This work is in part supported by a research grant from the &#8216;Meer Kennis met Minder Dieren&#8217; program (project number 114022501) of The Netherlands Organization for Health Research and Development (ZonMw) and the Dutch Heart Foundation CVON consortium grant RECONNECT.

1. Device Preparation
<ol>
	<li>Transfer the microfluidic 384-well plate to a sterile laminar-flow hood.</li>
	<li>Remove the lid and add 50 &#181;L of water or phosphate-buffered saline (PBS) to each of the 40 observation wells (Figure 1b, well B2) using a multichannel or repeating pipette. 
	NOTE: The protocol can be paused here. Leave the plate with the lid on in the sterile culture cabinet at room temperature (RT).</li>
</ol>
2. Prepare Gel and Coating
<ol>
	<li>Prepare 2.5 mL of 10 &#181;g/mL fibronectin (FN) coating solution. Dilute 25 &#181;L of 1 mg/mL fibronectin stock solution in 2.5 mL of Dulbecco&#8217;s PBS (dPBS, calcium and magnesium free). Place the solution in the water bath at 37 &#176;C till use.</li>
	<li>Prepare 100 &#181;L of collagen-1 solution. Add 10 &#181;L of HEPES (1 M) to 10 &#181;L of NaHCO3 (37 g/L) and mix by pipetting. Place the tube on ice and add 80 &#181;L of collagen-1 (5 mg/mL) to yield a neutralized collagen-1 concentration of 4 mg/mL. Use a pipette to mix carefully and avoid the formation of bubbles.</li>
	<li>Add 1.5 &#181;L of collagen-1 solution (4 mg/mL) to the gel inlet of each microfluidic unit (Figure 1b, well B1). Make sure the droplet of gel is placed in the middle of each well in order for the gel to enter the channel (see Figure 2a). 
	NOTE: Phaseguides prevent filling of the adjacent channels and enables gel patterning. Correct gel loading can be confirmed under a brightfield microscope by observing the meniscus formation through the &#8216;observation window&#8217; (well B2) or by flipping the plate upside down. If the gel did not completely fill the channel, an additional droplet of 1 &#181;L can be added.</li>
	<li>Place the microfluidic plate in an incubator (37 &#176;C, 5% CO2) for 10 min to polymerize collagen-1. 
	NOTE: Timing of the polymerization is crucial; due to the low volumes used in microfluidics, evaporation can already be observed after 15 min of incubation, which results in gel collapse or shrinkage.</li>
	<li>Take the plate out of the incubator and transfer to a sterile laminar-flow hood.</li>
	<li>Add 50 &#181;L of 10 &#181;g/mL FN coating solution to the outlet well of the top perfusion channel of every microfluidic unit (Figure 1b, well A3). Press the pipette tip against the side of the well for correct filling of the well without trapping air bubbles (see Figure 2b). The channel should fill, and the liquid should pin on the inlet (well A1) without filling the inlet well.</li>
	<li>Place the plate in the incubator (37 &#176;C, 5% CO2) for at least 2 days. 
	NOTE: The protocol can be paused here, as the collagen-1 gel together with the coating mixture is stable for at least 5 days in the incubator. If the coating mixture is refreshed, longer periods could be possible, but this has not been tested. Crucial is the level of FN-coating, as this prevents the dehydration of the collagen-1 gel.</li>
</ol>
3. Cell Seeding/Microvessel Culture
<ol>
	<li>Add 5 mL of fetal calf serum and 2.5 mL of pen/strep to 500 mL of basal endothelial cell culture medium and filter sterilize using a bottle top filter with 0.22 &#181;m pore size. This medium is now referred to as basal medium.</li>
	<li>Prepare vascular growth medium: Add 3 &#181;L of 50 &#181;g/mL vascular endothelial growth factor (VEGF) and 2 &#181;L of 20 &#181;g/mL basic fibroblast growth factors (bFGF) to 5 mL of basal medium.</li>
	<li>Thaw the frozen iPSC-ECs rapidly (&#60;1 min) in a 37 &#176;Cwater bath and transfer to a 15 mL tube and dilute in 10 mL of basal medium.</li>
	<li>Count the cells. 
	NOTE: A single vial contains 1 million cells in 0.5 mL with &#62;90% viability.</li>
	<li>Centrifuge the tube at 100 x g for 5 min. Aspirate supernatant without disturbing the cell pellet and resuspend in basal medium to yield a concentration of 2 x 107 cells/mL.</li>
	<li>Transfer the microfluidic 384-well plate from the incubator to a sterile laminar-flow hood.</li>
	<li>Aspirate FN-coating solution from the perfusion outlet well (A3) and replace with 25 &#181;L of basal medium in the outlet well (well A3).</li>
	<li>Add a 1 &#181;L droplet of cell suspension to every top perfusion inlet well (Figure 1b, well A1). The droplet should flatten in a few seconds (see Figure 3a,b for an illustration of this &#8216;passive pumping&#8217; method). 
	NOTE: Check under a microscope whether the seeding is homogeneous. If not, add another 1 &#181;L in the outlet and wait till the droplet flattens.</li>
	<li>Incubate the microfluidic well-plate for 1 h at 37 &#176;C, 5% CO2. After this, the cells should have adhered. If not, wait another 30 min.</li>
	<li>Remove the basal medium from the top perfusion outlet wells (Figure 1b, well A3). Add warm vessel culture medium in the top perfusion inlet and outlet (Figure 1b, wells A1 and A3). Place the plate on the rocker platform (set on 7&#176; angle, 8 min rocking interval) in the incubator (37 &#176;C, 5% CO2).</li>
	<li>Image the plate using a brightfield microscope with automated stage at day 1 and 2 post-seeding to confirm cell viability. After 2 days, a confluent monolayer should have formed against the collagen-1 scaffold. 
	NOTE: If the channels do not appear to be equally confluent, the microvessels can be cultured for an additional 24 h.</li>
</ol>
4. Study Angiogenic Sprouting Including Tip- and Stalk-cell Formation
<ol>
	<li>Prepare 4.5 mL of angiogenic sprouting medium by supplementing basal medium with 4.5 &#181;L of VEGF (50 &#181;g/mL stock), 4.5 &#181;L of phorbol 12-myristate-13-acetate (PMA) (2&#160;&#181;g/mL stock), and 2.25 &#181;L of sphingosine-1-phosphate (S1P) (1 mM stock).</li>
	<li>Prepare 8.5 mL of vessel growth medium (basal medium supplemented with 30 ng/mL VEGF and 20 ng/mL bFGF).</li>
	<li>Aspirate medium from the wells and add 50 &#181;L of fresh vessel culture medium in the top perfusion inlet and outlet wells and gel inlet and outlet wells (Figure 1b, wells A1, A3, B1 and B3).</li>
	<li>Add 50 &#181;L of angiogenic sprout mixture to each of the bottom perfusion channel inlet and outlet wells (Figure 1b, wells C1 and C3).</li>
	<li>Place the device back in the incubator (37 &#176;C, 5% CO2) on the rocker platform in order to form a gradient of angiogenic growth factors.</li>
	<li>Image 1 day and 2 days after addition of the angiogenic growth factors using a brightfield microscope with automated stage. 
	NOTE: Continue culturing the microvessels to study anastomosis (go to section 5) or fix and stain the microvessels to quantify sprouting length and morphology at day 2 and day 6 (go to section 6).</li>
</ol>
5. Study Anastomosis and Sprout Stabilization
<ol>
	<li>Image using a brightfield microscope with automated stage.</li>
	<li>Add 1 &#181;L of fluorescently labeled albumin (0.5 mg/mL) to the top perfusion inlet (Figure 1b, well A1) and mix using a 50 &#181;L pipette.</li>
	<li>Transfer the plate to a fluorescent microscope with automated stage and incubator set at 37 &#176;C. Set microscope at 10x objective and correct exposure settings (e.g., tetramethylrhodamine [TRITC]-channel, 20 ms exposure). Acquire time-lapse images every minute for 10 min.</li>
	<li>Remove the plate from the microscope and transfer the plate to a sterile laminar-flow hood.</li>
	<li>Remove all medium from the wells and replace both the vessel culture medium and angiogenic sprouting medium in the corresponding wells (see steps 4.3 and 4.4).</li>
	<li>Place the device back into the incubator (37 &#176;C, 5% CO2) to continue angiogenic sprouting.</li>
	<li>Repeat steps 5.1&#8722;5.6 to study the permeability at day 6.</li>
</ol>
6. Fixation, Staining, and Imaging
<ol>
	<li>Aspirate all culture media from all the wells. 
	NOTE: Residual medium or liquids in the microfluidic channels does not influence the fixation due to its low volume of 1&#8722;2 &#181;L.</li>
	<li>Add 25 &#181;L of 4% paraformaldehyde (PFA) in PBS to all the perfusion inlet (Figure 1b, A1 and C1) and outlet wells (Figure 1b, A3 and C3). Incubate for 10 min at RT. Place the device under a slight angle (&#177;5&#176;) to induce flow (e.g., by placing one side of the plate on a lid).</li>
	<li>Aspirate PFA from the wells. Wash all the perfusion inlets and outlets twice with 50 &#181;L of Hank&#8217;s balanced salt solution (HBSS). Aspirate HBSS from wells.</li>
	<li>Permeabilize at RT for 10 min by adding 50 &#181;L of 0.2% nonionic surfactant to all the perfusion inlets and outlets.</li>
	<li>Aspirate the nonionic surfactant from wells. Wash the perfusion channels twice by adding 50 &#181;L of HBSS to all perfusion inlet and outlet wells. Aspirate HBSS from wells.</li>
	<li>Stain the nuclei using Hoechst (1:2,000) and F-actin using phalloidin (1:200) in HBSS. Prepare 2.2 mL for 40 units, and add 25 &#181;L to each perfusion inlet and outlet well. Place the plate under a slight angle and incubate at RT for at least 30 min.</li>
	<li>Wash twice with 50 &#181;L of HBSS in all the perfusion inlets and outlets.</li>
	<li>Directly image using a fluorescent microscope with automated stage or store the plate protected from light at 4 &#176;C for later use.</li>
</ol>

