics of tip cells in vivo (Fig. 17d). This
in vitro model of tip cells allows studies into the differentiation and regulation of tip cells, as well as the discovery of
novel tip cell genes [422].
20.3 Network analysis of retinal vasculature
The size and topology of the developing retinal vasculature
can be used as a general readout to assess angiogenic processes. The overall diameter or the plexus covered during the
initial expansion phase of retinal vasculature development is
widely used as an indicator of decreased or increased angiogenesis. Moreover, branching frequency, branch length, and
Fig. 17 Identification of tip
cells. The tip cell is the leading
cell of an angiogenic sprout
with long filopodia extensions,
followed by stalk cells that proliferate and phalanx cells that
form a matured new capillary.
a Tip cells in the developing
mouse retina can be identified
by staining with isolectin B4
(IB4). b The mouse retinal vasculature (IB4) follows the astrocytic meshwork (GFAP) when
forming the superficial vascular
plexus during development. c A
combination of collagen type IV
IHC (in pink) for general staining of blood vessels and ISH
for PDGFB (black) for specific
identification of tip cells in the
developing mouse retina. d Tip
cells in endothelial cell cultures
are identified by CD34
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480
vessel diameter are strongly affected by tip/stalk imbalances
and can easily be assessed by image analysis (e.g., using
tools in ImageJ or Fiji) [430]. The plexus central to the
leading front is unstable and remodeled via the pruning of
capillaries, leading to reduced vessel density. In addition,
capillary free spaces form in the vicinity of arteries. These
readouts can indicate changes in vessel stability [431]. Furthermore, artery–vein differentiation is particularly noticeable in the developing retinal vasculature because of the radially alternating patterning of arteries and veins. Arteries are
easily labeled with an antibody against alpha smooth muscle actin, which is strongly upregulated in arterial smooth
muscle cells. Endothelial specific arterial (e.g., ephrin B2 or
DLL4) and venous markers (e.g., Eph B4) can also be used.
20.4 Limitations and challenges
Retinal whole-mount IHC and ISH can be tricky, and the
accessibility of antibodies in the retina may be dependent
on the protein or mRNA targeted. Transgenic reporter mice
partly overcome these problems greatly reducing background and show a more equally distributed expression, but
are not affordable for every laboratory. A further limitation
to the use of tip cell markers is that none of the current markers exclusively labels tip cells, as they can also be expressed
in other EC of the vascular plexus, arteries, or perivascular
cells. Therefore, identification of tip cells relies mainly on
their position, morphology, and their filopodial protrusions.
To confirm the endothelial identity of tip cells, staining with
multiple antibodies is necessary to distinguish between EC
and perivascular cell types.
Another major limitation of the developing mouse retinal
vasculature as a model system is that experimental manipulations are challenging. Although intraocular injections are
possible, they are prone to artefacts and requires highly
skilled and experienced operators. Genetic models are more
robust but usually require an inducible approach (e.g., the
tamoxifen-inducible Cre-lox system) because the retinal vasculature develops postnatally and cannot be studied when
vascular abnormalities lead to embryonic death. The associated mouse breeding can be very costly and time-consuming.
Thus, ex vivo and in vitro assays may be in many instances
more appropriate as they are faster, cheaper, and ethically
more responsible. However, more complex network development depends on in vivo approaches and can so far not be
studied in vitro.
20.5 Concluding remarks
The developing retinal vasculature is a very well-defined
and widely used model to assess vascular phenotypes in
mutant mice. It has been particularly useful in the study
of endothelial tip/stalk behavior in physiological context.
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However, inducible genetic tools are usually required and
the associated mouse breeding makes it a resource and timeintensive model.
21 Corneal angiogenesis assays
In the early 1970s, the Folkman’s group firstly set up an
in vivo angiogenesis by exploiting the anatomical features
of rabbit cornea and in particular the absence of blood vessel
and the easy access to monitor the neovascularization by a
slit lamp after implanting tumor cells or tissues or slowrelease pellets containing angiogenic inducers [432]. This
strategy was further extended to rodents (mouse and rats),
which are smaller and less expensive than rabbits and allow
studying specific molecular correlations by taking advantages of using genetic engineered mouse models [433–435].
Corneal assays are instrumental to characterize angiogenic
inducers and inhibitors, interactions between different factors, and to study cellular and molecular mechanisms of
angiogenesis [436]. Interestingly corneal assay can also be
used to study other biological processes. For example, studies of lymphangiogenesis were made possible through the
implantation of low-dose bFGF pellets, which allowed the
visualization of lymphatic vessels through specific molecular markers [437].
21.1 Assay overview
The protocol is based on the creation of a small pocket in
the cornea to introduce tissue samples, cells, or slow-release
pellets incorporating growth factors. Surgical procedures are
performed on anesthetized animals, in sterile conditions to
avoid inflammatory reactions and with the help of a stereotaxic microscope. Generally, the resins used are etylenvinyl acetate (Elvax 40W, 40% by weight ethylene–vinyl
acetate comonomer content with a “W” amide additive,
Dupont) or a mixture of sucralfate and poly(2-hydroxyethyl
methacrylate) (Hydron). To produce slow-release Elvax 40
pellets, 1 g of the polymer is washed in 100 ml of absolute
alcohol for at least 15 days and then dissolved in 10 ml of
methylene chloride to prepare 10% casting stock solution
(60 min in at 37 °C). Two hundred microliters of this solution is layered on a Teflon film, and the methylene chloride is
allowed to evaporate under a laminar flow hood. The resulting film of polymer is cut under a stereomicroscope into
1 × 1 × 0.5 mm pieces, which are used as implants. Ten
implants (see above) are monitored for 14 days after implantation. The casting solution is eligible for the use when the
implants performed with a specific casting solution do not
induce reactive inflammatory injury at microscope and histological examination of the cornea. To incorporate the test
substance, a predetermined amount of casting solution is
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mixed with a given amount of the molecule and processed
as above described. Alternatively, pellet is incubated with
a solution of the testing molecule dissolved in sterile phosphate buffer solution for 40 min at room temperature in a
plate rotator and then dried in a laminar airflow hood under
sterile condition.
Elvax-40 containing testing molecules can be stored up
to 1 month at − 80 °C. Empty pellets of Elvax-40 are used
as negative control, while positive control contains VEGFA (200 ng/pellet). In the case the experiment protocols
require the use of cells or tissues, it is used to implant 5
μl of medium containing 2–5 × 105 cells, or a fragment of
2–3 mg of fresh tissue.
A second resin is hydron stabilized by sucralfate. Ten
milligrams of sucralfate is added to solutions of sterile saline
(20–50 μl) containing the appropriate amount of testing substances (e.g., 0.05 mg VEGF-A as positive control) that are
then speed vacuumed until the mixture is dry (30–60 min).
To this suspension, 10 μl of 12% hydron in ethanol is added.
This suspension is layered on a sterilized 1.5 × 1.5 cm piece
of nylon mesh (pore size 0.4 × 0.4 mm). Both sides of the
mesh are covered with a thin layer of hydron and allowed
to dry in sterile conditions. Then, the fibers of the mesh are
pulled apart under a microscope, and only uniformly sized
pellets of 0.4 × 0.4 × 0.2 cm are selected for implantation.
All procedures are performed under sterile conditions. Such
pellets can be stored at − 80 °C for 1 month without loss of
bioactivity.
Pellets are implanted in anaesthetized animals, and the
eye is further anesthetized with few drops of local anesthetic.
A central incision (1.5 × 3 mm in rabbit; 1.5 × 1.5 mm in
mouse or rat) is produced into the corneal 3 mm (rabbit) or
1–1.5 mm (mouse, rat) from the limbus, and using a von
Graefe knife (2 × 30 mm), a micropocket is dissected to
insert the pellet with forceps. This procedure allows the diffusion of test molecule in the tissue, with the formation of
a gradient for the ECs of limbal vessels. The distance of
the pocket from the limbus can influence the angiogenic
response, which depends on the features of the angiogenic
inducer [438].
Observations of the implants are made with a slit lamp
stereomicroscope over a set period of time on anaesthetized
animals. Images are recorded for subsequent analysis. In
rabbit, an angiogenic response is positive when budding
of the vessels from the limbal plexus occurs after 3–5 days
and capillaries progress to reach the implant in 7–10 days.
Implants with an inflammatory reaction or that are unable
to induce angiogenesis within 10 days are discarded. The
angiogenic activity is evaluated on the basis of the angiogenic score calculated by the formula vessel density/cornea
x distance from the limbus [439, 440]. The number of positive implants over the total number is also scored.
481
In the mouse or in the rat, the vascular response is measured as the maximal vessel length [441, 442]. By using
the slit lamp microscope with an ocular grid, the y-axis of
the reticule is located along limbal vessel directly beneath
implant. It is possible to refer the eye as a clock with intervals from 1 to 12, and the measurement of the number of
clock hours with new vessels is performed during each
observation. Two measurements are obtained for each eye:
the linear response representing the maximal vessel length
(an average of the five longer vessels) extending from the
limbus toward the implant and the circumferential response
representing the neovascularization zone (measured in clock
hours). The vessel area is calculated according to this formula: vessel length × clock hour × 0.2π. At the end of the
experiments, cornea is fixed and analyzed for the presence
of inflammatory cells and to investigate the vessels features.
21.2 Limitations and challenges
Corneal assays present advantages and disadvantages in
different species. The rabbit’s size enables an easy manipulation of both whole animal and the eyes, and the inflammatory reactions are easily detectable as corneal opacity.
However, the experimental cost is higher in rabbit than in
mouse or rat. Multiple observations are easier in rabbit than
in mouse or rat and can be done in not anaesthetized animals. Furthermore, the angiogenic response in mouse and
rat is highly variable and a large number of animals are
required. Thanks to its size, the rabbit eye offers a large area
for the placement of stimuli in different forms including the
presence of two pellets. All species can be also used to test
the effect of local or systemic drug treatment on the local
angiogenic response. Obviously, the systemic drug treatment
of rabbit is more expensive than the treatment of mouse or
rat. There are some critical steps in performing a successful
corneal assay. The first issue is the preparation of pellets,
which need to be uniform, without any inflammatory activity and able to ensure a good distribution of the angiogenic
molecule. Surgical procedure is another challenge issue of
the assay, and it is important to calibrate the appropriate
depth of the incision to avoid the eye rupture. In mouse,
this assay permits to induce neo-angiogenesis on different
genetic backgrounds, which can be useful to evaluate the
effect of a specific gene on angiogenesis.
21.3 Concluding remarks
Corneal angiogenesis assay, which requires high technical
competences, represents a powerful tool to monitor and
quantify in vivo neo-angiogenesis, to test specific drug treatment and the effect of genetic manipulation. To possibility of
final histological examinations allows an initial description
of the cellular mechanisms sustaining the process.
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22 Mouse oxygen‑induced retinopathy
model
Pathological angiogenesis plays a pivotal role in the most
common causes of blindness including neovascular agerelated macular degeneration (AMD) [443], polypoidal
choroidal vasculopathy (PCV) [444], or diabetic retinopathy
and retinopathy of prematurity (ROP) [445]. Animal angiogenesis models are an essential tool to understand pathways
involved in physiological and pathological angiogenesis for
basic comprehension of the biology and to determine the
basis for in vivo anti-angiogenic drug development in translational cancer and ophthalmic research. Moreover, they are
critical in preclinical drug testing.
The oxygen-induced retinopathy (OIR) model was originally developed to reflect the pathological stages of human
ROP, vaso-obliteration of retinal vessels with oxygen exposure (phase I), followed by vaso-proliferation (phase II).
Oxygen exposure of neonatal dogs, cats, rats, and mice creates a relative hypoxic state in the retina with stimulation of
growth factors resulting in eventual vaso-proliferation as is
seen in preterm infants with ROP [446]. A major distinction
of this model is that mice, rats, dogs, and cats are born with
incompletely vascularized retinas, similar to humans born
preterm allowing the study of retinal vascular development.
Mice are the least expensive and most genetically manipulative animal model and therefore the most commonly used.
Murine retinal angiogenesis occurs postnatally in a tightly
regulated, predictable manner similar to humans allowing
for manipulations seen postnatally in preterm babies such
as oxygen exposure [445].
Since the introduction of the current mouse OIR model
in 1994, it has been extensively used in many studies of
angiogenesis and retinopathy. A Google Scholar search with
the keywords “oxygen-induced retinopathy mice” revealed
approx. 45,000 publications with an increase of 9500 publications in the last year, reflecting the feasibility and practicality of this in vivo model in many areas of vascular biology
research. The major advantages lies in its reproducibility,
affordable costs, short duration (10 days), and accessibility
of the retinal vasculature for isolation, imaging, and interventions [447, 448]. Further, the well-established platforms
for creating systemic or conditional transgenic strains and
access to a wide availability of recombinant proteins and
antibodies make it a powerful tool for basic angiogenesis
research.
Angiogenesis (2018) 21:425–532
postnatal day (P)7 until P12 and returned to room air (21%
oxygen) from P12 to P17 (Fig. 18a). Due to the high pO2
levels during the first hyperoxic phase (P7–P12), retinal vessels stop developing and immature capillaries in the central
retina regress resulting in a central zone of vaso-obliteration
(Fig. 18a, b′) [450]. Interestingly, the oxygen-induced vasoobliteration (VO) develops very rapidly with peak VO 48 h
after onset of oxygen exposure [446]. The retina starts revascularizing slowly under remaining oxygen exposure from
P9 to P12 [446, 451], likely reflecting increasing oxygen
demands of the developing retina. On return to room air at
P12, the VO area becomes hypoxic [452] and significant
upregulation of HIF-1-dependent, pro-angiogenic pathways including VEGF ensues resulting in neovascularization (NV) with its peak at p17 [453–455] (Fig. 18a, b″). A
comprehensive step-by-step protocol of this mouse model
has been published [456].
