Consensus guidelines for the use and interpretation of angiogenesis assays

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.

Table of contents

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

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

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 [1,2,3,4,5]. 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 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 [6,7,8] 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 [9,10,11]. 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 [15,16,17,18,19,20], (4) the unraveling of mechanisms regulating lymphangiogenesis [21, 22], (5) the interrelationship of angiogenesis and immunity [23,24,25], (6) the development of imaging as diagnostic strategy [26], (7) the study of drug resistance mechanisms [27,28,29], (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.

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 [34,35,36,37], 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 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.

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 [34,35,36]. 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 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).

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 video-lapse 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].

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 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.essenbioscience.com/en/products/incucyte). 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.vp-scientific.com/wounding_tissue_culture_experiments.php) by a Sciclone ALH 3000 workstation robot (Caliper Life Sciences, Hopkinton, MA). The additional precision of the 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).

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.

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 researcher-dependent observations [57]. This section presents different methods and elaborates on problems and pitfalls.

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.

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

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.

³H-thymidine has been used in proliferation assays for decades [19, 56, 58, 59]. Briefly, cells are pulsed with ³H-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].

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-(3-carboxymethoxyphenyl)-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].

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 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 from human, mouse, and viral sources that stimulate the VEGFR-2 and VEGFR-3 pathways [69, 70].

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.

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.

Table 1 Comparison of 3D models of vascular morphogenesis

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 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].

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 phase-contrast 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 micro-organ (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)

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-β-induced 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 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 [76,77,78], 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 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, whole-organ 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 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 time-lapse 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 [87,88,89]. 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 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 shear-sensing), 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 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.

Limitations and challenges

With all EC assays, the source of EC is critical to assay success. Although commercial versions of HUVEC and ECFC-EC 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.

Concluding remarks

Vascular morphogenesis requires the interaction between several cell types and their surrounding, 3D 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.

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 pro-angiogenic or anti-angiogenic compounds [100].

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].

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 aorta-derived 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)

Recently, rat aortic ring assay was adapted to human arteries by using matrigel as 3D hydrogel [102].

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 cross-sectioned 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.

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 [105,106,107,108]. 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.

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 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 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.

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 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 [105,106,107,108]. Finally, for the whole-mount immunohistochemical stain, gel thinness is critical for optimal antibody penetration.

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.

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 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.

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 hematoxylin-and-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 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 pan-endothelial 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 whole-slide 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).

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 (*)

Several groups have identified “tumor endothelial markers” or TEMs [123,124,125]. 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 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 and strong link between angiogenesis and immune suppression [129,130,131] 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].

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.

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 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.

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.

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].

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 micro-CT 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].

Fig. 5

Adapted from [648]

Intussusceptive angiogenesis—the methodological challenge. 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.

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.

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 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).

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 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.

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 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).

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-D-ΔNΔC. 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

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.addgene.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.

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 anti-CD-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 FITC-lectin (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 post-implantation (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 long-lasting 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 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.

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 half-way 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, 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 AAV-binding 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.

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.

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, 152,153,154,155]. To this end, bioassays have been developed in 3D collagen or fibrin matrices to address such issues using serum-free defined conditions [156, 157] (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].

Fig. 7

Serum-free defined model of human endothelial cell-pericyte tube co-assembly in 3D collagen matrices. a Human EC and GFP-labeled 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 supplement), 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

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 co-assemble to form tube networks with abluminally recruited pericytes in 3D matrices.

Assay overview

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 stromal-derived 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 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].

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, 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.

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 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.

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.

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 [161,162,163]. Anatomically, these various cell types are fine-tuned 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. Co-cultures 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 medium-throughput manner in an environment more relevant to the in vivo situation.

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].

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 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].

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

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). Gravity-enforced 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 re-inverted, 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 the additions of, for example, growth factors, therapeutics, or reporters (e.g., viability or proliferation stains; Fig. 8).

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.

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.

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 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.

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 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.

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 non-radioactive 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, ¹⁶O, 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 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.

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.

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].

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].

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 (OCR_ATP; 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 (OCR_BAS; green), the maximal mitochondrial OCR (OCR_MAX; orange), the proton leakage (blue), and the non-mitochondrial OCR (pink)

In order to map the different metabolic fates of glucose and other crucial substrates, a set of differently radioactive-labeled 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 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.

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 (OCR_BAS), ATP-dependent OCR (OCR_ATP), maximal respiration (OCR_MAX) and non-mitochondrial respiration (OCR_non-mito), proton leak, and spare respiration (Fig. 9c) [184].

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 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. 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.

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 colony-forming 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.

Assays to identify endothelial stem and progenitor cells

A simple working definition of a stem cell is a clonal, self-renewing 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 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 long-term 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 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, 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].

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 [195,196,197]; 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.

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.

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.

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 fluid-filled 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]. 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 [208,209,210]. 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 [212,213,214,215], stromal fibroblasts [216, 217], vascular smooth muscle cells [218], and bone marrow cells [219]. Applications for these microfluidic co-culture 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 flow-mediated angiogenesis (Fig. 10). For instance, multiple studies have shown that interstitial flow potently induces angiogenesis [220,221,222] 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 [220,221,223,223]. 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].

Fig. 10

(Adapted from [221])

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, through the central matrix gel. Sprouting occurs through the apertures that flank the central 3D matrix and is easily visualized and quantified

One limitation to PDMS replica molding is that this process produces microchannels and subsequent endothelial-lined 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 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].

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 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].

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].

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.

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, carcinoma-associated 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, time-consuming, 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 [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].

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 pro-angiogenic 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.

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 single-cell suspension after tissue dissociation, erythrocytes lysis, and cell labeling with fluorescent or paramagnetic-coupled antibodies. Analysis of blood-circulating 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

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 pro-angiogenic 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 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.

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 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.

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.

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.

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 [275,276,277,278]. 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 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. MO-based 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, 281,282,283,284,285,286,287,288,289,290,291,292,293,294,295,296,297,298,299,300,301]. 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, translation-blocking 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 MOs a better representation of targeted loss of gene function in these cases.

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 [302,303,304,305,306,307,308,309]. 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, genome-editing technologies and particularly CRISPR/Cas9-based methods have made reverse genetic targeting of virtually any gene easy and cost-effective for most laboratories [281,282,283]. 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.

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.

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.

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 vascular-autonomous 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 experiments or tissue-specific transgenic expression to “rescue” the phenotype (Fig. 12f–h).

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 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]

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 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 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 [314,315,316] 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.

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 (fss^y66) mutant (j) animals. Improper somite formation in fss^y66 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

Assessing circulatory flow and cardiac function

Even in animals with apparently normal overall development, defective blood flow (due, for example, to heart-specific 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 [317,318,319,320]. 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.

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 [324,325,326]. 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, 327,328,329,330,331,332]. 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 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).

Concluding remarks

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-of-function phenotypes, they remain valuable when rigorously 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 cell-autonomous 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.

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 [335,336,337,338,339,340,341,342,343]. There are certainly several hundreds of variations in the experimental protocol, all of them involving placing a test material on the CAM or an 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.