<ol>
	<li>Griffith, L. G., Swartz, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Capturing+complex+3D+tissue+physiology+in+vitro.">Capturing complex 3D tissue physiology in vitro.</a> Nature Reviews Molecular Cell Biology. 7 (3), 211-224 (2006).</li><li>Davis, G. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Control+of+vascular+tube+morphogenesis+and+maturation+in+3D+extracellular+matrices+by+endothelial+cells+and+pericytes.">Control of vascular tube morphogenesis and maturation in 3D extracellular matrices by endothelial cells and pericytes.</a> Methods in Molecular Biology. , 17-28 (2013).</li><li>Kim, S., Chung, M., Jeon, N. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+biomimetic+model+to+reconstitute+sprouting+lymphangiogenesis+in+vitro.">Three-dimensional biomimetic model to reconstitute sprouting lymphangiogenesis in vitro.</a> Biomaterials. 78, 115-128 (2016).</li><li>Kim, C., Kasuya, J., Jeon, J., Chung, S., Kamm, R. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+quantitative+microfluidic+angiogenesis+screen+for+studying+anti-angiogenic+therapeutic+drugs.">A quantitative microfluidic angiogenesis screen for studying anti-angiogenic therapeutic drugs.</a> Lab Chip. 15 (1), 301-310 (2015).</li><li>Kim, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineering+of+a+Biomimetic+Pericyte-Covered+3D+Microvascular+Network.">Engineering of a Biomimetic Pericyte-Covered 3D Microvascular Network.</a> PLoS One. 10 (7), e0133880 (2015).</li><li>Tourovskaia, A., Fauver, M., Kramer, G., Simonson, S., Neumann, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue-engineered+microenvironment+systems+for+modeling+human+vasculature.">Tissue-engineered microenvironment systems for modeling human vasculature.</a> Experimental biology and medicine. 239 (9), 1264-1271 (2014).</li><li>Junaid, A., Mashaghi, A., Hankemeier, T., Vulto, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+end-user+perspective+on+Organ-on-a-Chip:+Assays+and+usability+aspects.">An end-user perspective on Organ-on-a-Chip: Assays and usability aspects.</a> Current Opinion in Biomedical Engineering. 1, 15-22 (2017).</li><li>Pagano, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimizing+design+and+fabrication+of+microfluidic+devices+for+cell+cultures:+An+effective+approach+to+control+cell+microenvironment+in+three+dimensions.">Optimizing design and fabrication of microfluidic devices for cell cultures: An effective approach to control cell microenvironment in three dimensions.</a> Biomicrofluidics. 8 (4), 046503 (2014).</li><li>Berthier, E., Young, E. W., Beebe, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineers+are+from+PDMS-land,+Biologists+are+from+Polystyrenia.">Engineers are from PDMS-land, Biologists are from Polystyrenia.</a> Lab Chip. 12 (7), 1224-1237 (2012).</li><li>Nowak-Sliwinska, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Consensus+guidelines+for+the+use+and+interpretation+of+angiogenesis+assays.">Consensus guidelines for the use and interpretation of angiogenesis assays.</a> Angiogenesis. 21 (3), 425 (2018).</li><li>Dejana, E., Hirschi, K. K., Simons, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+molecular+basis+of+endothelial+cell+plasticity.">The molecular basis of endothelial cell plasticity.</a> Nature Communications. 8, 14361 (2017).</li><li>Geraghty, R. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Guidelines+for+the+use+of+cell+lines+in+biomedical+research.">Guidelines for the use of cell lines in biomedical research.</a> British Journal of Cancer. 111 (6), 1021-1046 (2014).</li><li>Passier, R., Orlova, V., Mummery, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Complex+Tissue+and+Disease+Modeling+using+hiPSCs.">Complex Tissue and Disease Modeling using hiPSCs.</a> Cell Stem Cell. 18 (3), 309-321 (2016).</li><li>van Duinen, V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perfused+3D+angiogenic+sprouting+in+a+high-throughput+in+vitro+platform.">Perfused 3D angiogenic sprouting in a high-throughput in vitro platform.</a> Angiogenesis. 22 (1), 157-165 (2019).</li><li>Wevers, N. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+perfused+human+blood-brain+barrier+on-a-chip+for+high-throughput+assessment+of+barrier+function+and+antibody+transport.">A perfused human blood-brain barrier on-a-chip for high-throughput assessment of barrier function and antibody transport.</a> Fluids Barriers CNS. 15 (1), 23 (2018).</li></ol>

P. Vulto and T. Hankemeier are shareholders of Mimetas BV. V. Borgdorff and A. Reijerkerk are employees of Ncardia BV. V. van Duinen, W. Stam, V. V. Orlova and A.J. van Zonneveld have no disclosures.