The two main quantifiable components in the OIR mouse
model are the NV (maximum at P17) and VO areas. If a
study is designed to determine the amount of vasculature lost
during the hyperoxic phase, the VO area can be measured at
P12 or even as early as P8 by manually outlining the VO in
an image software such as Adobe Photoshop (Fig. 18c). In
most cases, NV and VO are measured at P17 when abnormal
vessel growth is at its peak. It has to be kept in mind that
the VO area on P17 can differ between experiments with
same VO area at p12 depending on the extent of vascular
repair, which can be determined by examining the difference between VO at P12 and P17. The current method to
quantify NV is the SWIFT_NV method, which is a computer-aided semiquantitative technique detecting NV tufts
and clusters on retinal wholemounts using the free NIH software ImageJ [449]. In brief, using different macros, retinal
flat mounts are divided into four quadrants and background
fluorescence is removed allowing NV structures to stand out
clearly (Fig. 18c′, c″). This enables the user to set a fluorescent threshold marking NV, but not normal vessels. Artifacts can be manually excluded during this step. Recently,
a novel Web-based image analysis tool (http://oirseg.org/)
has been developed based on deep learning segmentation,
which greatly facilitates the quantification of NV structures
[457]. This method greatly improved efficiency, reliability,
and objectivity compared to the prior methods such as crosssectional techniques and whole-mount grading systems
based on predefined characteristics.
22.2 Limitations and challenges
22.1 Assay overview
The current OIR model was developed by Smith et al. [446]
and optimized by Stahl et al. in 2009 [449]. In brief, neonatal
mice with nursing mothers are placed into 75% oxygen from
13
Despite standardization of the OIR mouse model over the
last 23 years, several essential factors can result in substantial phenotype variability if not controlled. One of these factors is postnatal weight gain (PWG) during the OIR period.
A study from Stahl et al. [458] revealed that pups with poor
Angiogenesis (2018) 21:425–532
483
Fig. 18 Mouse model of oxygen-induced retinopathy (OIR). a The
mouse model of oxygen-induced retinopathy (OIR). Neonatal mice
and their nursing mother are placed into 75% oxygen from P7 to P12,
which induces loss of immature retinal vessels, leading to a central
zone of vaso-obliteration (VO). After returning to room air at P12,
the central avascular retina becomes hypoxic, inducing vascular
regrowth with pathologic neovascularization (NV). At P17, the maximum severity of NV is reached. NV starts to regress shortly after P17
and almost no VO or NV remains visible by P25. b Retinal whole
mount stained with isolectin-B4-Alexa (red) displaying a normal
vascular development at P17 under normoxic condition. b′ OIR P12
retinal whole mount showing an extensive VO area without NV. b″
OIR P17 retinal whole mount showing a decreased VO with NV at
its maximum. c Quantification of vaso-obliteration (VO) by manually
outlining the avascular area with image-processing software (Photoshop, Adobe Systems) c′ For computer-aided NV quantification, both
the original image and the VO image generated with Adobe Photoshop were imported into NIH’s free-access ImageJ software. The
SWIFT_NV macroset isolates the red color channel, subtracts background fluorescence, and divides the VO image into four quadrants.
c″ SWIFT_NV then allows the user to outline NV tufts but not normal vessels by setting a fluorescence threshold for each quadrant. The
macroset then quantifies all NV pixels from all four quadrants, reports
the result as neovascular total area, and creates an overlay of NV and
original image
postnatal weight gain (PWG) (defined as pups weighing 5 g
or less at P17) have a significantly delayed and prolonged
vaso-obliteration and NV compared to medium (5–7.5 g)
and extensive (≥ 7.5 g) PWG. Therefore, the weight of each
pup at P7 and again at P17 should be measured. Mice below
6 g and over 7.5 g at P17 should be excluded from the analysis. To avert poor PWG during OIR, several adjustments can
be considered. We recommend avoiding large litter sizes
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484
over eight pups by removing some pups (or if pups are too
heavy by adding pups to maintain appropriate weight gain)
and surrogating C57BL/6 pups at P2 or P3 with S129 lactating mice due to their lower susceptibility to oxygen toxicity
if weight gain is low. It is important to note that nursing
mothers are no longer fertile after exposure to OIR. Under
hyperoxia exposure and especially after transfer to normoxia,
we found that mothers cannibalize their pups. Therefore,
we recommend careful handling after return to normoxia as
stress leads to less lactation and increased cannibalization.
Another important variable, which needs to be considered, is the phenotype variability between different strains
and even within one strain due to vendor-related substrain
differences. These differences can be easily addressed by
using the same strain from a single vendor. To correct for
age and genetic heterogeneity as well as environmental differences, the use of littermate controls is strongly recommended. When using transgenic mice, controls with exactly
the same background will help to minimize strain-dependent
phenotype variability.
Last, it has to be noted that the OIR mouse model does
not cover all aspects of human ROP since mice are not born
preterm even though retinal development occurs postnatally
as it does with preterm infants. Thus, systemic factors resulting from preterm delivery are not taken into account, such
as low IGF-1 seen after preterm birth in humans. Also, mice
have no macula (the area of acute central vision) and have
fewer cones than humans. However, vascular development
is very similar.
22.3 Concluding remarks
The OIR mouse model is a robust, reliable, and quantifiable
model to investigate developmental and pathological angiogenesis when variables are carefully controlled.
23 Laser‑induced choroidal
neovascularization mouse model
Age-related macular degeneration (AMD) is one of the
major causes of vision impairment in the elderly [459].
Although AMD does not lead to complete blindness, loss
of central vision makes it difficult for patients to recognize
faces, drive, or read. Nonexudative or non-neovascular AMD
includes early and intermediate forms of AMD, as well as
geographic atrophy, in which progressive loss of retinal
cells leads to some loss of visual function. In 10–20% of
AMD patients, nonexudative AMD progresses to neovascular AMD, which accounts for ~ 90% of AMD-associated
vision loss with deterioration of central vision [460]. Neovascular AMD is characterized by choroidal neovascularization (CNV), with blood vessels from the choriocapillaris
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penetrating through Bruch’s membrane into the normally
avascular subretinal space [461]. However, in some patients,
pathological angiogenesis develops from retinal vessels
known as retinal angiomatous proliferation (RAP). In vitro
EC culture models of CNV lack complex in vivo cellular
interactions of photoreceptors, retinal pigment epithelium,
pericytes, inflammatory cells, and glial cells [462]. Therefore, reproducible animal models that mimic the pathological angiogenesis in the retina and choroid are needed to
study AMD [463]. No models, however, mimic the aging
aspects of the disease, nor do non-primates have a macula.
23.1 Laser CNV model advantages and limitations
A laser-induced in vivo model of CNV, first described in
1979, uses photocoagulation to disrupt Bruch’s membrane,
inducing the growth of new choroidal vessels into the subretinal area [464]. This model is similar to the majority of
neovascular AMD in which pathological angiogenesis arises
from the choroid. However, it differs from AMD as it is
a wounding model unlike neovascular AMD that is initiated with aging changes. The laser-induced CNV model
has been used successfully to predict the clinical efficacy of
anti-vascular endothelial growth factor (VEGF) therapy to
suppress neovascular growth in AMD [465]. Laser-induced
CNV has been used in mice, rats, rabbits, pigs, and nonhuman primates. Non-human primates’ retinal anatomy
(with a macula) is most similar to humans, but primates
are costly to maintain, and rarely develop AMD and only
do so after decades. Rodent models offer the advantages
of lower cost but lack an anatomical macula [466]. Several
gene-targeted mouse models, such as the very low-density
lipoprotein receptor knockout (Vldlr −/−), superoxide dismutase (SOD) knockout (Sod1 −/−), and apolipoprotein E
(APOE) e4 transgenic mice, demonstrate some aspects of
CNV such as spontaneous subretinal neovascular lesions
[462]. However, these models are difficult to further manipulate genetically to determine pathways involved in AMD
progression, whereas the laser-induced CNV model can be
used in transgenic animals to explore the molecular mechanisms of CNV formation.
C57BL/6J mice are usually recommended for laserinduced CNV experiments, because only pigmented mice
absorb laser energy well and respond reliably to laser burns.
Previous studies suggest that both gender and age of animals influence the outcome of laser-induced CNV [467].
In one study, mice of both genders more than 12 weeks old
developed more severe CNV than mice less than 8 weeks
old. Gender difference was only significant in 12–16-weekold mice, but not in the younger 6–8-week-old mice [467].
However, Zhu et al. [468] observed larger CNV lesions in
female mice at 5–8 weeks old. This discrepancy may be
due to differences in analysis time points and fluorescent
Angiogenesis (2018) 21:425–532
methods between the studies. Especially noteworthy, the
older female mice developed significantly larger CNV
lesions than both older male and younger female mice. The
larger area of CNV in older female mice is suggested to be
related to their high circulating levels of estrogen, which
increases pro-angiogenic functions of both EC and smooth
muscle cells in vivo and promotes wound healing in both
human and animal models. Moreover, compared with the
younger mice, the lesion area in the older mice had increased
variability. These studies suggest that mice of either gender weighing 15–23 g at 6–8 weeks of age are optimal for
the laser-induced CNV model for testing efficacy of drugs,
although age- and gender-matched mice may be essential for
specific experiments.
23.2 Assay overview
The general procedure of laser-induced CNV induction
involves careful mouse anesthesia, mouse positioning,
laser burn, with optional optical coherence tomography
and fundus fluorescein angiography, eye dissection, choroid staining and imaging, and CNV lesion quantification
(Fig. 19a). Only intact eyes (Fig. 19b) without observable
structural or morphological abnormalities are used for the
laser-induced CNV model. Eyes with anomalous structures
(Fig. 19c), cataract, or visible defects of the cornea or fundus are excluded. Mice are anesthetized with a mixture of
xylazine and ketamine, and pupils are dilated with topical
drops of Cyclomydril. 2 min after pupil dilation, lubricating
eyedrops are applied to the cornea. Four laser burns at equal
distance from the optic nerve (which optimally is approximately twice the diameter of the optic nerve) are induced
one by one in each eye by a green Argon laser pulse with
a wavelength of 532 nm (Fig. 19d). The distance between
laser burns must be at least double the diameter of the optic
nerve to avoid fusion of lesions. Major retinal and choroidal vessels should be avoided to prevent potential bleeding.
The formation of a vaporization bubble immediately after
laser photocoagulation indicates the success of a laser burn,
which correlates with rupture of Bruch’s membrane. Optical coherence tomography immediately after laser photocoagulation may be used to confirm the success of the laser
burn with visible rupture of Bruch’s membrane (Fig. 19e).
After laser photocoagulation, the eyes are gently rinsed with
sterile saline to remove the lubricating eyedrops and treated
with an antibiotic ointment, erythromycin. Mice are then
placed on a pre-warmed warming plate at 35 °C after the
laser treatment until they are fully awakened. Mice with or
without treatment can be subjected to fundus fluorescein
angiography to evaluate the levels of vascular leakage from
CNV lesions 6 days after laser burn (Fig. 19f, g). The in vivo
retinal structure may also be examined by optical coherence
485
tomography, if applicable, to determine the cross-sectional
area of CNV lesions 7 days after laser burns. To measure the
surface area of CNV lesions, the fluorescence-stained retinal
pigment epithelium/choroid/sclera flat mounts are imaged
(Fig. 19h, i) and quantified by researchers masked to treatment. The choroidal CNV samples may also be analyzed for
RNA or protein.
23.3 Limitations and challenges
The laser-induced CNV model in mice has been often characterized as variable and inconsistent. Establishing a set of
consistent exclusion criteria is necessary for ensuring reliable data analysis. In a typical study, ten mice per group
with four lesions per eye would optimally provide 80 data
points for each experimental condition. To account for data
or mouse loss, including (1) cataract and corneal epithelial edema before laser photocoagulation, (2) unsuccessful
laser burn without Bruch’s membrane rupture, (3) odd lesion
shape due to mouse movements during laser induction, (4)
death of mice post-laser treatment, or (5) damage of the
CNV lesions during tissue dissection and processing, more
mice may be needed and should be considered in a power
analysis to account for an anticipated intervention effect.
To accurately evaluate the laser-induced CNV, some
lesions should be excluded. Severe hemorrhages will
cause much larger CNV lesions, whereas choroidal damage will yield a CNV lesion much smaller than the fellow
CNV lesions in the same eye. First, choroidal hemorrhages
encroaching on the lesion should be analyzed and classified
carefully: (1) If the diameter of the bleeding area is less than
that of the lesion, the lesion will be eligible for inclusion of
analysis; (2) if the diameter of the bleeding area is more than
that of the lesion but less than two times of the lesion diameter, the lesion should be excluded from quantification; and
(3) if the diameter of bleeding area is more than two times
the lesion diameter, all lesions in the same eye should be
excluded from analysis. Second, excessive laser burns that
damage not only Bruch’s membrane but also the choroid and
retinal pigment epithelium should be excluded. These excessive burns can be seen clearly as a solid “hole” in the bright
field of choroid imaging. Lesions should also be excluded
if (1) the lesion is fused with another lesion, (2) the lesion
is either more than five times larger than the mean of the
lesions under the same experimental conditions, or (3) the
lesion is the only one eligible for statistical analysis among
all lesions in an eye. Previous studies provide the optimal
settings and conditions to make use of the laser-induced
CNV model for the goal of improving the consistency and
reproducibility of experimental results for AMD and other
pathological angiogenesis research [467].
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Angiogenesis (2018) 21:425–532
Fig. 19 Experimental flowchart of the image-guided laser-induced
CNV model and data collection. a Overview of the procedure for
CNV induction involving mouse preparation and followed by experimental treatment, sample preparation, and analysis. b Representative
image of normal fundus (Green check mark). c Representative image
of anomalous structure (white arrow) in the eye, which is not suitable for laser photocoagulation (Red X). d Representative image of
normal fundus with four laser burns shown as bright white spots. e
Representative image of a successful laser burn (white arrow) with
3D optical coherence tomography. f, g Representative ocular fundus
fluorescein angiography images at 5 and 10 min after the injection of
fluorescent dye at day 6 after laser burn. h Representative images of
flat-mounted choroid with IB4 staining at day 7 after laser photocoagulation. Scale bar: 200 μm. ON: optic nerve. (i) Higher magnification
of the laser-induced CNV lesion highlighted in panel H. Scale bar:
50 μm. Reproduced with permission from [467]
23.4 Concluding remarks
24 Transparent window preparations
for angiogenesis studies in mice
The laser-induced choroidal neovascularization model is an
attractive model to study AMD. The model is available in
many species and is, despite the fact that the initiation of the
pathology is through wounding and not because of aging,
suitable for the development of drugs for the treatment of
AMD.
13
Our ability to observe living tissues with the microscope
improved dramatically in the 1920s when Sandison began
implanting transparent windows over wounds in rabbit ears.