This method describes the culture of 40 perfusable endothelial microvessels within a robust and scalable microfluidic cell culture platform. Compared to traditional 2D and 3D cell culture methods, this method shows how a physiological relevant cellular microenvironment that includes gradients and continuous perfusion can be combined with 3D cell culture with adequate throughput for screening purposes.
One of the major advantages over comparable microfluidic assays is that this method does not rely on pumps for perfusion but uses a rocker platform to induce continuous perfusion in all microfluidic units simultaneously. This ensures that the assay is robust and scalable: plates can be stacked on a rocker platform. Importantly, all microfluidic units remain individually addressable, which allows this method to be implemented within drug screening including the generation of a dose-response curve. Furthermore, without a pump, imaging and medium replacement is far simpler with less risk of (cross)-contamination.
Another advantage of this method is usage of a standardized, pre-manufactured platform, while comparable microfluidic cell culture platforms need to be fabricated by the end-users. This availability facilitates the adoption of this assay among other academic and pharmaceutical research groups, leading to standardization. Also, unlike microfluidic prototypes, the 384-well plate interface ensures compatibility with the current lab equipment (e.g., aspirators, plate handlers and multichannel pipettes), facilitating the integration within the current screening infrastructure.
There are several critical steps in performing this assay. The collagen-1 gel should completely fill the gel channel. During gel loading, this filling can be observed by inspecting the microfluidic channels either through the observation window (Figure 2a) or by flipping the plate upside down (as shown in Figure 1a). While filling, the collagen gel should remain in the center channel, without flowing into adjacent perfusion channels. We noticed that the quality of the collagen-1 gel is crucial for proper assay performance. Collagen-1 batches with too high viscosity will lead to incomplete filling of the gel channel. After 10 min of polymerization at 37 &#176;C, the gel should be homogenous and clear. If collagen-1 is not stored properly (e.g., due to fluctuating temperatures in the fridge), collagen will polymerize within the channels with clearly visible fiber formation. This can result in invasion of the ECs into the gel without addition of angiogenic factors, but without proper lumen development.
When the cells are seeded, the fibronectin coating solution is removed from the wells, leaving only the microfluidic channels filled with coating solution. Aspiration of the coating solution from microfluidic channels could cause gel disruption or gel aspiration. The cell suspension needs to replace/displace this coating solution. This works best when the cell suspension is seeded using the passive pumping method, as directly pipetting the cell suspension into the channels show less reproducible seeding densities.
As the microvessels form a stable monolayer against the gel, these small differences only result in different times needed for reaching confluency. Thus, the assay start point is determined by confluency rather than culture time. If necessary, the culturing time can be extended until a clear monolayer has been formed against the gel.
Within the wells, air bubbles can be trapped by incorrect filling of the wells (see Figure 2b). These air bubbles will restrict the flow of medium, even when the device is placed on a rocker platform, and result in collapse of the microvessel and improper gradient formation. Pressing the pipette tip against the side wall of the wells will increase the success of completely filling the well. If an air bubble is trapped within the wells, it can be removed by gently inserting a sterile pipette tip into the glass bottom. Air bubbles can also occur within the microfluidic channels. When medium has been removed from the wells, evaporation of medium is noticeable from the microfluidic channels after 30 min (due to the microliter volumes within the channels). Thus, medium changes are preferably performed as quick as possible. When medium is added in a channel with evaporated medium, air bubbles will be trapped within the microfluidic channels. These air bubbles within the microfluidic channels can be removed manually by placing a P20 pipette directly on either the inlet or outlet and forcing medium through the microfluidics from the opposite well. Successful removal of the air bubbles results in a small but noticeable decrease in volume in the other well. Table 1 lists common errors and how to troubleshoot them.
The lack of a pump is a limitation when continuous imaging is required, as the rocker platform limits the user to image at sequential time intervals. Furthermore, the perfusion of medium in this platform consists of bi-directional flow with low levels of shear stress, while vasculature in vivo is exposed to unidirectional flow with higher levels of shear stress. While we do not observe negative effects of the bi-directional flow with regards to the angiogenic sprouting, flow is an important biomechanical stimulus and preferably controlled. However, while there are commercially available pump setups, interfacing with the 384-well plate remains challenging and pump setups severely hamper the scalability of this assay.
The possibility to use iPSC-ECs to study angiogenic sprouting opens up new opportunities in disease modeling and drug research. In contrast to primary ECs, these cells can be generated in nearly limitless quantities with a stable genotype and by using genome editing techniques, cells can be generated that including gene knockouts and knock-ins. However, as the protocols to differentiate ECs from iPSC are relatively new, it is still unclear what leads to iPSC-ECs that best reflect primary ECs and which subtypes of EC are or can be generated. Also, there are still remaining questions regarding their relevancy. For example, do iPSC-ECs still exhibit the plasticity that is typical for endothelial cells? And to what degree do iPSC-derived cells respond to and interact with their cellular microenvironment? The standardized platform presented here could be used to answer some of these questions in order to further validate the usage of iPSC-derived ECs in vitro.
The most straight-forward future direction for this assay will be the integration of other cell types that play an important role during angiogenesis, such as pericytes and macrophages. This will facilitate the ability to study the role of macrophages during anastomosis between sprouts or the adherence of pericytes after capillary formation. Also, it is possible to culture various other cell types within or against an extracellular matrix (e.g., we have shown the culture of neurons and various epithelial structures such as proximal tubules and small intestines), which can be combined with the vascular beds generated using this method. Finally, it will be interesting to study angiogenic sprouting in synthetic hydrogels, as their defined composition further increases the standardization of the assay and allows tuning of stiffness and binding motives that affect cell-matrix interactions.
In conclusion, this method shows the feasibility of iPSC-derived ECs in a standardized and scalable 3D angiogenic assay that combines physiological relevant culture conditions in a platform that has the required robustness and scalability to be integrated within the drug screening infrastructure.

An erratum was issued for:&#160;<a href="https://www.jove.com/t/59678">Standardized and Scalable Assay to Study Perfused 3D Angiogenic Sprouting of iPSC-derived Endothelial Cells In Vitro</a>. The Protocol section was&#160;updated.
Step 2.6 of the Protocol was updated from:
Add 50 &#181;L of 10 &#181;g/mL FN coating solution to the outlet well of the top perfusion channel of every microfluidic unit (Figure 1b, well A3). Press the pipette tip against the side of the well for correct filling of the well without trapping air bubbles (see Figure 2b). The channel should fill, and the liquid should pin on the outlet (well C1) without filling the outlet well.
to:
Add 50 &#181;L of 10 &#181;g/mL FN coating solution to the outlet well of the top perfusion channel of every microfluidic unit (Figure 1b, well A3). Press the pipette tip against the side of the well for correct filling of the well without trapping air bubbles (see Figure 2b). The channel should fill, and the liquid should pin on the inlet (well A1) without filling the inlet well.
Step 3.7 of the Protocol was updated from:
Aspirate FN-coating solution from the perfusion inlet (well A1). Add 25 &#181;L of basal medium in the inlet wells (well A1).
to:
Aspirate FN-coating solution from the perfusion outlet well (A3) and replace with 25 &#181;L of basal medium in the outlet well (well A3).

An erratum was issued for:&#160;<a href="https://www.jove.com/t/59678">Standardized and Scalable Assay to Study Perfused 3D Angiogenic Sprouting of iPSC-derived Endothelial Cells In Vitro</a>. The Protocol section was&#160;updated.

Erratum: Standardized and Scalable Assay to Study Perfused 3D Angiogenic Sprouting of iPSC-derived Endothelial Cells In Vitro

In vitro models play a fundamental role in the discovery and validation of new drug targets of vascular diseases. However, current in vitro screening models that have sufficient throughput only have limited physiological relevance1, which hinders the translation of findings from in vitro to in vivo. Thus, to advance pre-clinical vascular drug research, improved in vitro models of vasculature are necessary that combine high-throughput screening with a physiologically relevant 3D cellular micro-environment.
Within the last decade, significant progress has been made to increase the physiological relevance of in vitro models of vasculature. Instead of culturing endothelial cells on flat surfaces such as tissue-culture plastics, endothelial cells can be embedded in 3D scaffolds, such as fibrin and collagen gels2. Within these matrices, the endothelial cells show a more physiologically relevant phenotype associated with matrix degradation and lumen formation. However, these models only demonstrate a subset of the many processes that occur during angiogenic sprouting as important cues from the cellular microenvironment are still lacking.
Microfluidic cell culture platforms are uniquely suited to further increase the physiological relevance of in vitro models of vasculature. For example, endothelial cells can be exposed to shear stress, which is an important biomechanical stimulus for vasculature. Also, the possibility to spatially control fluids within microfluidics allows the formation of biomolecular gradients3,4,5,6. Such gradients play an important role in vivo during the formation and patterning during angiogenesis. However, while microfluidic cell culture platforms have shown unparalleled physiological relevancy over traditional 2D and 3D cell culture methods, they often lack the necessary throughput, scalability and standardization that is required for drug screening7. Also, many of these platforms are not commercially available and require the end-users to microfabricate their devices prior to use8. This not only requires manufacturing apparatus and technical knowledge, but also limits the level of quality control and negatively affects reproducibility9.
To date, primary human endothelial cells remain the most widely used cell source to model angiogenesis in vitro10. However, primary human cells have a number of limitations that hinder their routine application in screening approaches. First, there is a limited possibility to scale up and expand primary cell-derived cultures. Thus, for large scale experiment, batches from different donors need to be used, which result in genomic differences and batch-to-batch variations. Second, after a few passages, primary endothelial cells generally lose relevant properties when cultured in vitro11,12.
Endothelial cells derived from human induced pluripotent stem cells (iPSC) are a promising alternative: they resemble primary cells, but with a more stable genotype that is also amenable to precise genome editing. Furthermore, iPSCs are able to self-renew and thus can be expanded in nearly unlimited quantities, which make iPSC-derived cells an attractive alternative to primary cells for usage within in vitro screening models13.
Here, we describe a method to culture endothelial cells as perfusable 3D microvessels in a standardized, high-throughput microfluidic cell culture platform. Perfusion is applied by placing the device on a rocker platform, which ensures robust operation and increases the scalability of the assay. As the microvessels are continuously perfused and exposed to a gradient of angiogenic factors, angiogenic sprouting is studied in a more physiological relevant cellular microenvironment. While the protocol is compatible with many different sources of (primary) endothelial cells14,15, we focused on using human iPSC-derived ECs in order to increase the standardization of this assay and to facilitate its integration within vascular drug research.

<table>
	<tbody>
		<tr>
			<td>0.2-10 &#956;L Electronic repeater pipette</td>
			<td>Sigma</td>
			<td>Z654566-1EA</td>
			<td></td>
		</tr>
		<tr>
			<td>1 M HEPES</td>
			<td>Gibco</td>
			<td>15630-056</td>
			<td></td>
		</tr>
		<tr>
			<td>3-lane OrganoPlate</td>
			<td>Mimetas</td>
			<td>4003-400B</td>
			<td></td>
		</tr>
		<tr>
			<td>4% PFA in PBS</td>
			<td>Alfa Aesar</td>
			<td>J61899</td>
			<td></td>
		</tr>
		<tr>
			<td>Basal culture medium</td>
			<td>NCardia</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Collagen-1</td>
			<td>Cultrex</td>
			<td>3447-020-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Combitips Advanced Biopur 2.5 mL</td>
			<td>VWR</td>
			<td>613-2071</td>
			<td></td>
		</tr>
		<tr>
			<td>dPBS</td>
			<td>Gibco</td>
			<td>14190-094</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Repeater M4 pieptte</td>
			<td>VWR</td>
			<td>613-2890</td>
			<td></td>
		</tr>
		<tr>
			<td>Fibronectin</td>
			<td>Sigma-Aldrich</td>
			<td>F4759-1MG</td>
			<td>1 mg/mL in milliQ water</td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Gibco</td>
			<td>14025-050</td>
			<td></td>
		</tr>
		<tr>
			<td>Hoechst</td>
			<td>Molecular Probes</td>
			<td>H3569</td>
			<td>10 mg/mL solution in water</td>
		</tr>
		<tr>
			<td>iPSC-derived endothelial cells</td>
			<td>NCardia</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Sigma</td>
			<td>S5761</td>
			<td>37 g/L in milliQ water</td>
		</tr>
		<tr>
			<td>OrganoPlate Perfusion Rocker Mini</td>
			<td>Mimetas</td>
			<td></td>
			<td>Rocker platform used to provide flow</td>
		</tr>
		<tr>
			<td>Pen/Strep</td>
			<td>Sigma</td>
			<td>P4333</td>
			<td></td>
		</tr>
		<tr>
			<td>Phalloidin</td>
			<td>Sigma</td>
			<td>P1951</td>
			<td>1 mg/mL in DMSO</td>
		</tr>
		<tr>
			<td>PMA</td>
			<td>Focus-Biomolecules</td>
			<td>10-2165</td>
			<td>2 &#956;g/mL in PBS containing 0.1% DMSO</td>
		</tr>
		<tr>
			<td>S1P</td>
			<td>Ecehelon Biosciences</td>
			<td>S-2000</td>
			<td>1 mM in methanol containing 1% acetic acid</td>
		</tr>
		<tr>
			<td>TRITC-albumin</td>
			<td>Invitrogen</td>
			<td>A34786</td>
			<td></td>
		</tr>
		<tr>
			<td>VEGF</td>
			<td>Peprotech</td>
			<td>450-32-10</td>
			<td>50 &#956;g/mL in water containing 0.1% BSA</td>
		</tr>
	</tbody>
</table>

standardized and scalable assay to study perfused 3d angiogenic sprouting of ipsc-derived endothelial cells in vitro

Pre-clinical drug research of vascular diseases requires in vitro models of vasculature that are amendable to high-throughput screening. However, current in vitro screening models that have sufficient throughput only have limited physiological relevance, which hinders the translation of findings from in vitro to in vivo. On the other hand, microfluidic cell culture platforms have shown unparalleled physiological relevancy in vitro, but often lack the required throughput, scalability and standardization. We demonstrate a robust platform to study angiogenesis of endothelial cells derived from human induced pluripotent stem cells (iPSC-ECs) in a physiological relevant cellular microenvironment, including perfusion and gradients. The iPSC-ECs are cultured as 40 perfused 3D microvessels against a patterned collagen-1 scaffold. Upon the application of a gradient of angiogenic factors, important hallmarks of angiogenesis can be studied, including the differentiation into tip- and stalk cell and the formation of perfusable lumen. Perfusion with fluorescent tracer dyes enables the study of permeability during and after anastomosis of the angiogenic sprouts. In conclusion, this method shows the feasibility of iPSC-derived ECs in a standardized and scalable 3D angiogenic assay that combines physiological relevant culture conditions in a platform that has the required robustness and scalability to be integrated within the drug screening infrastructure.