This allowed him to view the underlying tissue noninvasively and longitudinally [469, 470]. In the 1940s, Algire
modified Sandison’s method to visualize the dorsal skin in
mice [471]. With this powerful tool, he was able to perform
detailed studies of angiogenesis during wound healing and
Angiogenesis (2018) 21:425–532
487
Table 3 Angiogenesis assays and vascular analysis
References
Development of novel imaging techniques
Direct measurement of interstitial diffusion and convection of albumin using fluorescence photobleaching
[707]
Tumor induction of VEGF promoter activity in stromal cells
[506]
Interstitial pH and pO2 gradients in solid tumors in vivo
[505]
In vivo measurement of gene expression, angiogenesis, and physiological function in tumors
[487]
Two-photon fluorescence correlation microscopy to reveal transport in tumors
[498]
Quantum dots to spectrally distinguish multiple species within the tumor milieu in vivo
[490]
Three-dimensional in vivo microscopy using optical frequency domain imaging
[485]
Simultaneous measurement of RBC velocity, flux, hematocrit, and shear rate in vivo
[495]
In vivo validation of MRI vessel caliber index with intravital optical microscopy in a mouse brain tumor model
[515]
Video-rate resonant scanning multiphoton microscopy
[496]
Next-generation in vivo optical imaging with short-wave infrared quantum dots
[708]
Analysis of vessel function
Regulation of transport pathways in tumor vessels: role of tumor type and microenvironment
[501]
Effect of tumor–host interactions on distal angiogenesis and tumor growth
[709]
Kinetics of vascular normalization in response to VEGFR2 blockade
[510]
pH/pO2
Calibration and application of fluorescence ratio imaging of pH gradients
[503]
Noninvasive measurement of microvascular and interstitial oxygen profiles
[504]
Simultaneous in vivo high-resolution measurements of interstitial pH and pO2 gradients
[505]
Extracellular matrix and interstitial transport
Fluorescence photobleaching with spatial Fourier analysis for measurement of diffusion
[502]
Dynamic imaging of collagen in tumors in vivo using second-harmonic generation
[491]
In vivo imaging of ECM remodeling by tumor-associated fibroblasts
[492]
Angiotensin inhibition enhances drug delivery by decompressing tumor blood vessels
[500]
Anti-VEGF therapy induces ECM remodeling and mechanical barriers to therapy
[481]
Immune cells
VEGF and bFGF regulate natural killer cell adhesion to tumor endothelium
[497]
Ly6Clow monocytes drive immunosuppression and confer resistance to anti-VEGFR2 therapy
[511]
Analysis of lymphatic vessel function
Conventional and high-speed intravital multiphoton laser scanning microscopy
[508]
Lymphatic metastasis in the absence of functional intratumor lymphatics
[710]
A genetic Xenopus laevis tadpole model to study lymphangiogenesis
[711]
Investigation of the lack of angiogenesis in the formation of lymph node metastases
[486]
Drug delivery, nanoparticles
Vascular normalization improves the delivery of nanomedicines in a size-dependent manner
[499]
Compact high-quality CdSe-CdS core–shell nanocrystals with narrow emission linewidths and suppressed blinking
[489]
Magneto-fluorescent core–shell supernanoparticles
[712]
Engineered vasculature
Tissue engineering: creation of long-lasting blood vessels
[407]
Paradoxical effects of PDGF-BB overexpression in EC in vivo
[713]
Engineered blood vessel networks connect to host vasculature via wrapping-and-tapping anastomosis
[509]
Generation of functionally competent durable engineered blood vessels from human pluripotent stem cells
[398]
Mathematical analysis of angiogenesis
Scale-invariant behavior and vascular network formation in normal and tumor tissue
[517]
Cancer, angiogenesis, and fractals
[519]
Scaling rules for diffusive drug delivery in tumor and normal tissues
[518]
Assessing therapies
Herceptin acts as an anti-angiogenic cocktail
[714]
Targeting placental growth factor/neuropilin 1 pathway inhibits growth and spread of medulloblastoma
[478]
Vascular endothelial protein tyrosine phosphatase inhibition in tumor vasculature and metastatic progression
[512]
A cerebellar window for intravital imaging of medulloblastoma in mice
[479]
Tissue isolation chambers
Implantable tissue isolation chambers for analyzing tumor dynamics in vivo
[493]
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Angiogenesis (2018) 21:425–532
Fig. 20 Tumor angiogenesis imaging. Vasculature in the brain (left) and the dorsal skin (right) visualized using IR frequencies to image deeper
into tissue; blood flow creates the contrast, so it is noninvasive (from [485])
tumor growth. Intravital imaging techniques have provided
unprecedented insight into tumor vessel formation and
function [472–474]. By implanting transparent windows in
various anatomical locations in the mouse, it is possible to
observe angiogenesis associated with tumor development,
wound healing, and immune/inflammatory responses with
high resolution, and in the native microenvironment. To
date, transparent window preparations in mice have been
developed to observe angiogenesis and lymphangiogenesis
in the dorsal skin [475], brain [476, 477], cerebellum [478,
479], liver [480, 481], lung [482], pancreas [483, 484],
breast [485], and lymph node [486], Table 3.
24.1 Development and capabilities
Advances in intravital microscopy have come with the
development of new window models, but also with innovations in microscopy. Multiphoton laser scanning microscopes provide high resolution, 3D images of vessel anatomy, gene expression, and network topology, even in deeper
regions of tumors [487]. More recently, frequency domain
optical coherence tomography (OCT) imaging has been
applied in window chambers to improve depth penetration
and contrast, providing unprecedented images of vasculature noninvasively and longitudinally [485]. Another imaging technology, short-wave infrared imaging (SWIR), has
been adapted from the defense industry to improve imaging
depth penetration and reduce scattering [488]. The development of novel probes for improving contrast has also
improved the ability to monitor angiogenesis in vivo [489,
490]. Furthermore, advanced imaging technologies such as
second-harmonic generation imaging allow us to see important components of the angiogenic process such as ECM
fibers [491]. This ability has improved our understanding
of the involvement of matrix components in the angiogenic
process [491–493].
13
Chronic window preparations allow analysis of vessel
function, including hemodynamics [494–496], immune cell
trafficking [497], transport of materials in the vascular and
interstitial spaces [472, 487, 498–501], the binding kinetics of drugs in vivo [502], and other microenvironmental
parameters such as perivascular pH and pO2 [503–505].
Using intravital reporters, it is possible to study spatiotemporal expression of genes [506], to quantify vessel structure, function, and dynamics [485, 493], and to probe the
abnormal organization, structure, and function of angiogenic vasculature (e.g., hyper-permeability, heterogeneous,
and compromised blood flow) [475, 477, 487, 507, 508].
When combined with sophisticated imaging and analysis
techniques, we can quantify blood velocity, hematocrit, and
shear rate in vivo [495].
Intravital microscopy through transparent windows has
been used to study many aspects of neovascularization and
vessel dynamics (Fig. 20). In addition to tumor angiogenesis
and wound healing, they have been valuable for understanding the integration of engineered tissue grafts into existing
vascular beds [398, 407]. By following various cell populations as angiogenic host vessels make new connections with
those of the graft, it is possible to understand the cellular
mechanism of anastomosis [509].
Intravital imaging also allows assessment of therapies that
affect the vasculature. For example, it is possible to quantify
the abnormalities in structure and function that are hallmarks
of tumor angiogenesis, and then assess how these abnormalities change when the vasculature is normalized with antiangiogenic or anti-fibrotic therapies [478, 500, 508–513].
Intravital optical microscopy [514] has also been useful for
calibrating clinical imaging modalities in patients [515].
With information about the structure and dynamics of the
angiogenic network, it is possible to analyze performance
and efficiency of the vasculature at a deeper level. These
analyses allow for further probing of the pathophysiology of
Angiogenesis (2018) 21:425–532
489
Fig. 21 MMTV tumor vasculature in the cranial window pillar TIC.
a MPLSM/SHG image of the indicated region in the bright-field
image (b). Near the edge of the PDMS, the vasculature extends radially into the central chamber. At this time point (day 7 after implantation), the vasculature is mature and has normal morphology in the
regions far from the tumor. Four feeding arterioles (red arrowheads)
and three venules (blue arrowheads) are indicated. These vessels have
significant flow and have acquired smooth muscle cells in their walls
(red, αSMA+-DsRed). Note that the arterioles generally have more
αSMA signal than the venules, as expected. c The vasculature near
the growing tumor has dramatically different morphology and flow,
as observed in other animal models and human tumors. The tumor
was not fluorescently labeled in this group, but is visible as the mass
extending from the central tumor (T) in b (from [493])
Fig. 22 Imaging of vasculature in the cranial window preparation.
Vascular sprouts entering a cranial window tissue isolation chamber. Time sequence of new vasculature (green) migrating toward the
top left into a cranial window TIC, past the edge of the PDMS disk
(dashed line) (imaged using MPLSM and SHG). Alignment of collagen fibers (white) is evident, and alpha-SMA+ cells can be seen
on the PDMS surface (red). The vasculature (green, FITC-dextran)
extends by forming perfused loops and sprouts. As the matrix remodels, the vessels also remodel as they advance. a: D1: day 1, D3: day 3,
D6: day 6 post-TIC implantation. A pillar structure, which defines the
height of the chamber, is indicated with * (from [493])
tumor angiogenesis and the implications of abnormal vasculature for drug and nutrient delivery [516–519].
Microfabrication technologies popularized in the past
two decades have allowed the design and implementation
of more sophisticated chamber designs. By creating customdesigned window chambers from polydimethylsiloxane, a
clear, biocompatible polymer, it is possible to control the
interactions between host and implanted tissues and to
improve imaging by partially constraining the growing tissues [493] (Fig. 21).
24.2 Advantages of intravital microscopy/chronic
window chambers for angiogenesis studies
Intravital microscopy in window chambers is characterized
by the following features: (1) It allows studies in an in vivo
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490
microenvironment, while ex vivo or in vitro studies require
more extensive validation; (2) it enables visualization and
measurement of the dynamics of the same vessels and cells
as the new vasculature forms and changes in response to
various treatments (Fig. 22); (3) it permits a variety of functional studies of angiogenic vessels which are being increasingly recognized in the angiogenesis field; (4) it allows dissection of the role of various mechanical (shear stresses,
hoop stresses, etc.) and biochemical signals known to govern
angiogenesis, (5) it allows investigation of the role of different organ microenvironments on vessel formation and function; and (6) a wide range or reagents are readily available
for the murine system (Abs, genetic manipulation, etc.).
24.3 Limitations and challenges of intravital
microscopy and window chambers
Intravital microscopy and window chambers experience some
limits: (1) They are labor-intensive, making screening studies expensive; (2) creating the window may cause transient
inflammation and inappropriate implantation of the window
can induce tissue damage; the use of compromised windows
affects angiogenesis in the windows (wound healing response);
(3) for single-photon microscopes, depth penetration is limited
to a few cell layers due to light scattering. Multiphoton microscopes permit deeper imaging but still limited to a few hundred
micrometers. OCT, Doppler OCT, and SWIR permit deeper
imaging, but are more expensive; (4) commercially available
advanced imaging techniques are very expensive and the most
advanced imaging techniques are not widely available.
24.4 Concluding remarks
Optical microscopy provides the high-resolution imaging
needed to distinguish cell dynamics, ECM components, and
intracellular features. By placing transparent windows over
various organs in animal models, we can take advantage of
optical microscopy to follow the formation of new vasculature
noninvasively for time periods of weeks or months. This enables studies of neovascularization in normal processes such as
wound healing and diseases such as cancer and allows detailed
analyses of responses to drugs that target blood vessels.
25 The RIP1‑Tag2 transgenic mouse model
The RIP1-Tag2 mouse model was one of the earliest oncogenic mouse models used in pioneering studies by Hanahan
and Folkman to identify and characterize the angiogenic
switch and multi-step progression to pancreatic neuroendocrine tumors. Due to its high vascularity, and synchronous
and rapid tumor development, it became a valuable preclinical tool for developing and evaluating response and relapse
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Angiogenesis (2018) 21:425–532
from anti-angiogenic therapy from its earliest stages to the
current day. Studies in the RIP1-Tag2 model predicted efficacy and helped to motivate clinical trials that led to the
approval of two compounds, everolimus and sunitinib, for
human pancreatic neuroendocrine tumors (PNET) in 2011.
25.1 The angiogenic switch
The RIP1-Tag2 mouse was developed by Douglas Hanahan
as a result of his interest in using newly developed transgenic
technology to use a well-characterized rat insulin promotor
to express a viral oncogene, SV40 T antigen, in the beta
cells of the pancreatic islets of Langerhans in order to study
oncogenic transformation of normal tissues [520]. The first
transgenic mice died at an early age from hypoglycemia,
consequent to the rapid proliferation of beta-islet cells,
which produced high insulin levels. Tumors from viable
mice were characteristically bright red and highly vascularized, and it was immediately proposed that this was likely
the result of what was later called the “angiogenic switch,”
which Judah Folkman had postulated, must occur for tumors
to grow beyond a small size [521]. But when and how is
this switch activated? Co-culture of the total islet population
with EC in a collagen matrix resulted in proliferation and
migration of EC toward only a small subset of “angiogenic”
islets to form capillary tubes, elegantly confirming that the
angiogenic switch occurred in hyperplastic islets prior to
tumor formation [522]. This also implies that a soluble factor [521], later identified as VEGF through multiple lines
of evidence, was critical. In normal adult mammals, the
vasculature is quiescent, and new blood vessels are formed
through angiogenesis, wherein new capillaries sprout from
existing vessels in a process largely driven through VEGF
signaling. Hanahan and Folkman proposed that this angiogenic switch is a discrete early step in tumorigenesis and
that the stages of progression that characterize RIP1-Tag2
tumorigenesis, where progressively smaller subsets of quiescent islets sequentially become hyperplastic, angiogenic, and
finally progress to invasive carcinomas (Fig. 23a), also exist
in human tumors [523]. In contrast to the popular hypothesis at that time that the angiogenic switch is induced by
upregulation of pro-angiogenic factors, RT-PCR analysis
of all VEGF ligands and their receptors, flt-1 and flt-2, as
well as acidic fibroblast growth factor (FGF1) indicated that
their expression levels were similar in islets before and after
the angiogenic switch [524, 525]. The fact that normal nontransgenic islets already contained high levels of various
pro-angiogenic factors suggested another mechanism in the
activation of angiogenesis. Indeed, the switch from vascular
quiescence to an angiogenic state involved the matrix metalloproteinase MMP-9, which was secreted by infiltrating
myeloid cells. MMP-9 rendered ECM-sequestered VEGF
bioavailable to its receptor, thus triggering an angiogenic
Angiogenesis (2018) 21:425–532
491
Fig. 23 RIP1-Tag2 mouse model. a Multi-step progression to tumors
in RIP-Tag2. Although oncogene expression begins during embryonic development (E8.5), the pancreatic islets initially have a normal
anatomical and histological appearance (“normal” stage). Beginning at 4–5 weeks of age, hyperplastic and dysplastic islets begin
to appear to comprise about 50% of islets by 10 weeks. Angiogenic
islets appear beginning around 6 weeks of age, and represent 10%
of all islets at 10.5 weeks. Angiogenic islets are recognized by their
dilated blood vessels and microhemorrhages. Tumors form beginning at 9–10 weeks and represent 2–4% of all the islets by 14 weeks.