The microfluidic 3D cell culture platform consists of 40 perfused microfluidic units (Figure 1a,b), which is used to study angiogenic sprouting of perfused microvessels against a patterned collagen-1 gel (Figure 1c). These microvessels are continuously perfused and exposed to a gradient of angiogenic growth factors (Figure 3a-d). The angiogenic sprouts can be either studied 2 days after gradient exposure or cultured for more than 5 days after gradient exposure to study anastomosis and sprout stabilization (see timeline, Figure 1d).
Seeding the iPSC-ECs using the passive pumping method should result in homogenous seeding densities (Figure 4a,b). Culture under continuous perfusion resulted in confluent microvessels in 2 days, with the cells completely lining the circumference of the microfluidic channel and the formation of a confluent monolayer against the patterned collagen-1 gel.
Exposure to a gradient of angiogenic factors resulted in directional angiogenic sprouting of the microvessels within the patterned collagen-1 gel (Figure 5a-g). Clear tip cell formation and invasion into the collagen-1 gel was visible 24 h after addition of the angiogenic gradient, while stalk cells including lumen formation were visible after 48 h (Figure 5a).
After fixation and staining, the capillary network can be visualized using phalloidin to stain F-actin and using Hoechst 33342 to stain the nucleus (Figure 5b,c). These sprouts can be quantified (e.g., shape and length14). Without addition of growth factors, no invasion into the collagen-1 gel should be observed (Figure 5d). Confocal imaging was used to determine the sprout diameter and to confirm lumen formation (Figure 5e-g).
The sprouts continue to grow towards the direction of the gradient and reach the opposite perfusion channel within 3&#8722;4 days after addition of angiogenic growth factors. This results in remodeling of the vascular network, with a clear reduction in the number of angiogenic sprouts (Figure 6a). Lumen formation was assessed by perfusion of the vascular network with fluorescently labeled macromolecules (e.g., albumin or dextrans). Perfusing the microvessels with 0.5 mg/mL labeled albumin before and after anastomosis revealed a clear difference in sprout permeability after 10 min (Figure 6b-e), which suggests that the capillaries stabilize and mature after anastomosis.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59678/59678fig1.jpg" /> 
Figure 1: Microfluidic cell culture protocol for iPSC-derived microvessels. (a) The bottom of the microfluidic cell culture device is shown displaying the 40 microfluidic units that are integrated underneath the 384-well plate. Larger view displays one of the 40 microfluidic units. (b) Each microfluidic unit is positioned underneath 9 wells with 3 inlet wells and 3 outlet wells. The microfluidic channels are separated by ridges (&#8216;phaseguides&#8217;), which enable the patterning of hydrogels in the central channel (&#8216;gel channel&#8217;) while there is still contact with the adjacent channels (&#8216;perfusion channels&#8217;). (c) Method to culture a perfused microvessel within the microfluidic device, which is used to study gradient driven angiogenic sprouting through a patterned collagen-1 matrix. (d) Timeline for studying angiogenic sprouting and/or anastomosis. This figure has been modified from van Duinen et al.14. <a href="https://www.jove.com/files/ftp_upload/59678/59678fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59678/59678fig2.jpg" /> 
Figure 2: Loading procedures for gel and medium. (a) Examples of correct and incorrect gel deposition. Correct filing results in a patterned collagen-1 gel in the middle channel, which is subsequently polymerized. (b) Examples of correct and incorrect filling of the wells. Wells are filled in the order of 1&#8722;4 to prevent air-bubble trapping within the microfluidic channels. <a href="https://www.jove.com/files/ftp_upload/59678/59678fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59678/59678fig3.jpg" /> 
Figure 3: Continuous hydrostatic pressure driven flow and gradient stabilization. (a) Hydrostatic pressure differences between wells result in passive levelling and flow within the microfluidic channels. (b) Placing the device on a rocker platform set at 7&#176; and 8 min cycle time results in continuous, bi-directional perfusion within the microfluidic channels. (c) Gradients are formed by introducing two different concentrations within the wells, which are continuously refreshed by passive leveling. (d) Gradient visualization using fluorescein isothiocyanate (FITC)-dextran. Bi-directional flow stabilizes the gradient up till 3 days. Scale bar = 200 &#181;m. This figure has been modified from van Duinen et al.14. <a href="https://www.jove.com/files/ftp_upload/59678/59678fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59678/59678fig4.jpg" /> 
Figure 4: Passive pumping method for cell seeding. (a) Passive pumping is driven by pressure differences that are caused by differences in surface tension. This results in a flow from the droplet (high internal pressure) towards the reservoir (low internal pressure). (b) Time lapse of a droplet (gray outline) that is placed on top of the inlet (white outline) of the microfluidic channel (blue outline). Right after addition (Figure 4b, i), the droplet on top of the inlet shrinks (Figure 4b, ii: 1 s after addition; iv: 2 s after addition), which results in a flow towards the outlet. This continues until the droplet meniscus is pinned by the inlet (Figure 4b, iv). Scale bar = 400 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59678/59678fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59678/59678fig5.jpg" /> 
Figure 5: Robust 3D sprouting of iPSC-EC microvessels. (a) Sprouting of iPSC-EC over time. Microvessels were grown for 48 h (right) and then stimulated with an angiogenic cocktail containing 50 ng/mL VEGF, 500 nM S1P, and 2 ng/mL PMA. The first tip-cells that invade the collagen-1 scaffold (middle) are visible 24 h after exposure. The first lumens are visible (arrows) 48 h after exposure (right) while the tip-cells have migrated further in the direction of the gradient. (b) Array of 15 microvessels that were stimulated with VEGF, S1P and PMA for 2 days and stained for F-actin (yellow) and nuclei (blue). Scale bar = 200 &#181;m. (c) Stimulated microvessel (positive control). (d) Unstimulated microvessel (negative control). (e) The maximum projection of a single capillary within the gel. (f) Same as (g) but focused on the middle. Dotted line indicates the position of the orthogonal view in panel g. Scale bars (a-d: 200 &#181;m; e-g: 20 &#181;m). <a href="https://www.jove.com/files/ftp_upload/59678/59678fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/59678/59678fig6.jpg" /> 
Figure 6: Visualization of angiogenic sprout permeability before and after anastomosis. (a) Anastomosis with basal channel triggers pruning and maturation of angiogenic sprouts. Closeup of capillary bed at 2, 4, 6 and 7 days after stimulation with angiogenic growth factors. (b) Angiogenic sprouts after 2 days after addition of angiogenic growth factors. Angiogenic sprouts are formed within gel, but are not yet connected to the bottom perfusion channel. (c) Perfusion of the microvessel with 0.5 mg/mL albumin-Alexa 555 solution. Fluorescent images obtained at 0 and 10 min. (d,e) Same as in panels b and c, but after 7 days of stimulation. Sprouts are connected to the other side and formed a confluent microvessel in the basal perfusion channel. Scale bars = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59678/59678fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Problem</td>
			<td>Cause</td>
			<td>Solution</td>
		</tr>
		<tr>
			<td rowspan="3">Collagen-1 does not enter or fill the channel completely</td>
			<td>Collagen-1 droplet is not placed on top of the inlet</td>
			<td>Carefully place the droplet on top of the inlet from the gel channel</td>
		</tr>
		<tr>
			<td>Volume of collagen-1 is too low</td>
			<td>Use 1.5 &#181;L of the gel to fill the channel completely</td>
		</tr>
		<tr>
			<td>Collagen-1 is too viscous</td>
			<td>Use another batch of collagen-1</td>
		</tr>
		<tr>
			<td>Collagen-1 flows into perfusion channels</td>
			<td>Collagen-1 is pipetted directly into the inlet of gel channel</td>
			<td>Carefully place the droplet on top of the inlet from the gel channel</td>
		</tr>
		<tr>
			<td rowspan="2">Collagen-1 is not clear/fiber formation</td>
			<td>Collagen-1 is not stored properly</td>
			<td>Store collagen-1 at 4 &#176;C, do not freeze</td>
		</tr>
		<tr>
			<td>NaHCO3 and HEPES are not mixed well before adding collagen-1</td>
			<td>Carefully mix the NaHCO3 and HEPES by pipetting before adding the collagen-1</td>
		</tr>
		<tr>
			<td rowspan="2">Droplet does not shrink using passive pumping method</td>
			<td>Droplet adheres to side of the well</td>
			<td>Aspirate droplet and add new droplet on top of the inlet</td>
		</tr>
		<tr>
			<td></td>
			<td>Make sure the outlet well is filled with at least 20 &#181;L of medium</td>
		</tr>
		<tr>
			<td rowspan="3">No sprouting is observed</td>
			<td>Growth factors are not added or aliquots are not stored properly</td>
			<td>Prepare fresh angiogenic sprouting medium</td>
		</tr>
		<tr>
			<td>Air bubble blocks perfusion/ gradient formation</td>
			<td>Remove air bubbles using a P20 or P200 pipette</td>
		</tr>
		<tr>
			<td>Volume differences between wells</td>
			<td>Volumes in all wells need to be equal in order to form a linear gradient</td>
		</tr>
		<tr>
			<td rowspan="2">Cells not viable</td>
			<td>Plate not placed on rocker platform/rocker platform turned off</td>
			<td>Make sure rocker platform is on and has the right cycle time/angle (8 min/7&#176;)</td>
		</tr>
		<tr>
			<td>No perfusion possible due to presence of air bubbles</td>
			<td>Remove air bubbles using a P20 or P200 pipette</td>
		</tr>
		<tr>
			<td>No lumens are formed, cells migrate as single cells</td>
			<td>Angiogenic sprouting mixture was added before a monolayer was formed</td>
			<td>Wait an additional 24 h before adding the angiogenic growth factors</td>
		</tr>
		<tr>
			<td>Major variation in sprouting density</td>
			<td>Differences in cell densities after seeding</td>
			<td>Check if cell density is homogenous and comparable between microfluidic units. Add another droplet of cell suspension if necessary</td>
		</tr>
	</tbody>
</table>
Table 1: Troubleshooting common errors.