About half of the tumors at end stage evidence either focal or widespread invasion to the surrounding acinar tissue. RIP-Tag2 mice die
at approximately 14 weeks of age primarily due to hyperinsulinemia.
b Anti-angiogenic Therapy Response and Relapse. Tumors treated
with anti-angiogenic therapy using an RTK inhibitor starting at
12–15 weeks “Response,” or 12–20 weeks “Relapse,” when vessels
have significantly rebounded. Tumors are stained with anti-insulin in
blue, vessels are stained with anti-CD31 in red, and surrounding exocrine pancreas is stained with amylase in green
switch; this identification was confirmed using co-culture
studies of angiogenic islets derived from MMP-9 KO RIP1Tag2 mice and EC in collagen matrices—in contrast to wildtype islets, the MMP-9 KO islets did not induce EC capillary
formation [526]. The result that VEGF-A is a pivotal factor
in the angiogenic switch was further confirmed with an islet
beta cell-specific knockout of VEGF-A in RIP1-Tag2 mice,
which impaired both angiogenic switching and subsequent
tumor growth [525].
25.2 Preclinical and clinical studies
These and many other concurrent studies identifying the
VEGF/VEGFR2 axis as a powerful therapeutic target led to
the development of numerous anti-angiogenic compounds.
Sarah Parangi and colleagues were the first who used the
RIP1-Tag2 model to perform preclinical studies with a
combination of three early compounds with anti-angiogenic activity—AGM-1470, the antibiotic minocycline,
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and interferon alpha/beta [527]. The combination can reduce
tumor growth, but not prevent it, and was an early preclinical success for anti-angiogenic therapy that helped to propel
this class of inhibitors into the clinic [527]. The rapid and
synchronous multi-step tumor progression that characterized the RIP1-Tag2 model made it then ideal for designing
three different trial formats (Fig. 23a): early treatment at
the hyperplastic stage to block the angiogenic switch commencing prior to tumor formation (prevention trial), treatment of mice bearing small tumors to determine whether
tumor growth and progression can be stopped (intervention
trial), and treatment of mice with substantial tumor burden and near death to test whether drugs can induce tumor
regression and promote survival benefits (regression trial).
These trials were first tested with AGM-1470, batimastat
(BB-94), Fc-endostatin, Fc-angiostatin, or a combination
of the latter two compounds and produced distinct efficacy
profiles in the various trial formats [528]. These studies suggested that those anti-angiogenic compounds needed to be
fine-tuned to target specific stages of disease progression
and that combinatorial strategies can broaden the effects
and enhance survival. Subsequent studies illustrating the
importance of targeted therapies that disrupt both tumor EC
and pericytes brought this notion to fruition in combination
therapy trials with two different Sugen inhibitors, SU5416
and SU6668, that target VEGFR2 and PDGFR, respectively [529]. While the VEGFR inhibitor SU5416 was most
potently effective against early-stage disease, SU6668 was
shown to block further growth of end-stage tumors suggesting that PDGFR+pericytes in tumors present a complimentary target to EC for efficacious anti-angiogenic therapy.
Combination therapies of these compounds were more
efficacious against all stages of islet carcinogenesis than
either single agent [529]. Congruently, subsequent studies
with sunitinib (SU11248), which targets both VEGFR2 and
PDGFR, showed remarkable efficacy in preclinical studies
in RIP1-Tag2 mice and produced increased survival relative
to monotherapy targeting VEGFR2 [530]. Collectively, these
studies were used to incentivize the use of sunitinib in clinical trials for PNET. These clinical trials produced an impressive extension in progression-free survival that leads to the
approval of sunitinib in 2011 [531], along with everolimus
that primarily targets mTOR signaling. Interestingly, mTOR
inhibitors also are very efficacious and produced increased
survival in the RIP1-Tag2 model [532]. Thus, the RIP1-Tag2
model predicted the beneficial effects of both sunitinib and
everolimus. However, significantly increased overall survival
of RIP1-Tag2 mice, but not PNET patients treated with these
inhibitors, can be reflective of the consequence of the different life spans of mouse versus human (approximately
15 weeks versus 70 years); therefore, a relatively short life
extension can be significant. Also, preclinical studies in the
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RIP1-Tag2 mice are confounded by the fact that tumors are
multifocal and that mice die from hypoglycemia, with relatively low (collective) tumor burdens and little metastasis
relative to that characteristic of other mouse tumor models or
human cancer. However, it is also noticed that evaluation of
overall survival in clinical studies is confounded by the fact
that placebo-treated PNET patients (unlike RIP1-Tag2 mice)
elect to “cross-over” to the treatment cohort, likely reducing
the calculated overall survival benefit in patients [533].
25.3 RIP1‑Tag2 versus PNET tumors
Another major complication in interpreting preclinical studies using the RIP1-Tag2 mouse lies in fundamental biological differences from PNET. Tumors in the RIP1-Tag2 model
are rapidly progressive and rarely form metastases, in contrast to the more indolent PNET tumors, although a subset
of PNET tumors is more aggressive and metastasized [534].
Further, the tumor gene driving RIP1-Tag2 oncogenesis is
a viral protein that interferes with p53 and retinoblastoma
tumor suppressor function, while the genes associated with
human PNET are predominantly chromatin-remodeling
genes, DNA repair genes, mTOR-PI3K pathway genes, and
menin mutations [534]. Notwithstanding these differences,
expression profiling of human PNET tumors split them into
three distinct groups, and remarkably two of them correspond to expression profiles identified in RIP1-Tag2 tumors
[535]. Interestingly, human PNET tumors rarely adapt to
culture, and while many cell lines from RIP1-Tag2 tumors
(βTC cell lines) have been generated, they are surprisingly
difficult to culture in comparison with other mouse tumor
cell lines in spite of their aggressive growth in vivo.
25.4 Adaptation to anti‑angiogenic therapy
The RIP1-Tag2 mouse represents a powerful model to
assess therapeutic response and relapse to anti-angiogenics due to their very synchronous tumor progression, and
this has made it a potent model to study adaptation to antiangiogenic therapy. The response phase is characterized by
tumor stasis and markedly reduced vascularity and vessel
normalization blocking hemorrhage formation that produces whitish tumors, while the angiogenic relapse phase
is characterized by tumor regrowth and revascularization
(Fig. 23b). McDonald and colleagues investigated the kinetics and mechanisms underlying vessel regrowth upon withdrawal of anti-angiogenic therapy and found that although
two different VEGFR inhibitors, AG-013736 or AG-028262,
significantly regressed tumor vasculature, they left behind
empty sleeves of pericyte-covered basement membrane
which functioned as a scaffold for rapid revascularization
after treatment withdrawal. They concluded that targeting
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pericytes and the sleeves of basement membrane appeared
to be an important strategy to produce an enduring response
to anti-VEGF therapy [536]. Casanovas et al. [537] performed preclinical studies in the RIP1-Tag2 model, which
demonstrated that after a period of response to continuous
anti-VEGFR2 monotherapy, tumors rebounded although
blockade of VEGFR2 signaling persisted. They found that
the mechanism for regrowth was induction of alternative
pro-angiogenic factors, including the fibroblast growth factor
family (FGF) and ephrins. In addition, the response phase
is characterized by the angiostatic and immunostimulatory
polarization of myeloid cells that contributes to pruning and
normalization of the vasculature [538]. Tumors respond to
these changes by activating and repolarizing myeloid cells
to an angiogenic and immunosuppressive phenotype and by
initiating an adaptive immune response by upregulating the
negative immune checkpoint regulator, programmed cell
death ligand PDL1; both immune adaptations limited the
efficacy of VEGF/VEGFR inhibitors [538–540]. Another
form of evasion from anti-angiogenic therapy that was
characterized in the RIP1-Tag2 model (and glioblastoma)
is that of increased invasion and metastasis following pharmacological and genetic targeting of the VEGF signaling
axis [530], the consequence of increased hypoxia induced
by vascular fallout. While increased invasion has not yet
been found in PNET treated with anti-angiogenic therapy, it
appears bevacizumab treatment can cause increased invasion
in a subset of patients with glioblastoma multiforme [541],
again highlighting the predictive power of this model.
25.5 Limitations and challenges
In spite of the fact that preclinical studies in the RIP1-Tag2
model predicted the efficacy of two approved therapies, the
model is multifocal, rapidly progressive, and driven by a
viral oncogene unlike its human counterpart. Because some
PNET patients progress on available targeted therapies, it
is desirable to further develop biomarkers of therapeutic
response together with Patient Derived Xenograft (PDX)
models from resistant tumors in humanized mouse models in order to identify beneficial secondary or concurrent
therapies [533].
25.6 Concluding remarks
The RIP1-Tag2 model of beta islet-cell carcinogenesis has
been instrumental in studying mechanistic underpinnings
of tumor angiogenesis and in revealing adaptations to the
environmental stresses elicited by anti-angiogenic therapies,
leading to valuable insights and predictions regarding clinical successes and failure.
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26 The MMTV‑PyMT breast cancer model
The MMTV-PyMT mouse breast cancer model was
described in 1992 by Guy, Cardiff, and Muller [542] and has
been broadly employed for studying several aspects of breast
cancer progression, such as the regulation of angiogenesis,
cancer cell intravasation and dissemination, and metastatic
colonization. In this transgenic model, the polyomavirus
middle T antigen (PyMT) is expressed in the mammary
epithelium from the long terminal repeat (LTR) promoter
of the mouse mammary tumor virus (MMTV). The transforming activity of the PyMT is dependent on its association
with several oncogenic proteins, resulting in the multifocal
growth of mammary adenocarcinomas that metastasize to
the lung [542].
26.1 Tumor angiogenesis in MMTV‑PyMT mice
Lin et al. have examined breast cancer progression in
MMTV-PyMT mice [543–545]. Early malignant transition
occurs between 8 and 12 weeks of age and is associated
with increased nuclear pleomorphism, peri-acinar infiltration by leukocytes, and angiogenesis [544]. The angiogenic
switch—the conversion of a quiescent, non-angiogenic vasculature into one that is actively growing and infiltrative—
generally demarcates the malignant progression of benign
lesions [5]. Although the transition from hyperplasia to the
adenoma/mammary intraepithelial neoplasia (MIN) involves
the enlargement of the tumor mass in MMTV-PyMT mice,
the density of functional blood vessels remains constant in
these premalignant stages and is comparable to that of normal mammary glands [545].
A change in blood vessel distribution and density occurs
during the transition from premalignant to malignant stages.
In early adenocarcinomas, the acinar structures are replaced
in the core of the lesion by small solid nodules displaying
increased nuclear pleomorphism and cytologic atypia [545].
A high-density vascular network becomes evident in the
solid nodular area, whereas surrounding premalignant acini
are not associated with angiogenesis. Therefore, the development of adenocarcinomas, but not premalignant adenoma/
MIN, is accompanied by angiogenesis in MMTV-PyMT
mice. Of note, both high- and low-density vascular networks
are found in early or late adenocarcinomas [545]. Variation
in the features and density of blood vessels in transgenic
(autochthonous) tumors is likely controlled by a multitude of
parameters, including the spatiotemporally regulated expression of pro- and anti-angiogenic factors and inflammatory
mediators, as well as biophysical forces, which dynamically
control vessel functionality, growth, and regression in distinct tumor microenvironments [5].
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26.2 Role of macrophages in MMTV‑PyMT tumor
angiogenesis
Monocytes and macrophages have been implicated in
MMTV-PyMT tumor angiogenesis and progression [543,
545]. MMTV-PyMT mice carrying a homozygous mutation in the colony-stimulating factor-1 (Csf1) gene (Csf1op/op)
recruit fewer tumor-associated macrophages (TAMs) than
Csf1-proficient mice [543]. The macrophage-deficient
phenotype of MMTV-PyMT/Csf1op/op mice is associated
with delayed tumor progression (decreased incidence of
invasive adenocarcinomas and pulmonary metastasis) and
defective tumor angiogenesis [543, 545]. Conversely, the
genetic overexpression of Csf1 in the mammary epithelium
of MMTV-PyMT mice results in the premature accumulation of macrophages around hyperplastic lesions and adenomas and accelerates both the development of an angiogenic
vasculature and tumor progression [545]. These findings are
consistent with results obtained using a macrophage-depleting CSF1 receptor (CSF1R) inhibitor [546] and agree with
studies in other mouse tumor models, such as RIP1-Tag2
transgenic mice [547].
Conditional overexpression of vascular endothelial
growth factor-A (Vegfa) in the mammary epithelium of
4–6-week-old MMTV-PyMT mice was sufficient to induce
the formation of enlarged, angiogenic blood vessels around
premalignant acini [548]. Also, VEGF-A overexpression rescued macrophage infiltration, stimulated the formation of a
high-density vascular network, and restored tumor progression in MMTV-PyMT/Csf1op/op mice [548]. These findings
suggested that macrophages can promote tumor angiogenesis in MMTV-PyMT mice, at least partly, through VEGFA [548]. In another study, conditional deletion of Vegfa
specifically in myeloid-lineage cells, which encompass
macrophages, altered vascular patterning in MMTV-PyMT
mice [549]. This phenotypic conversion involved decreased
vessel tortuosity and density and increased pericyte coverage, which are consistent with an attenuated angiogenic
response. While overall VEGF-A levels were unabated by
loss of myeloid-derived VEGF-A (likely due to substantial
VEGF-A production by cancer cells), the phosphorylation
of endothelial VEGF receptor 2 (VEGFR2) was decreased
in the tumors [549]. These findings suggest that myeloid
cells, or subsets thereof, provide a non-redundant source of
bioactive and pro-angiogenic VEGF-A in the perivascular
microenvironment of MMTV-PyMT tumors. Macrophages
enwrap angiogenic blood vessels in MMTV-PyMT tumors
[550, 551], which may explain the importance of macrophage-derived VEGF-A for tumor angiogenesis [549].
However, it should be noted that attenuation of angiogenesis
does not necessarily translate into tumor inhibition. Indeed,
Stockmann et al. observed accelerated tumor progression
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in MMTV-PyMT mice with myeloid-specific Vegfa deletion [549]. This finding can be attributed to the “normalized” structure of the blood vessels, which may have led to
improved tumor oxygenation and cancer cell growth [552].
Besides VEGF-A, TAMs secrete additional pro-angiogenic factors [5]. For example, the genetic deletion of Wnt7b
in TAMs reduced the expression of WNT/β-catenin mitogenic target genes in vascular endothelial cells and decreased
the MVD in MMTV-PyMT adenocarcinomas [553].
MMTV-PyMT mice have also been instrumental for
studying macrophage–vascular interactions in the mammary
tumors. The genetic inactivation of Tie2/Tek in TAMs did
not block their recruitment to tumors, but impaired their
ability to associate with immature blood vessels and sustain tumor angiogenesis [551]. Further to supporting angiogenesis, perivascular TAMs enhance vascular permeability
through VEGF-A, which facilitates cancer cell intravasation
and metastasis in MMTV-PyMT mice [554]. The association
of macrophages with tumor blood vessels is increasingly recognized as an important determinant for early breast cancer
metastasis in both MMTV-PyMT mice and patients with
breast cancer [554–556].
26.3 Anti‑angiogenic therapy in MMTV‑PyMT mice
The pharmacological inhibition of VEGF-A signaling attenuates tumor angiogenesis in both transgenic MMTV-PyMT
mice and mice carrying orthotopic MMTV-PyMT tumor
transplants [557, 558]. More marked anti-angiogenic effects
are observed when VEGF-A and angiopoietin-2 (ANGPT2)
are neutralized concomitantly in the mammary tumors [557,
558]. Dual VEGF-A and ANGPT2 blockade also leads to
substantial tumor inhibition and significantly extends the
survival of transgenic MMTV-PyMT mice [558]. Further
to suppressing tumor angiogenesis and abating the density
of the vascular network, combined VEGF-A and ANGPT2
blockade normalized the structure of the blood vessels
that survived in viable tumor regions and facilitated the
extravasation and perivascular accumulation of activated,
interferon-γ (IFNγ)-expressing CD8+ T cells [558]. These
tumor phenotypes were likely due to the complementary
actions of ANGPT2 and VEGF-A inhibitors on the tumor
microenvironment: Whereas VEGF-A blockade has more
marked vascular-pruning activity, at least acutely [559],
ANGPT2 blockade limits angiogenesis [551] and enforces
the maturation and stabilization of the remaining vessels,
making them permissive to T cell extravasation and trafficking [560, 561]. Thus, MMTV-PyMT mice provide a useful
model for studying the kinetics of angiogenesis inhibition
and vascular normalization in response to anti-angiogenic
drugs.
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26.4 Limitations and challenges
Although angiogenesis has been extensively studied in
primary adenocarcinomas, little is known of the mechanisms controlling vascularization of pulmonary metastases
in MMTV-PyMT mice. Because macroscopic pulmonary
metastases become evident only in aging mice (12–16 weeks
of age), the development of multifocal primary tumors may
impose significant constraints on the investigator’s ability to examine metastatic lesions whose mass would be
large enough to stimulate angiogenesis. In this regard, it is
increasingly appreciated that metastatic tumors may rely,
at least partly, on vessel co-option and be less dependent
on angiogenesis than primary tumors [562]. Further studies
are required to establish whether pulmonary metastases that
develop in MMTV-PyMT mice vascularize through angiogenesis, vessel co-option, or both.
27 Tumor implantation models
27.1 Implantation of angiogenic tumors
Tumors may be implanted in mice as xenografts or in a
syngeneic context. Implantation may also be done subcutaneously or orthotopically at the correct organ site where
the tumor usually grows. There are many angiogenic tumor
models to cite only a few such as glioma (U87 human glioblastoma, mouse GL261 glioma), breast (MDA-MB-231,
66cL4 mouse mammary tumor), lung (Lewis lung carcinoma, human LNM35 lung) melanoma (B16 mouse
melanoma), prostate (human prostate CWR22Rv1 tumor)
[563]. Tumor cells may be injected as single-cell suspension, embedded in matrix such as matrigel or implanted
as spheroids or organoids. Cells may be transduced with a
suitable reporter construct (such as luciferase), and tumor
development may be followed by imaging. Usually at a given
time point, animals are killed and tumors are processed for
immunohistochemistry, RNA expression analysis, or protein
detection (immunoblot). The angiogenic response is analyzed by immunohistochemistry or immunofluorescence
using appropriate antibodies for endothelial cells (CD31,
CD34) or pericytes (alpha2SM actin, desmin). This analysis
will give information about vessel density, vessel size distribution, and vessel quality (tight or loose pericyte coverage).
The angiogenic response is usually high in the viable core
of the tumor. At the invading front, tumor cells may use
preexisting vessels for invasion.
It is of note that tumors may exhibit different angiogenic
responses when they are either implanted in orthotopically
or subcutaneously as it has been shown in the case of astrocytomas [564].
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27.2 Detection of “non‑angiogenic” tumors, which
rely instead on “vessel co‑option”
While there is an overwhelming body of preclinical/experimental and clinical evidence accumulated over 40 years
showing the importance of sprouting neo-angiogenesis in
tumor growth, progression, and metastasis, it is increasingly appreciated that there are many circumstances in
which tumors can grow and expand without inducing any
new blood vessel capillaries. This realization has major
implications for understanding some of the clinical efficacy
limitations including failures of anti-angiogenic drugs. Thus,
new blood vessels can be formed by a process of the “splitting” of existing vessels, a process known as intussusception,
as discussed above. Or, certain tumors, especially neural
crest-derived cancers, may form blood forming channels by
“vasculogenic mimicry” whereby tumor cells line the vessel
lumen, rather than authentic host EC. However, the most
dominant manifestation of non-angiogenic tumors is likely
“vessel co-option” and it is particularly prominent in vascular-rich organs that are the most common sites of metastatic
disease, namely lung, liver, lymph nodes, and brain [117].
Broadly speaking, vessel co-option refers to the ability
of tumors to parasitize preexisting blood vessels of the tissue, bypassing the requirement of developing new ones.
The modern era of vessel co-option may have begun in
1996–1997 with the publication of two pathology reports by
Pezzella and colleagues. Therein, a pattern of tumor growth
of non-small cell lung cancer and lung metastases was recognized which made use of alveolar air spaces for expansion
and alveolar vessels for blood supply without any indication
of angiogenesis [121, 565]. Histopathological descriptions
of similar growth pattern in the lungs and analogous patterns
in the brain and liver date to the late 1800s and early 1900s.
The distinction between so-called newly formed versus preexisting blood vessels was noted in the past, but the biological significance of each form was generally not appreciated.
This has changed in the era of anti-angiogenic therapy—for
example, targeting the vascular endothelial growth factor
(VEGF) pathway to impair and cause regression of new
vessel sprouts does not deplete preexisting vessels. Tumor
exploitation of vessel co-option has recently been shown to
account for instances of intrinsic and acquired resistance
to anti-angiogenic drugs, both preclinically [122, 566, 567]
and clinically [119]. Moreover, growth patterns associated
with vessel co-option are emerging as important for patient
prognosis [120, 568].
There is currently no “assay” as such to quantitate the
extent of vessel co-option in tumors. Conventional measurement of tumor microvessel densities (MVD) within tumors
has high potential for error when one considers that normal tissue vessel densities—as well as tumors where cancer
cells surround these vessels—may be high [569]. Rather,
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histopathological analysis is currently used with certain
key features being looked for. Some examples are briefly
described below; more detailed descriptions can be found
elsewhere [117, 569].
27.3 Some “hallmarks” of detecting vessel
co‑option in tumors growing within the lungs,
liver, and brain
Though the histological structures of these organs are distinct, tumors within these sites that co-opt preexisting blood
vessels—either as part of its overall mass or throughout—
often have some of the following features:
Fig. 24 Visualization of vasculature in transplanted mouse models.
Immunofluorescence and immunohistochemical staining of formalin-fixed mouse lung samples to enable differentiation between vessel co-option and angiogenesis in tumors. a, b Vessel co-opting
tumors are observed in spontaneously formed lung metastases from
mice with orthotopically implanted then surgically resected MDAMB-231/LM2-4 breast tumors. In a sections are stained for alveolar
cell marker podoplanin, EC marker CD34 and nuclei marker DAPI.
Tumor cells can be seen filling alveolar spaces along the border and
incorporating alveolar capillaries into the tumor core. In b is the corresponding section stained for HLA human cell marker and hematoxylin to show the presence of tumor cells with respect to host
stroma and lung parenchyma. A bronchiole is also seen to be taken
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1. The architecture of the normal host tissue is preserved
within the tumor. This is in contrast to angiogenic
tumors, which tend to destroy the surrounding tissue.
Tissue architecture preservation has been aided by
immunohistochemistry for basement membrane and
epithelial markers showing continuity of form across
the tissue–tumor interface.
2. Lack of surrounding capsule or desmoplastic tumor
boundary. Angiogenic tumors often form defined masses
or nodules, a symptom of the inflammatory response of
the host to the tumor, but this is lacking in “purely” vessel co-opting tumors.
3. Invasive, infiltrative, and replacing tumor growth.
Minimal compressive tumor growth and compression
of normal tissue structures. Rather, tumor cells tend to
into the tumor and gradually filled with tumor cells. The tumor border is irregular. c, d Angiogenic growth is observed in spontaneously
formed lung metastases from mice bearing intra-renal implanted
RENCA tumor cells that later underwent nephrectomy. Sections are
stained for alveolar and bronchial epithelium cell marker cytokeratin
7, EC marker CD34 and nuclei marker DAPI. RENCA tumors grow
in “cannonball” shape, compressing lung tissue and excluding them.
The lung–tumor interface of another nodule is shown at high magnification in d. Lung tissue is compressed or “pushed” aside to allow
tumor expansion. The tumor border is smooth, and microvessels are
not associated with alveolar epithelium within the tumor. Scale bar
represents 200 µm. Regions in dashed boxes are expanded on the
right. “T” = tumor. Arrow = columnar bronchial epithelium
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infiltrate along preexisting vessels, often in direct contact with epithelial–endothelial basement membranes.
Epithelial cells may become destroyed or stripped from
their vessels during this process. Tumor borders are
irregular.
4. Incorporation of characteristic cells and structures of
the tissue into the tumor. Parenchymal epithelial cells
(e.g., lung pneumocytes, liver hepatocytes, neuralglia)
and larger intact structures (e.g., lung bronchioles and
large vessels in the lungs, portal triads in the liver) are
taken up into the expanding infiltrative tumor mass and
get locked into the tumor.
5. Maintained vessel morphology and marker expression.
Co-opted vessels are known to be remodeled by surrounding tumor tissue and therefore may not in all cases
appear “normal.” Newly recruited co-opted vessels in
particular may maintain morphological and molecular
characteristics of tissue endothelium (e.g., GLUT-1 and
p-glycoprotein endothelial expression in brain tumors).
6. Low rates of endothelial cell proliferation relative to
angiogenic tumors. Specific molecular markers of coopted blood vessels are not currently known; thus, several criteria are used to distinguish between vessel subtypes and differential growth patterns of tumors. Vessel
co-option was first identified in tumors from patients;
however, experimental mouse models have also been
shown to recapitulate the growth patterns observed in
human tumors.
The major question now for future studies is whether
“anti-vascular” therapeutic strategies can be developed that
target co-opted vessels, in addition or instead of angiogenic
vessels, without impacting the normal vasculature; some
possibilities have recently emerged, for example using antibody–drug conjugates [570].
Shown in Fig. 24 are preclinical examples of both vessel co-opting and angiogenic tumors. Lung metastases from
mice with breast cancer cell (MDA-MB-231/LM2-4) vs.
kidney cancer (RENCA). The former develops purely nonangiogenic vessel co-opting lung metastases, whereas the
latter forms both angiogenic (shown here) and vessel coopting lung metastases. Staining for lung-specific epithelial
and EC markers shows the architecture of blood vessels in
lung tissue and metastases. In vessel co-opting lung metastases, alveolar cells surrounding capillaries are taken up by the
expanding mass at the tumor border, but alveolar epithelial
cells are gradually shed from vessels toward the tumor center
while capillaries remain.
27.4 Limitations and challenges
Implantation models are generally very useful in deciphering
mechanisms of the tumor angiogenesis and the role of given
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angiogenesis stimulator or inhibitor, receptors or modulating
intracellular factors. The only drawback is that the adaptive
immune system in xenotransplantations is absent and thus
the data must be interpreted with care. The other challenge
is the implantation site. Usually, it is better to orthotopically
implant tumor cells since the correct host tissue is present
in this case. Another challenge is whether or not to implant
cell suspensions or spheroids. Spheroids seem in many cases
better suited because they already adopt a 3D tissue configuration which preserves some of their genetic program that is
encountered in a 3D tissue.
Co-optive growth may be induced by anti-angiogenic
therapy as it is encountered in glioblastoma for instance.
Thus, a switch from angiogenesis to co-option may occur in
this case. It would be important to develop therapies against
co-opted vessels but this may be damaging because there
are for the moment no specific markers discovered for coopted vessels.
27.5 Concluding remarks
Tumor implant models are of great value to decipher tumor
vessel interactions. Despite several limitations, this is still
important methodology that is useful in characterizing many
molecular mechanisms of tumor angiogenesis.
28 Mouse hind limb ischemia model
Peripheral arterial disease (PAD) is caused by atherosclerosis and occlusion of peripheral blood vessels and results in
intermittent claudication or in more severe stage in critical
limb ischemia. The prevalence of PAD increases with age,
affecting 6% of individuals aged 50–60 years, and 10–20%
of individuals aged > 70 years [571, 572]. In the upcoming years, the number of PAD patients will increase even
more due to the aging of the population and the increase in
number of patients with obesity and type 2 diabetes. Unfortunately, current therapies for improving the perfusion of
the lower limbs in patients with PAD are hardly effective
[573, 574]. Therefore, much effort has been put in the last
decade in defining new strategies to improve the blood flow
to the lower extremities and to promote blood vessel growth.