Watch this Scientific Journal Video about Standardized and Scalable Assay to Study Perfused 3D Angiogenic Sprouting of iPSC-derived Endothelial Cells In Vitro at JoVE.com

Standardized and Scalable Assay to Study Perfused 3D Angiogenic Sprouting of iPSC-derived Endothelial Cells In Vitro

This method describes the culture of iPSC-derived endothelial cells as 40 perfused 3D microvessels in a standardized microfluidic platform. This platform enables the study of gradient-driven angiogenic sprouting in 3D, including anastomosis and stabilization of the angiogenic sprouts in a scalable and high-throughput manner.

Pre-clinical drug research of vascular diseases requires in vitro models of vasculature that are amendable to high-throughput screening. However, current ...

standardized-scalable-assay-to-study-perfused-3d-angiogenic-sprouting

Research

JoVE Journal

Bioengineering

30.3K Views.  Leiden University Medical Center. This method describes the culture of iPSC-derived endothelial cells as 40 perfused 3D microvessels in a standardized microfluidic platform. This platform enables the study of gradient-driven angiogenic sprouting in 3D, including anastomosis and stabilization of the angiogenic sprouts in a scalable and high-throughput manner.

Video: Standardized and Scalable Assay to Study Perfused 3D Angiogenic Sprouting of iPSC-derived Endothelial Cells In Vitro

Multiparametric Optical Mapping of the Langendorff-perfused Rabbit Heart

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not have been accomplished by other approaches1. With the use of the calcium- and voltage-sensitive dyes, optical mapping allows measurement of transmembrane action potentials and calcium transients with high spatial resolution without the physical contact with the tissue. This makes measurements of the cardiac electrical activity possible under many conditions where the use of electrodes is inconvenient or impossible1. For example, optical recordings provide accurate morphological changes of membrane potential during and immediately after stimulation and defibrillation, while conventional electrode techniques suffer from stimulus-induced artifacts during and after stimuli due to electrode polarization1. 
The Langendorff-perfused rabbit heart is one of the most studied models of human heart physiology and pathophysiology. Many types of arrhythmias observed clinically could be recapitulated in the rabbit heart model. It was shown that wave patterns in the rabbit heart during ventricular arrhythmias, determined by effective size of the heart and the wavelength of reentry, are very similar to that in the human heart2. It was also shown that critical aspects of excitation-contraction (EC) coupling in rabbit myocardium, such as the relative contribution of sarcoplasmic reticulum (SR), is very similar to human EC coupling3. Here we present the basic procedures of optical mapping experiments in Langendorff-perfused rabbit hearts, including the Langendorff perfusion system setup, the optical mapping systems setup, the isolation and cannulation of the heart, perfusion and dye-staining of the heart, excitation-contraction uncoupling, and collection of optical signals. These methods could be also applied to the heart from species other than rabbit with adjustments to flow rates, optics, solutions, etc.
Two optical mapping systems are described. The panoramic mapping system is used to map the entire epicardium of the rabbit heart4-7. This system provides a global view of the evolution of reentrant circuits during arrhythmogenesis and defibrillation, and has been used to study the mechanisms of arrhythmias and antiarrhythmia therapy8,9. The dual mapping system is used to map the action potential (AP) and calcium transient (CaT) simultaneously from the same field of view10-13. This approach has enhanced our understanding of the important role of calcium in the electrical alternans and the induction of arrhythmia14-16.

This article describes the basic procedures for conducting optical mapping experiments in the Langendorff-perfused rabbit heart using the panoramic imaging system, and the dual (voltage and calcium) imaging modality.

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not ...

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Eye Irritation Test (EIT) for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial (RhCE) Tissue Model

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and accurate assessment of ocular toxicity in man are needed. To address this need, we have developed an eye irritation test (EIT) which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model that is based on normal human cells. The EIT is able to separate ocular&#160;irritants and corrosives (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category). The test utilizes two separate protocols, one designed for liquid chemicals and a second, similar protocol for solid test articles. The EIT prediction model uses a single exposure period (30 min for liquids, 6 hr for solids) and a single tissue viability cut-off (60.0% as determined by the MTT assay). Based on the results for 83 chemicals (44 liquids and 39 solids) EIT achieved 95.5/68.2/ and 81.8% sensitivity/specificity and accuracy (SS&#38;A) for liquids, 100.0/68.4/ and 84.6% SS&#38;A for solids, and 97.6/68.3/ and 83.1% for overall SS&#38;A. The EIT will contribute significantly to classifying the ocular irritation potential of a wide range of liquid and solid chemicals without the use of animals to meet regulatory testing requirements. The EpiOcular EIT method was implemented in 2015 into the OECD Test Guidelines as TG 492.

Eye Irritation Test EIT for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial RhCE Tissue Model

We have developed an eye irritation test which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model. The test is able to discriminate between ocular irritant and corrosive materials (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category).

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and ...

Improved 3D Hydrogel Cultures of Primary Glial Cells for In Vitro Modelling of Neuroinflammation

In the central nervous system, numerous acute injuries and neurodegenerative disorders, as well as implanted devices or biomaterials engineered to enhance function result in the same outcome: excess inflammation leads to gliosis, cytotoxicity, and/or formation of a glial scar that collectively exacerbate injury or prevent healthy recovery. With the intent of creating a system to model glial scar formation and study inflammatory processes, we have generated a 3D cell scaffold capable of housing primary cultured glial cells: microglia that regulate the foreign body response and initiate the inflammatory event, astrocytes that respond to form a fibrous scar, and oligodendrocytes that are typically vulnerable to inflammatory injury. The present work provides a detailed step-by-step method for the fabrication, culture, and microscopic characterization of a hyaluronic acid-based 3D hydrogel scaffold with encapsulated rat brain-derived glial cells. Further, protocols for characterization of cell encapsulation and the hydrogel scaffold by confocal immunofluorescence and scanning electron microscopy are demonstrated, as well as the capacity to modify the scaffold with bioactive substrates, with incorporation of a commercial basal lamina mixture to improved cell integration.

Improved 3D Hydrogel Cultures of Primary Glial Cells for In Vitro Modelling of Neuroinflammation

Herein, we present a protocol for the 3D culture of rat brain-derived glia cells, including astrocytes, microglia, and oligodendrocytes. We demonstrate primary cell culture, methacrylated hyaluronic acid (HAMA) hydrogel synthesis, HAMAphoto-polymerization and cell encapsulation, and sample processing for confocal and scanning electron microscopic imaging.

In the central nervous system, numerous acute injuries and neurodegenerative disorders, as well as implanted devices or biomaterials engineered to enhance ...

Gene Expression Analysis of Endothelial Cells Exposed to Shear Stress Using Multiple Parallel-plate Flow Chambers

We describe a workflow for the analysis of gene expression from endothelial cells subject to a steady laminar flow using multiple monitored parallel-plate flow chambers. Endothelial cells form the inner cellular lining of blood vessels and are chronically exposed to the frictional force of blood flow called shear stress. Under physiological conditions, endothelial cells function in the presence of various shear stress conditions. Thus, the application of shear stress conditions in in vitro models can provide greater insight into endothelial responses in vivo. The parallel-plate flow chamber previously published by Lane et al.9 is adapted to study endothelial gene regulation in the presence and absence of steady (non-pulsatile) laminar flow. Key adaptations in the set-up for laminar flow as presented here include a large, dedicated environment to house concurrent flow circuits, the monitoring of flow rates in real-time, and the inclusion of an exogenous reference RNA for the normalization of quantitative real-time PCR data. To assess multiple treatments/conditions with the application of shear stress, multiple flow circuits and pumps are used simultaneously within the same heated and humidified incubator. The flow rate of each flow circuit is measured continuously in real-time to standardize shear stress conditions throughout the experiments. Because these experiments have multiple conditions, we also use an exogenous reference RNA that is spiked-in at the time of RNA extraction for the normalization of RNA extraction and first-strand cDNA synthesis efficiencies. These steps minimize the variability between samples. This strategy is employed in our pipeline for the gene expression analysis with shear stress experiments using the parallel-plate flow chamber, but parts of this strategy, such as the exogenous reference RNA spike-in, can easily and cost-effectively be used for other applications.

Here, a workflow for the culture and gene expression analysis of endothelial cells under fluid shear stress is presented. Included is a physical arrangement for simultaneously housing and monitoring multiple flow chambers in a controlled environment and the use of an exogenous reference RNA for quantitative PCR.

We describe a workflow for the analysis of gene expression from endothelial cells subject to a steady laminar flow using multiple monitored parallel-plate ...

Advanced 3D Liver Models for In vitro Genotoxicity Testing Following Long-Term Nanomaterial Exposure

Due to the rapid development and implementation of a diverse array of engineered nanomaterials (ENM), exposure to ENM is inevitable and the development of robust, predictive in vitro test systems is essential. Hepatic toxicology is key when considering ENM exposure, as the liver serves a vital role in metabolic homeostasis and detoxification as well as being a major site of ENM accumulation post exposure. Based upon this and the accepted understanding that 2D hepatocyte models do not accurately mimic the complexities of intricate multi-cellular interactions and metabolic activity observed in vivo, there is a greater focus on the development of physiologically relevant 3D liver models tailored for ENM hazard assessment purposes in vitro. In line with the principles of the 3Rs to replace, reduce and refine animal experimentation, a 3D HepG2 cell-line based liver model has been developed, which is a user friendly, cost effective system that can support both extended and repeated ENM exposure regimes (&#8804;14 days). These spheroid models (&#8805;500 &#181;m in diameter) retain their proliferative capacity (i.e., dividing cell models) allowing them to be coupled with the &#8216;gold standard&#8217; micronucleus assay to effectively assess genotoxicity in vitro. Their ability to report on a range of toxicological endpoints (e.g., liver function, (pro-)inflammatory response, cytotoxicity and genotoxicity) has been characterized using several ENMs across both acute (24 h) and long-term (120 h) exposure regimes. This 3D in vitro hepatic model has the capacity to be utilized for evaluating more realistic ENM exposures, thereby providing a future in vitro approach to better support ENM hazard assessment in a routine and easily accessible manner.

This procedure was established to be used for developing advanced 3D hepatic cultures in vitro, which can provide a more physiologically relevant assessment of the genotoxic hazards associated with nanomaterial exposures over both an acute or long-term, repeated dose regimes.

Due to the rapid development and implementation of a diverse array of engineered nanomaterials (ENM), exposure to ENM is inevitable and the development of ...