These strategies that are based on the concept of therapeutic
neovascularization are often also called therapeutic angiogenesis or more general, neovascularization [573].
Neovascularization comes in three anatomical and physiological forms; angiogenesis, arteriogenesis, and vasculogenesis [574]. Angiogenesis describes an expansion of the
microvasculature, because of sprouting of EC from preexisting capillaries, followed by their proliferation, migration,
and capillary formation and is mainly hypoxia driven [246].
By contrast, arteriogenesis describes the remodeling of
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Fig. 25 Laser Doppler perfusion imaging and angiography
of blood flow. Analysis of blood
flow recovery in time by laser
Doppler perfusion imaging (left
panels) and angiography (right
panels)
existing arterioles into collateral arteries so that they can
deliver more blood flow to the limb [575]. Finally, adult vasculogenesis describes the incorporation of circulating (progenitor) cells into the regenerating microvasculature [573].
In PAD, neovascularization primarily occurs in the form of
13
arteriogenesis or collateral formation, although angiogenesis
in more distally located ischemic areas does occur.
To test new therapeutic approaches to induce neovascularization, as well as unraveling the complex cellular and
molecular mechanisms involved in the regulation of the neovascularization process, mouse models mimicking critical
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limb ischemia have been developed. Although these models
may vary in crucial aspects, commonly they are referred to
as the mouse hind limb ischemia model. Being in vivo models they are complex and not always straightforward to interpret, but they faithfully reproduce the physiology of blood
recovery in PAD after stenosis of the original conduit. In this
section, we will discuss the critical aspects of the variations
in the use of the mouse hind limb ischemia model, their
consequences for interpretation of the data obtained, and the
limitations and pitfalls in their use.
499
however, severe ischemia in the calf muscles in the excision
model is warranted.
A good comparison of the various models has also been
hampered by the lack of a proper description of the anatomy
of the blood vessels in the mouse limb. Hopefully the recent
excellent description of the arterial anatomy of the murine
hind limb by Kochi et al. [593] will contribute to a better understanding of the outcomes of the different ischemia
models.
28.1 Mouse model for hind limb ischemia
28.2 Variants of the surgical procedure to induce
mouse hind limb ischemia
Couffinal et al. were among the first to describe a mouse
model for hind limb ischemia [576]. They induced acute
hind limb ischemia by ligating the proximal end of the femoral artery, and the distal portion of the saphenous artery,
followed by excision of ligated part of the femoral artery and
attached side branches. The recovery of blood flow in the
ischemic limb due to angiogenesis and arteriogenesis was
monitored by laser Doppler perfusion imaging (LDPI) [576],
Fig. 25. Subsequently, many groups used similar approaches
and variants of this model [577, 578]. Surgical procedures
range from a single ligation of the femoral or iliac artery
[579–581] to a complete excision of the artery [576] and
sometimes even the vein and the nerve were dissected as
well [582, 583]. Several excellent review papers have been
written on these variations in the hind limb ischemia model
[584–587]. When choosing a particular variant of the mouse
hind limb ischemia model for a study, it is important to
define the goal of the study. For instance, when testing new
pro-arteriogenic approaches, there should be an appropriate
therapeutic window in which an improvement of blood flow
recovery can be monitored. In mice that rapidly form new
collaterals, for example, C57BL/6 mice with a single ligation
of the femoral artery, it is difficult to monitor an increase
toward an even faster collateral formation [588], and therefore a model with more severe injury is required [589, 590].
On the other hand, such a fast model is ideal for mechanistic
studies in which the effects of the deficiency or inhibition
of crucial factors on blood flow recovery are studied [591,
592]. The fast recovery model is also associated with less
animal discomfort.
Another crucial question is which aspect of neovascularization will be studied. Arteriogenesis is the remodeling
of preexisting arterioles to larger diameter collateral blood
vessels required for restoring the blood flow to the distal
ischemic parts of the limb, whereas angiogenesis is the
sprouting of new (capillary) vessels into the ischemic tissue.
One can imagine that the excision model to induce hind limb
ischemia results in the disruption of the preexisting arterioles in the adductor muscle, and thus the presumed substrate
for collateral formation. In pro-angiogenic intervention,
General aspects of the surgical procedures Before surgery,
mice are anaesthetized with an intraperitoneal injection of a
combination of midazolam, medetomidine, and fentanyl. In
all models, the femoral vein and nerve are preserved. After
surgery, the skin is closed with 6/0 Ethilon sutures (procedures adapted from [585]).
Single electrocoagulation of femoral artery Directly after
incision of the skin in the left inguinal region, the subcutaneous fat pad in the thigh becomes visible and is pulled
aside distally. After dissection of the artery from the nerve
and vein, ischemia is induced by electrocoagulation of the
left femoral artery, proximal to the superficial epigastric
artery. Electrocoagulation results in complete occlusion of
the artery. After electrocoagulation, the proximal end of the
artery moves proximally into the surrounding tissue and the
distal end moves distally, so there will be a distance of a few
millimeters between both ends after the surgical procedure.
Single electrocoagulation of iliac artery A larger skin
incision in the inguinal region is made for this procedure.
Furthermore, there is no need to cleave the fat pad. For exposure of the iliac artery, a retroperitoneal approach is used. By
carefully moving the peritoneum proximally with a cotton
swab, a good exposure of the iliac artery is possible. In addition, preparation of the artery from the vein is necessary. The
internal iliac artery serves as a landmark; direct proximally
of the internal iliac artery an electrocoagulation of the common iliac artery is performed.
Double electrocoagulation of both femoral artery and
iliac artery For a double coagulation model, both common iliac artery and femoral artery are electrocoagulated.
First, an electrocoagulation of the common iliac artery is
performed followed by electrocoagulation of the femoral
artery. These coagulations are at the same anatomical levels
as in the single electrocoagulation procedures of the femoral
artery and the iliac artery.
Total excision of femoral artery The upper thigh is
exposed through an inguinal to knee incision. A proximal
part of the femoral artery is removed after double ligation.
Then, all side branches are isolated and coagulated. The distal ligation site is at the popliteal artery level distal from the
13
500
saphenous artery bifurcation. The whole artery in between
is removed.
28.3 Analysis of blood flow recovery
and neovascularization
Traditionally, blood flow recovery is analyzed by Laser
Doppler Perfusion Imaging (LDPI) where the flow in the
footpads of the ligated and contralateral limb is measured
and expressed as the ratio between the ligated and unligated
footpad. The restoration of the blood flow in time is used as
a readout for neovascularization (angiogenesis and arteriogenesis combined) in the ischemic limb.
Next to the LDPI-based flow analysis, immunohistochemical analyses of the neovascularization are used. Usually, the
angiogenic response is studied in the distal part of the limb,
the ischemic calf muscle (gastrocnemius), using EC staining
(CD31, von Willebrand is in absolute numbers per area or as
number of blood vessels related to number of myocytes). In
addition, the arteriogenic response, (i.e., newly formed collateral arteries) is commonly studied in the adductor muscle.
They are usually stained with alpha smooth muscle actin to
demonstrate the arterial nature, whereas size and location
are indicative of their collateral nature.
In addition to these two commonly used methods to monitor the neovascularization in the ischemic limb of the mouse,
sometime a functional test is used in which the use of the
foot and limb is analyzed for instance by assessment of the
plantar/dorsiflexion [594]. In the more severe models, necrosis of toes or even feet ensues, leading to auto-amputation.
Semiquantitative scores have been developed and used to
assess efficacy of interventions. Moreover, the increased sensitivity of current imaging methods may represent a major
step forward. For example, the use of micro-CT analysis
(which previously was typically used as a postmortem procedure due to the low sensitivity and lack of good contrast
agents [585], has now become a method for in vivo imaging of the microvasculature [595], enabling the analysis of
the neovascularization over time. Recently, it was demonstrated by Hendrikx et al. [596] that single photon emission
computed tomography perfusion can be used to analyze
neovascularization processes in the mouse hind limb, even
with such a resolution that they can demonstrate that LDPI
analysis underestimates the revascularization processes in
this model.
28.4 Limitations and challenges
One of the major limitations of the currently used hind limb
ischemia model is the acute nature of the ischemia, whereas
in the patients with peripheral artery disease the ischemia
develops gradually. Although an interesting approach
for gradual induction of ischemia in mice using ameroid
13
Angiogenesis (2018) 21:425–532
constrictors has been described by Yang et al. [597], this
model has not received large follow-up.
The capacity for vascular regeneration and neovascularization differs strongly between commonly used mouse
strains [598, 599]. For many years, it has been known that
C57BL/6 mice have a stronger neovascularization capacity
than Balb/c mice [581, 600] and only recently, it has been
discovered that this difference in regenerative response is
linked not only to a difference in the preexisting collateral
bed in the various strains [598] but that it also can be attributed to a specific gene locus in chromosome 7 of the mouse
[601–604]. Because of their slow regenerative response,
Balb/c mice are frequently used for hind limb ischemia studies, on the assumption that this slow response better mimics the situation in patients with PAD that display a poor
regenerative capacity. A recent study by Nossent et al. [605]
compared the mRNA expression pattern of angiogenesis and
arteriogenesis related genes in the hind limb of C57BL/6
and Balb/c mice. This showed that in Balb/c mice a stronger
upregulation of pro-angiogenic and pro-arteriogenic genes
can be observed when compared to C57BL/6 mice, despite
the poorer regenerative phenotype in Balb/c mice. These
findings suggest that rather than a more accurate model
of human critical limb ischemia, Balb/c mice lack a thus
far unknown factor that is crucial for vascular regenerative
response.
Since inflammation and the immune system play an
important role in the regulation of the angiogenic and arteriogenic responses in the hind limb ischemia model and
several immune cells including monocytes, T cells, and NK
cells are crucial in neovascularization [579, 581, 591, 592,
606–609], the use of immune-compromised mouse strains
for cell therapy approaches with human cells is not ideal
and the results of these studies should be interpreted with
caution. This becomes even clearer when the fate of injected
cells, and their rapid disappearance even from syngeneic
mouse is taken into account [609]. A very comprehensive
review on the use of various immunocompromised mouse
strains in hind limb ischemia models for human cell therapy
validations has recently been published by Thomas et al.
[594] and provides valuable information on the choice of
the proper model.
Last but not least, a major limitation for most of the hind
limb ischemia mouse studies is that they are performed in
healthy young mice, whereas the PAD patients usually are
older, and have atherosclerosis, type-2 diabetes, or other
comorbidities. These comorbidities strongly affect the neovascularization responses in mice [610, 611]. Moreover,
induction of vascular remodeling may also affect the underlying atherosclerotic disease leading to arterial occlusion.
This phenomenon is described as the Janus phenomenon of
neovascularization by Epstein et al. [612].
Angiogenesis (2018) 21:425–532
28.5 Concluding remarks
Many variants of the mouse hind limb ischemia model are
used; researchers should be aware of the different options,
select the model that fits best the goal of their experiments,
and properly describe the type of model used. It should also
be noted that although this model is commonly referred to
as an angiogenesis model, the mode of neovascularization is
a true combination of collateral formation and angiogenesis.
29 Large animal models for myocardial
angiogenesis
There is little doubt that large animal models are needed for
translational and preclinical studies of myocardial angiogenesis. Advantages of large animal models over smaller
animals are that the heart better resembles human heart in
size, anatomy, and function. Also, the same imaging equipment and treatment strategies, including similar dosage, can
be used. Due to ethical reasons and a good resemblance to
human, sheep and pigs are nowadays most widely used large
animals in myocardial angiogenesis studies. Recent developments in these models are briefly described in this chapter.
29.1 Acute ischemia models
Acute myocardial ischemia models have been widely used, as
they are rather easy to replicate in large number of animals.
It will also be easy to make interventions if any arrhythmias
occur. The models used are either surgical or catheter-based,
very consistent, and variation between the animals is usually
low. In surgical models, manipulation of the coronary arteries
from a thoracotomy opening to induce ischemia is a widely
used method [613]. However, from the thoracotomy opening, surgery affects the potential therapeutic approaches as
pericardial attachments complicate further interventions and
make it difficult to evaluate possible effusate accumulation,
which often complicates pro-angiogenic therapies.
On the contrary to surgical models, catheter-based endovascular models leave much of the heart intact and post-operational complications are much less frequent. Widely used
catheter-based ischemia models include ischemia–reperfusion model [614], infarction due to an intracoronary injected
solution [615] and intracoronary coil occlusion [616].
29.2 Chronic ischemia models
A healthy animal has several mechanisms to respond to
acute ischemia, which chronically ischemic patients may
not have anymore. This has led to the development of more
slowly developing occlusion and ischemia models to counteract and expand these adaptation mechanisms and to better
501
mimic the lack of necrosis, acute inflammation and strong
endogenous angiogenic stimuli, which takes place in acute
ischemia models [617].
Very widely used gradual obstruction model is a surgical ameroid constrictor model [618]. However, as a surgical
model, this has the same disadvantages as described above.
Endovascular models have also been recently developed.
Catheter-based models include the use of copper stents
[619], copper-plated stents [620], or oversized steel stents
[621], where in-stent restenosis causes stenosis in the coronary artery. However, in these stent models, the stenosis
size is variable, and its development cannot be controlled.
There is also a ligated hourglass-shaped stent-graft model
[622], but in this model, the exact time of total occlusion is
not known. A recent bottleneck stent model can be varied
for either reversible myocardial ischemia or ischemic cardiomyopathy [623], depending on if the stent is allowed to
occlude in this model. Occlusion is controlled by gradual
reduction of antiplatelet and antithrombotic therapy.
29.3 Non‑ischemic heart failure models
In addition to ischemic heart failure models, non-ischemic
models of heart failure have been developed that might be
used to study neovascularization as a therapeutic approach
for the improvement of cardiac function. In a recent model,
heart failure is induced by chronic pulmonary hypertension
[624]. To induce left ventricular hypertrophy and heart failure, different variations of aortic banding have been used
[625]. Pacing models to induce heart failure have also been
extensively used [626].
29.4 Myocardial stainings
Immunohistochemical stainings are used to histologically
detect the effects of angiogenesis. Hematoxylin–eosin staining
is used for tissue morphology characterization. Endothelial
cells can be stained with PECAM-1, which is also known as
CD31. Lymphatic endothelium can be stained with LYVE-1
or PROX-1. These pericytes, the smooth muscle cells surrounding blood vasculature, are stainable with α-SMA.