An Open Source Technology Platform to Manufacture Hydrogel-Based 3D Culture Models in an Automated and Standardized Fashion

Current mixing steps of viscous materials rely on repetitive and time-consuming tasks which are performed mainly manually in a low throughput mode. These issues represent drawbacks in workflows that can ultimately result in irreproducibility of research findings. Manual-based workflows are further limiting the advancement and widespread adoption of viscous materials, such as hydrogels used for biomedical applications. These challenges can be overcome by using automated workflows with standardized mixing processes to increase reproducibility. In this study, we present step-by-step instructions to use an open source protocol designer, to operate an open source workstation, and to identify reproducible mixtures. Specifically, the open source protocol designer guides the user through the experimental parameter selection and generates a ready-to-use protocol code to operate the workstation. This workstation is optimized for pipetting of viscous materials to enable automated and highly reliable handling by the integration of temperature docks for thermoresponsive materials, positive displacement pipettes for viscous materials, and an optional tip touch dock to remove excess material from the pipette tip. The validation and verification of mixtures are performed by a fast and inexpensive absorbance measurement of Orange G. This protocol presents results to obtain 80% (v/v) glycerol mixtures, a dilution series for gelatin methacryloyl (GelMA), and double network hydrogels of 5% (w/v) GelMA and 2% (w/v) alginate. A troubleshooting guide is included to support users with protocol adoption. The described workflow can be broadly applied to a number of viscous materials to generate user-defined concentrations in an automated fashion.

This protocol serves as a comprehensive tutorial for standardized and reproducible mixing of viscous materials with a novel open source automation technology. Detailed instructions are provided on the operation of a newly developed open source workstation, the usage of an open source protocol designer, and the validation and verification to identify reproducible mixtures.

Current mixing steps of viscous materials rely on repetitive and time-consuming tasks which are performed mainly manually in a low throughput mode. These ...

Fabrication of 3D Cardiac Microtissue Arrays using Human iPSC-Derived Cardiomyocytes, Cardiac Fibroblasts, and Endothelial Cells

Generation of human cardiomyocytes (CMs), cardiac fibroblasts (CFs), and endothelial cells (ECs) from induced pluripotent stem cells (iPSCs) has provided a unique opportunity to study the complex interplay among different cardiovascular cell types that drives tissue development and disease. In the area of cardiac tissue models, several sophisticated three-dimensional (3D) approaches use induced pluripotent stem cell-derived cardiomyocytes (iPSC-CMs) to mimic physiological relevance and native tissue environment with a combination of extracellular matrices and crosslinkers. However, these systems are complex to fabricate without microfabrication expertise and require several weeks to self-assemble. Most importantly, many of these systems lack vascular cells and cardiac fibroblasts that make up over 60% of the nonmyocytes in the human heart. Here we describe the derivation of all three cardiac cell types from iPSCs to fabricate cardiac microtissues. This facile replica molding technique allows cardiac microtissue culture in standard multi-well cell culture plates for several weeks. The platform allows user-defined control over microtissue sizes based on initial seeding density and requires less than 3 days for self-assembly to achieve observable cardiac microtissue contractions. Furthermore, the cardiac microtissues can be easily digested while maintaining high cell viability for single-cell interrogation with the use of flow cytometry and single-cell RNA sequencing (scRNA-seq). We envision that this in vitro model of cardiac microtissues will help accelerate validation studies in drug discovery and disease modeling.

Here, we describe an easy-to-use methodology to generate 3D self-assembled cardiac microtissue arrays composed of pre-differentiated human-induced pluripotent stem cell-derived cardiomyocytes, cardiac fibroblasts, and endothelial cells. This user-friendly and low cell requiring technique to generate cardiac microtissues can be implemented for disease modeling and early stages of drug development.

Generation of human cardiomyocytes (CMs), cardiac fibroblasts (CFs), and endothelial cells (ECs) from induced pluripotent stem cells (iPSCs) has provided ...

Stepwise Cell Seeding on Tessellated Scaffolds to Study Sprouting Blood Vessels

The cardiovascular system is a key player in human physiology, providing nourishment to most tissues in the body; vessels are present in different sizes, structures, phenotypes, and performance depending on each specific perfused tissue. The field of tissue engineering, which aims to repair or replace damaged or missing body tissues, relies on controlled angiogenesis to create a proper vascularization within the engineered tissues. Without a vascular system, thick engineered constructs cannot be sufficiently nourished, which may result in cell death, poor engraftment, and ultimately failure. Thus, understanding and controlling the behavior of engineered blood vessels is an outstanding challenge in the field. This work presents a high-throughput system that allows for the creation of organized and repeatable vessel networks for studying vessel behavior in a 3D scaffold environment. This two-step seeding protocol shows that vessels within the system react to the scaffold topography, presenting distinctive sprouting behaviors depending on the compartment geometry in which the vessels reside. The obtained results and understanding from this high throughput system can be applied in order to inform better 3D bioprinted scaffold construct designs, wherein fabrication of various 3D geometries cannot be rapidly assessed when using 3D printing as the basis for cellularized biological environments. Furthermore, the understanding from this high throughput system may be utilized for the improvement of rapid drug screening, the rapid development of co-cultures models, and the investigation of mechanical stimuli on blood vessel formation to deepen the knowledge of the vascular system.

Engineered tissues heavily rely on proper vascular networks to provide vital nutrients and gases and remove metabolic waste. In this work, a stepwise seeding protocol of endothelial cells and support cells creates highly organized vascular networks in a high-throughput platform for studying developing vessel behavior in a controlled 3D environment.

The cardiovascular system is a key player in human physiology, providing nourishment to most tissues in the body; vessels are present in different sizes, ...

A Microphysiological System to Study Leukocyte-Endothelial Cell Interaction during Inflammation

Leukocyte-endothelial cell interactions play an important role in inflammatory diseases such as sepsis. During inflammation, excessive migration of activated leukocytes across the vascular endothelium into key organs can lead to organ failure. A physiologically relevant biomimetic microfluidic assay (bMFA) has been developed and validated using several experimental and computational techniques, which can reproduce the entire leukocyte rolling/adhesion/migration cascade to study leukocyte-endothelial cell interactions. Microvascular networks obtained from in vivo images in rodents were digitized using a Geographic Information System (GIS) approach and microfabricated with polydimethylsiloxane (PDMS) on a microscope slide. To study the effect of shear rate and vascular topology on leukocyte-endothelial cell interactions, a Computational Fluid Dynamics (CFD) model was developed to generate a corresponding map of shear rates and velocities throughout the network. The bMFA enables the quantification of leukocyte-endothelial cells interactions, including rolling velocity, number of adhered leukocytes in response to different shear rates, number of migrated leukocytes, endothelial cell permeability, adhesion molecule expression and other important variables. Furthermore, by using human-related samples, such as human endothelial cells and leukocytes, bMFA provides a tool for rapid screening of potential therapeutics to increase their clinical translatability.

In this protocol, a biomimetic microfluid assay, which can reproduce a physiologically relevant microvascular environment and reproduce the entire leukocyte adhesion/migration cascade, is employed to study leukocyte-endothelial cell interactions in inflammatory disease.

Leukocyte-endothelial cell interactions play an important role in inflammatory diseases such as sepsis. During inflammation, excessive migration of ...