29.5 Challenges and limitations
Large animal models are the last translational step from
bench to the first clinical trials. Therefore, using these
models requires careful planning. Efficient delivery of the
therapeutic agent, be it cellular, viral, or growth factor-based
therapy, is a very critical factor for successful treatments.
Earlier, mainly intramyocardial injections were used as
these can provide robust and efficacious transduction, but
more recent studies have used intracoronary infusion and
retrograde coronary sinus infusion to achieve more global
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502
Angiogenesis (2018) 21:425–532
Fig. 26 Perfusion imaging is crucial in detecting functional changes
in the vasculature. In normoxic pig myocardium 6 days after intramyocardial AdVEGF-B186 and AdVEGF-DΔNΔC gene transfer, myocardial perfusion is increased at stress conditions in the treated region
(gt) as measured with PET. Color scale is absolute; darkest blue is
0 ml/min/g, green is 1.5 ml/min/g, and deepest red is 3.0 ml/min/g
or over (a–c). In control AdCMV group, relative perfusion, i.e., the
ratio of absolute perfusion in the gene transfer area to that of the
control anteroseptal area, did not increase at stress, whereas with
both AdVEGF-B186 and AdVEGF-DΔNΔC, the relative perfusion was
higher in comparison with the control group. Relative perfusion at
rest was 12 and 13% and at stress 40 and 34% higher for AdVEGFB186 and AdVEGF-DΔNΔC, respectively d, e. Ad, adenoviral; CMV,
cytomegalovirus; ctrl, control anteroseptal area; gt, posterolateral
gene transfer area [654]
transduction of the heart. These new delivery methods bring
along new challenges to achieve efficacious transduction for
angiogenetic therapies.
In large animal pro-angiogenic studies, study endpoints
must be carefully decided. Therapeutic angiogenesis itself is
not the de facto aim. These neovessels should be functional
and enhance myocardial function. Therefore, functional analyses should have a significant role in evaluating the effects
of pro-angiogenic therapies. Previously, microspheres have
been used for the evaluation of myocardial perfusion, but
with improved technology, imaging modalities such as computed tomography (CT), magnetic resonance imaging (MRI),
and positron emission tomography (PET) should play more
prominent roles as real-time perfusion, anatomical and functional imaging can be performed simultaneously (Fig. 26).
These modalities also allow for a long-term follow-up, as
imaging can be performed sequentially, providing a better
understanding of the long-lasting effects of the treatments.
Potential limitation of angiogenic therapies is the potential effect of the treatment on neural networks of the heart.
It is entirely possible that the pro-angiogenic therapies
also induce alterations in nerves producing arrhythmias.
As a result, potential sudden deaths and other potential
arrhythmia-based problems should be carefully monitored.
Also, large animals do not have comorbidities that the
patients might possess. For instance, reactions to angiogenic
stimuli significantly vary in the ischemic and normoxic myocardium. Rapid growth of heart also causes challenges, as
relatively young tissue is more resistant to injury and can
regenerate better than tissues in elderly patients.
13
29.6 Concluding remarks
Large animal models are invaluable in neovascularization
studies, as they most closely resemble human patients. The
model must be carefully selected, and further thought must
be given when deciding suitable endpoints, which should
nowadays include functional imaging studies.
30 Guidelines for purity of recombinant
proteins in angiogenesis assays
This brief section aims to provide a few practical guidelines for purity requirements of recombinant proteins.
Indeed, many proteins are tested in the in vitro and in vivo
Angiogenesis (2018) 21:425–532
angiogenesis assays described throughout this article,
including pro- and anti-angiogenic cytokines or growth
factors and antibodies. Antibodies, owing to their long systemic half-life, are widely used in preclinical in vivo models.
While in the majority of cases reliable commercial sources
ensure high quality of reagents and hence reproducibility of
experimental results, at times controversial or unexpected
findings have emerged that may be, at least in part, traced
back to purity and quality control issues.
30.1 Endotoxin
One of the major confounders and a key source of variability, both in vitro and in vivo, is represented by endotoxin or
lipopolysaccharide (LPS) contamination [627, 628]. LPS is
an integral part of the outer cell membrane of Gram-negative bacteria and is released following bacterial lysis. LPS
consists of a lipid component, known as Lipid A, a core
oligosaccharide and a long heteropolysaccharide chain, the
O-specific chain, representing the surface antigen (O-antigen). The O-antigen is strain specific. Lipid A is the most
conserved component of endotoxin and is responsible for
the majority of the biological effects attributed to LPS [628,
629]. LPS is heat stable and is not destroyed under regular
sterilizing conditions. The limulus amebocyte lysate (LAL)
assay is commonly used for the detection of endotoxin,
and LAL kits are available from many manufacturers. This
assay is based on a cell lysate of the horseshoe crab Limulus
polyphemus that coagulates in the presence of even very low
Fig. 27 Effects of imidazole on endothelial cell proliferation. BCECs
(bovine choroidal microvascular endothelial cells) were cultured in
the presence of low-glucose DMEM in the presence of 10% bovine
calf serum as previously described [645]. Elution buffer was tested
up to a final concentration of 5 mM imidazole, with or without 10 ng/
ml VEGF165. Note that the concentration of imidazole typically used
to elute His-tagged proteins is 500 mM. After 5-6 days, cell proliferation was determined by fluorescence readings at 590 nm. Asterisks
denote significant differences compared to no addition (0) groups by t
test (***p < 0.001, **p < 0.01)
503
levels of endotoxins. Endotoxin levels are expressed as units
(EU): 1 EU is generally equivalent to 100 pg of LPS.
It is now well established that LPS interacts with tolllike receptor(TLR)-4 [630]. This is a key pathway in innate
immunity [631]. Upon activation, TLRs signal as dimers,
complexing via their intracellular Toll/interleukin-1 receptor
(TIR) domains with a family of adaptor proteins in TLRspecific patterns. This results in the activation of multiple
downstream pathways in a cell-type specific fashion [632].
It is also well established that, besides LPS, a variety of
endogenous and exogenous ligands can have agonistic or
antagonistic effects on TLR4, accounting for a wide variety
of biological effects [631].
30.2 Effects of endotoxin in angiogenesis assays
LPS can have diverse effects in angiogenesis-related assays.
It can induce both cell injury and activation in cultured
endothelial cells, even at very low concentrations [633, 634].
In vivo, LPS has been reported to induce tumor angiogenesis
[635] and metastasis as well as tumor invasion [636]. On the
other hand, a variety of studies have shown the opposite, i.e.,
anticancer effects of LPS in mouse models and in humans
[637]. Indeed, as above noted, TLR4 stimulation can activate
multiple signaling cascades, including the MAP kinase and
NF-kB pathways [632]. These pathways may activate the
secretion of proinflammatory cytokines, such as IL-6 and
IL-8, or anti-inflammatory type I IFNs, including IFN-γ.
To further complicate the picture, it has been reported that,
depending on p53 status, TLR4 stimulation may have opposite effects on promotion of breast cancer cell growth [638].
A common misconception is that only proteins expressed
in bacteria can be contaminated by LPS. In reality, a variety
of sources have been found to be contaminating [628]. Tap
water, air and people’s fingers can lead to endotoxin contamination. It has been reported that one of the most common
sources of endotoxin is laboratory water since distillation
and deionizing columns do not remove endotoxin [628].
Pyrogen-free water is helpful, but LPS can be still introduced
by inadequate handling of containers during washing procedures. A variety of chemical reagents and buffers are also
potential sources of endotoxin [628]. It is not uncommon to
find high levels of endotoxin in biological products such as
albumin, collagen or gelatin. Indeed, it has been argued that,
unless specific steps to prevent LPS contamination have been
taken and the LPS level is indicated, one should assume that
a reagent is almost certainly contaminated [628]. Therefore,
LPS levels ideally should be tested routinely and certainly
whenever unexpected results are obtained.
In biotech settings, the generally acceptable LPS levels
for antibodies or other protein reagents to be tested in preclinical studies are < 1 EU (~ 100 pg)/mg protein. Achieving such low levels requires significant planning and taking
13
504
multiple steps to prevent contamination [628], besides the
usual chromatographic steps aimed at achieving high purity
of the recombinant protein. Treatment of columns and equipment with 1 N NaOH for 1 h (> 10 h when using 0.1 N
NaOH) is typically employed to inactivate LPS. Methods
to remove LPS in contaminated preparations have also been
described, although their success is relatively mixed [629].
It is noteworthy that even some relatively recent controversies in the tumor angiogenesis field might be attributed,
at least in part, to LPS (or other contaminants). For example,
Paez-Ribes and colleagues reported in 2009 the unexpected
finding that the anti-VEGFR2 monoclonal antibody DC101
promotes invasiveness and metastasis in the Rip-Tag model of
insulinoma [530]. This finding was in conflict with previous
studies reporting inhibition of invasiveness by DC101 in other
models [639] and with subsequent reports that did not confirm
induction of tumor invasiveness and metastasis by DC101 in
the Rip-Tag model [557, 640]. While the DC101 employed
in the majority of studies was purified and distributed by
ImClone Systems [641], Paez-Ribes and colleagues purified
the antibody themselves from supernatants of hybridoma cells
that had been deposited at the American Tissue Culture Collection [530]. Unfortunately, the authors did not describe any
steps to remove LPS nor did they report the LPS levels in their
preparations of DC101 [530]. Given the high doses of DC101
required for in vivo studies (100 mg/kg/week), it is conceivable that even a relatively modest contamination may affect
the results. It remains to be determined whether LPS was truly
responsible for the reported tumor invasiveness.
However, LPS is by no means the only confounder in angiogenesis-related assays. Commonly used reagents or contaminants, for example detergents employed to solubilize samples
or misfolded or aggregated proteins, can have major effects on
endothelial cells. In the course of earlier studies in which we
screened libraries of secreted proteins to identify growth stimulators and inhibitors [642], we observed a surprisingly high rate
of inhibitory “hits” using an early version of the library [643].
However, the majority of these “hits” could not be confirmed
using more stringently purified proteins. It became apparent
that basal or VEGF-stimulated endothelial cell growth could
be easily inhibited by a variety of impurities, as above noted.
Imidazole, the buffer typically employed to elute His-tagged
protein bound to Nickel columns, a common procedure to purify
recombinant proteins, was also found to have profound effects
in endothelial cell proliferation assays. Figure 27 illustrates the
effects of imidazole on proliferation of bovine microvascular endothelial cells. As little as 3 mM significantly inhibited
VEGF-stimulated endothelial cell growth. At 4–5 mM imidazole, there was a dramatic inhibition of both basal and VEGFstimulated endothelial cell growth. Considering that 500 mM
is the concentration of imidazole commonly used to elute proteins from Nickel columns, a several hundred-fold final dilution
or, preferably, a buffer exchange is advisable before adding the
13
Angiogenesis (2018) 21:425–532
sample to endothelial cells. It has been suggested that similar
issues might account for the opposite reports by two groups on
the effects on endothelial cell growth of VEGF-Ax, a VEGF
isoform arising from read through translation [644, 645].
30.3 Concluding remarks
It is clear from the above that contamination of reagents can
have major effects in both in vitro and in vivo assays. The
most well-known contaminant with major consequences is
bacterial endotoxin, or LPS. Endotoxin is not only present in
recombinant preparations but can be co-purified from many
other sources. Next to endotoxin, other contaminants can
have major effects as well. It is therefore required to check
compound preparations for the presence of contaminants and
take action for purification when necessary.
31 Conclusions
This paper describes a large number of assays and analysis
methods that can be used for the assessment of angiogenesis. A proper interpretation of these assays is warranted for
obtaining relevant mechanistical insight into the sequence of
angiogenesis processes in different tissues and under changing conditions. We have aimed to provide insight into the use
and interpretation of the mostly used assays and have tried
to provide the reader with the challenges, limitations, and
pitfalls in making use of these assays.
Acknowledgements We apologize for not being able to cite the work of
all other studies related to this topic because of space restrictions. This
work was supported by funding from: Dutch Cancer Society (VU20125480 to JRvB and AWG; VU2014-7234 to AWG and PNS); European
Research Council (ERC) Starting Research Grant (EU-ERC680209,
to PNS); Advanced Research Grant (EU-ERC269073 to PC); FWO
Postdoctoral Fellowships (to JK); Federal Government Belgium
grant (IUAP P7/03 to PC); long-term structural Methusalem funding
by the Flemish Government (to PC); Research Foundation Flanders
(FWO, to PC); Foundation Leducq Transatlantic Network (ARTEMIS, to PC); Foundation Against Cancer (to PC); AXA Research
Fund (to PC); INSERM recurrent funding (to AB); Association pour
la Recherche sur le Cancer (ARC, to AB); Ligue Nationale du Cancer
(to AB); Swiss National Science Foundation (31003A_159824 and
CRSII3_154499 to CR); British Heart Foundation (to AMR and ND).
Research reported in this manuscript was supported by the National
Institutes of Health under Award Nos: R01 HL096384, R01 HL27030
to JB; R01 AR069038, R01 HL128452 and R21 AI123883 to JMMM; R01 CA188404 and R01 CA201537 to GB. The content is solely
the responsibility of the authors and does not necessarily represent the
official views of the National Institutes of Health.
Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativeco
mmons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate
credit to the original author(s) and the source, provide a link to the
Creative Commons license, and indicate if changes were made.
Angiogenesis (2018) 21:425–532
Abbreviations
AAV: Adeno-associated virus; AMD: Age-related
macular degeneration; BME: Basement membrane
extracts; BOEC: Blood outgrowth endothelial cells;
BrdU: 5-Bromo-2′-deoxyuridine; CAF: Cancer-activated fibroblasts; CAM: Chorioallantoic membrane;
CD31: Platelet endothelial cell adhesion molecule;
CD45: Leukocyte common antigen; CD144: Vascular
endothelial cadherin; CNV: Choroidal neovascularization;
CRC: Colorectal carcinoma; CT: Computer tomography;
D: Definitive differentiated cells; DAPI: 4′,6-Diamidino2-phenylindole; EACA: Epsilon aminocaproic acid;
EBM: Endothelial basal medium; EC: Endothelial cell(s);
ECAR: Extracellular acidification rate; ECFC: Endothelial colony-forming cells; ECM: Extracellular matrix;
EdU: 5-Ethynyl-2′deoxyuridine; ELISA: Enzyme-linked
immunosorbent assay; ENU: N-ethyl-N-nitrosourea;
EPC: Endothelial progenitor cells; EST: Expressed
sequence tag; EVP: Endovascular progenitor cells;
FGF: Fibroblast growth factor; FACS: Fluorescenceactivated cell sorting; GFP: Green fluorescent protein; GS-B4: Griffonia simplicifolia isolectin B4;
HDMEC: Foreskin-derived human dermal microvascular
EC; H&E: Hematoxylin and eosin; HIF: Hypoxia-inducible factor; HILIC: Hydrophilic liquid chromatography;
hiPSC: Human-induced pluripotent stem cell; HGP: Histopathological growth pattern; HSVSMC: Saphenous vein
smooth muscle cells; HUAEC: Human umbilical artery
EC; HUVEC: Human umbilical vein EC; ICAM: Intercellular adhesion molecule; IL-3: Interleukin 3; LDPI: Laser
Doppler perfusion imaging; LPS: Lipopolysaccharide;
MACS: Magnetic-activated cell sorting; MDSC: Myeloid-derived suppressor cells; MMP: Matrix metalloproteinase; MO: Morpholino; MP: Main population;
MPC: Mesenchymal progenitor cells; MRI: Magnetic
resonance imaging; MSC: Mesenchymal stem cells;
MTT: 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; NMR: Nuclear magnetic resonance;
NV: Neovascularization; OCR: Oxygen consumption rate;
OIR: Oxygen-induced retinopathy; oxPPP: Oxidative pentose phosphate pathway; PAD: Peripheral arterial disease;
PCNA: Proliferating cell nuclear antigen; PCR: Polymerase chain reaction; PCV: Polypoidal choroidal vasculopathy; PDMS: Polydimethylsiloxane; PET: Positron
emission tomography; PI: Propidium iodide; PNET: Pancreatic neuroendocrine tumor; PV: Particle velocimetry;
PWG: Postnatal weight gain; RCC: Renal cell carcinoma;
ROP: Retinopathy of prematurity; RS II: Reduced serum
supplement II; RTCA: Real-time cell analysis; SCF: Stem
cell factor; SCID: Severe combined immune deficiency;
SDF: Stromal-derived factor; SEM: Scanning electron
microscopy; SMA: Smooth muscle actin; SOD: Superoxide dismutase; SP: Side-population; SPARC: Secreted
505
protein acidic and rich in cysteine; TA: Transient amplifying cells; TAM: Tumor-associated macrophages;
TCA: Tetracarboxylic acid; TEM: Tumor endothelial
marker; TLR: Toll-like receptor; TME: Tumor microenvironment; TNF: Tumor necrosis factor; UEA: Ulex
europaeus agglutinin; VCAM: Vascular cell adhesion
molecule; VEGF: Vascular endothelial cell growth factor;
VESC: Vascular endothelial stem cells; VHL: Von Hippel–Lindau; VMO: Vascularized micro-organ; VO: Vasoobliteration; WGA: Wheat germ agglutinin
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Affiliations
Patrycja Nowak‑Sliwinska1,2 · Kari Alitalo3 · Elizabeth Allen4 · Andrey Anisimov3 · Alfred C. Aplin5 ·
Robert Auerbach6 · Hellmut G. Augustin7,8,9 · David O. Bates10 · Judy R. van Beijnum11 · R. Hugh F. Bender12 ·
Gabriele Bergers13,4 · Andreas Bikfalvi14 · Joyce Bischoff15 · Barbara C. Böck7,8,9 · Peter C. Brooks16 ·
Federico Bussolino17,18 · Bertan Cakir19 · Peter Carmeliet20,21 · Daniel Castranova22 · Anca M. Cimpean23 ·
Ondine Cleaver24 · George Coukos25 · George E. Davis26 · Michele De Palma27 · Anna Dimberg28 ·
Ruud P. M. Dings29 · Valentin Djonov30 · Andrew C. Dudley31,32 · Neil P. Dufton33 · Sarah‑Maria Fendt34,35 ·
Napoleone Ferrara36 · Marcus Fruttiger37 · Dai Fukumura38 · Bart Ghesquière39,40 · Yan Gong19 ·
Robert J. Griffin29 · Adrian L. Harris41 · Christopher C. W. Hughes12 · Nan W. Hultgren12 · M. Luisa Iruela‑Arispe42 ·
Melita Irving25 · Rakesh K. Jain38 · Raghu Kalluri43 · Joanna Kalucka20,21 · Robert S. Kerbel44 · Jan Kitajewski45 ·
Ingeborg Klaassen46 · Hynda K. Kleinmann47 · Pieter Koolwijk48 · Elisabeth Kuczynski44 · Brenda R. Kwak49 ·
Koen Marien50 · Juan M. Melero‑Martin51 · Lance L. Munn38 · Roberto F. Nicosia5,52 · Agnes Noel53 · Jussi Nurro54 ·
Anna‑Karin Olsson55 · Tatiana V. Petrova56 · Kristian Pietras57 · Roberto Pili58 · Jeffrey W. Pollard59 · Mark J. Post60 ·
Paul H. A. Quax61 · Gabriel A. Rabinovich62 · Marius Raica23 · Anna M. Randi33 · Domenico Ribatti63,64 ·
Curzio Ruegg65 · Reinier O. Schlingemann46,48 · Stefan Schulte‑Merker66 · Lois E. H. Smith19 · Jonathan W. Song67,68 ·
Steven A. Stacker69 · Jimmy Stalin66 · Amber N. Stratman22 · Maureen Van de Velde53 · Victor W. M. van Hinsbergh48 ·
Peter B. Vermeulen50,72 · Johannes Waltenberger70 · Brant M. Weinstein22 · Hong Xin36 · Bahar Yetkin‑Arik46 ·
Seppo Yla‑Herttuala54 · Mervin C. Yoder71 · Arjan W. Griffioen11
1
Molecular Pharmacology Group, School of Pharmaceutical
Sciences, Faculty of Sciences, University of Geneva,
University of Lausanne, Rue Michel-Servet 1, CMU,
1211 Geneva 4, Switzerland
2
Translational Research Center in Oncohaematology,
University of Geneva, Geneva, Switzerland
3
Wihuri Research Institute and Translational Cancer Biology
Program, University of Helsinki, Helsinki, Finland
4
Laboratory of Tumor Microenvironment and Therapeutic
Resistance, Department of Oncology, VIB-Center for Cancer
Biology, KU Leuven, Louvain, Belgium
5
Department of Pathology, University of Washington, Seattle,
WA, USA
6
University of Wisconsin, Madison, WI, USA
7
European Center for Angioscience, Medical Faculty
Mannheim, Heidelberg University, Heidelberg, Germany
13
8
Division of Vascular Oncology and Metastasis Research,
German Cancer Research Center, Heidelberg, Germany
9
German Cancer Consortium, Heidelberg, Germany
10
Division of Cancer and Stem Cells, School of Medicine,
University of Nottingham, Nottingham, UK
11
Angiogenesis Laboratory, Department of Medical Oncology,
VU University Medical Center, Cancer Center Amsterdam,
De Boelelaan 1117, 1081 HV Amsterdam, The Netherlands
12
Department of Molecular Biology and Biochemistry,
University of California, Irvine, CA, USA
13
Department of Neurological Surgery, Brain Tumor Research
Center, Helen Diller Family Comprehensive Cancer Center,
University of California, San Francisco, CA, USA
14
Angiogenesis and Tumor Microenvironment Laboratory
(INSERM U1029), University Bordeaux, Pessac, France
Angiogenesis (2018) 21:425–532
531
15
Vascular Biology Program and Department of Surgery,
Harvard Medical School, Boston Children’s Hospital,
Boston, MA, USA
38
Edwin L. Steele Laboratories, Department of Radiation
Oncology, Massachusetts General Hospital and Harvard
Medical School, Boston, MA, USA
16
Center for Molecular Medicine, Maine Medical Center
Research Institute, Scarborough, ME, USA
39
Metabolomics Expertise Center, VIB Center for Cancer
Biology, VIB, Leuven, Belgium
17
Department of Oncology, University of Torino, Turin, Italy
40
18
Candiolo Cancer Institute-FPO-IRCCS, 10060 Candiolo,
Italy
Department of Oncology, Metabolomics Expertise Center,
KU Leuven, Leuven, Belgium
41
Molecular Oncology Laboratories, Oxford University
Department of Oncology, Weatherall Institute of Molecular
Medicine, John Radcliffe Hospital, Oxford, UK
19
Department of Ophthalmology, Harvard Medical School,
Boston Children’s Hospital, Boston, MA, USA
20
42
Laboratory of Angiogenesis and Vascular Metabolism,
Department of Oncology and Leuven Cancer Institute (LKI),
KU Leuven, Leuven, Belgium
MCDB, University of California, Los Angeles, CA, USA
43
Laboratory of Angiogenesis and Vascular Metabolism,
Center for Cancer Biology, VIB, Leuven, Belgium
Department of Cancer Biology, Metastasis Research Center,
The University of Texas MD Anderson Cancer Center,
Houston, TX, USA
44
Department of Medical Biophysics, Biological Sciences
Platform, Sunnybrook Research Institute, University
of Toronto, Toronto, ON, Canada
45
Department of Microscopic Morphology/Histology,
Angiogenesis Research Center, Victor Babes University
of Medicine and Pharmacy, Timisoara, Romania
Department of Physiology and Biophysics, University
of Illinois, Chicago, IL, USA
46
Department of Molecular Biology, Center for Regenerative
Science and Medicine, University of Texas Southwestern
Medical Center, Dallas, TX, USA
Ocular Angiogenesis Group, Departments of Ophthalmology
and Medical Biology, Academic Medical Center, University
of Amsterdam, Amsterdam, The Netherlands
47
The George Washington University School of Medicine,
Washington, DC, USA
48
Department of Ophthalmology, University of Lausanne,
Jules-Gonin Eye Hospital, Fondation Asile des Aveugles,
Lausanne, Switzerland
49
Department of Pathology and Immunology, University
of Geneva, Geneva, Switzerland
50
HistoGeneX, Antwerp, Belgium
51
Department of Immunology, Genetics and Pathology,
Uppsala University, Uppsala, Sweden
Department of Cardiac Surgery, Harvard Medical School,
Boston Children’s Hospital, Boston, MA, USA
52
29
Department of Radiation Oncology, University of Arkansas
for Medical Sciences, Little Rock, AR, USA
Pathology and Laboratory Medicine Service, VA Puget
Sound Health Care System, Seattle, WA, USA
53
30
Institute of Anatomy, University of Bern, Bern, Switzerland
Laboratory of Tumor and Developmental Biology,
GIGA-Cancer, University of Liège, Liège, Belgium
31
Department of Microbiology, Immunology, and Cancer
Biology, University of Virginia, Charlottesville, VA, USA
54
Department of Biotechnology and Molecular Medicine,
University of Eastern Finland, Kuopio, Finland
32
Emily Couric Cancer Center, The University of Virginia,
Charlottesville, VA, USA
55
33
Vascular Sciences, Imperial Centre for Translational
and Experimental Medicine, National Heart and Lung
Institute, Imperial College London, London, UK
Department of Medical Biochemistry and Microbiology,
Science for Life Laboratory, Uppsala Biomedical Center,
Uppsala University, Uppsala, Sweden
56
Department of oncology UNIL-CHUV, Ludwig Institute
for Cancer Research Lausanne, Lausanne, Switzerland
57
Division of Translational Cancer Research, Department
of Laboratory Medicine, Lund, Sweden
58
Genitourinary Program, Indiana University-Simon Cancer
Center, Indianapolis, IN, USA
59
Medical Research Council Centre for Reproductive Health,
College of Medicine and Veterinary Medicine, University
of Edinburgh, Edinburgh, UK
60
Department of Physiology, Maastricht University, Maastricht,
The Netherlands
61
Einthoven Laboratory for Experimental Vascular Medicine,
Department Surgery, LUMC, Leiden, The Netherlands
21
22
23
24
Division of Developmental Biology, Eunice Kennedy Shriver
National Institute of Child Health and Human Development,
National Institutes of Health, Bethesda, MD, USA
25
Ludwig Institute for Cancer Research, Department
of Oncology, University of Lausanne, Lausanne, Switzerland
26
Department of Medical Pharmacology and Physiology,
University of Missouri, School of Medicine and Dalton
Cardiovascular Center, Columbia, MO, USA
27
28
34
35
School of Life Sciences, Swiss Federal Institute
of Technology, Lausanne, Switzerland
Laboratory of Cellular Metabolism and Metabolic
Regulation, VIB Center for Cancer Biology, Leuven,
Belgium
Laboratory of Cellular Metabolism and Metabolic
Regulation, Department of Oncology, KU Leuven
and Leuven Cancer Institute, Leuven, Belgium
36
University of California, San Diego, La Jolla, CA, USA
37
Institute of Ophthalmology, University College London,
London, UK
13
532
62
63
Angiogenesis (2018) 21:425–532
Laboratory of Immunopathology, Institute of Biology
and Experimental Medicine, National Council of Scientific
and Technical Investigations (CONICET), Buenos Aires,
Argentina
Department of Basic Medical Sciences, Neurosciences
and Sensory Organs, University of Bari Medical School,
Bari, Italy
64
National Cancer Institute “Giovanni Paolo II”, Bari, Italy
65
Department of Oncology, Microbiology and Immunology,
Faculty of Science and Medicine, University of Fribourg,
Fribourg, Switzerland
66
Institute of Cardiovascular Organogenesis and Regeneration,
Faculty of Medicine, WWU, Münster, Germany
13
67
Department of Mechanical and Aerospace Engineering, The
Ohio State University, Columbus, OH, USA
68
Comprehensive Cancer Center, The Ohio State University,
Columbus, OH, USA
69
Tumour Angiogenesis and Microenvironment Program, Peter
MacCallum Cancer Centre and The Sir Peter MacCallum,
Department of Oncology, University of Melbourne,
Melbourne, VIC, Australia
70
Medical Faculty, University of Münster,
Albert-Schweitzer-Campus 1, Münster, Germany
71
Department of Pediatrics, Indiana University School
of Medicine, Indianapolis, IN, USA
72
Translational Cancer Research Unit, GZA Hospitals,
Sint-Augustinus & University of Antwerp, Antwerp, Belgium
Andreas Bikfalvi