iPS Cell Differentiation into Brain Microvascular Endothelial Cells

Abstract

The blood–brain barrier is a tissue structure that modulates the selective entry of molecules into the brain compartment. This barrier offers protection to the brain microenvironment from toxins or any fluctuations in the composition of the blood plasma via a layer of endothelial cells connected by tight junctions and supported by pericytes and astrocytes. Disruption of the barrier can be either a cause or a consequence of central nervous system pathogenesis. Therefore, research based on understanding the structure, function, and the mechanisms of breaching the blood–brain barrier is of primary interest for diverse disciplines including drug discovery, brain pathology, and infectious disease. The following protocol describes a detailed differentiation method that uses defined serum components during stem cell culture to deliver cellular cues in order to drive the cells towards brain endothelial cell lineage. This method can be used to obtain reproducible and scalable cultures of brain microvascular endothelial cells with barrier characteristics and functionality. These endothelial cells can also be stored long term or shipped frozen.

1. Introduction

The brain tissue is separated from the blood flow by the blood– brain barrier (BBB). The cells that compose this barrier strictly maintain and regulate the exchange of materials such as nutrients and toxins between the systemic blood circulation and the central nervous system (CNS) [1–4]. This structure acts primarily as a physical barrier due to the presence of complex tight junction (TJ) proteins in the brain endothelial cells that restrict the passage of most molecules across the BBB into the brain parenchyma [3, 5]. The main component of the BBB is the endothelial cell lining the blood capillaries in the brain, which are surrounded by other brain cell types such as neurons and glial cells. The organized interaction of these different cell types is referred to as the neurovascular unit (Fig. 1) [3, 6].

Fig. 1.

Schematic representation of cellular components forming the blood–brain barrier. Oxygen in the blood circulation is supplied to the brain by the internal carotid and vertebral arteries that, as they deepen into the brain tissue, branch into pial arteries, arterioles, and capillaries that run along and throughout the brain. The capillaries are the highest in density and responsible for most of the interactions between the brain parenchyma and the blood components

Researchers modeling the BBB have traditionally used animal models, cultured primary brain microvascular endothelial cells (BMECs), and a few cell lines derived from human BMECs (Table 1); however, all these have limitations, including species differences in the case of animal models and low barrier tightness due to the extended subculture of BMECs and immortalized BMECs cell lines [12, 13]. The accessibility of primary human BMECs also presents a challenge. Hence, there is a great need of in vitro models that are reliable and can fully replicate the complex relationships existing within the BBB neurovascular unit in vivo. Desired characteristics include both physiological functions and disease phenotypes such as barrier breakdown and the ensuing immune response [14, 15]. Here, we provide a detailed step-by-step protocol for a stem cell differentiation scheme with proven applications in drug permeability prediction [16], evaluation of nutrient uptake by the BBB [12], and barrier dysfunction in the context of neurological disorders [17]. The method consists of differentiation of pluripotent stem cells guided by serum and growth factor addition to culture medium and subsequent selection of BBB microvascular endothelial cells via capture by basement membrane proteins (Fig. 2). The endothelial cells can then either be directly used for experimental manipulations or frozen down using conventional cell culture techniques and shipped to other labs, facilitating experimental design and implementation of this model. This procedure allows for consistent production of cultures with desired barrier phenotypes such as high barrier tightness as measured by their transendothelial electrical resistance (TEER) values that average between 2000 and 4000 Ω.cm².

Table 1.

Reported TEER values for BBB culture in different models

	Reported TEER values (Ω.cm2)
In vivo
Rat BBB	30–5900 [7]
Transwell culture
iPSC monolayer co-culture with astrocytes	1450 [8]
iPSC monolayer co-culture with astrocytes	2000–8000[9]
Primary BMECs	70 [10]
hCMEC/D3	60 [10]
Dynamic BBB model
Primary BMECs	1200 [10]
hCMEC/D3	1200 [10]
b.End3	20 [11]
Microfluidic based model
b.End3	150 [11]
b.End3 co-culture with astrocytes	300 [11]

Fig. 2.

Differentiation time-course diagram. Stem cells are cultured on Matrigel-coated plates for 1–3 days, then medium is changed to growth medium for a minimum of 5 days until culture flasks are confluent and morphological changes are observed on the cell monolayer. After this, the cells are cultured for 2 days on EC medium with supplements to favor endothelial cell growth. Finally, the cells are selected for and subcultured using collagen IV and fibronectin coated culture vessels

2. Materials

2.1. Stem Cell Culture

1. Stem cells: hESCs (H9)11 or hiPSCs (iPS(IMR90)-4, iPSC DF19–9-11T.H).

2. Corning^® Matrigel^® Growth Factor Reduced (GFR) Basement Membrane Matrix.

3. T-75 cell culture flasks.

4. Stem cell culture medium (suggested: mTESR1, STEMCELL Technologies, 85857).

5. Dissociation solution (suggested: ReLeSR or Versene solution).

6. Astrocyte Medium.

7. Trypsin/EDTA Solution.

8. Dulbecco’s phosphate-buffered saline (DPBS).

9. Growth medium: DMEM/Ham’s F12, 20% Knockout Serum Replacer, 1× MEM nonessential amino acids, 1 Mm l-glutamine, 0.1 mM β-mercaptoethanol, 100 ng/mL Human basic fibroblast growth factor (bFGF), 10 μM Retinoic Acid (RA).

10. Endothelial cell (EC) medium: Human Endothelial Serum-Free Medium, 1× B27 supplement or 1× N2 supplement, 1× Antibiotic-Antimycotic: Streptomycin, Amphotericin B and Penicillin (PSA), 10 μM ROCK Inhibitor Y-27632.

11. StemPro^™ Accutase^™ Cell Dissociation Reagent.

12. Endothelial cell freezing medium: EC medium supplemented with RA and bFGF, 10% Fetal Bovine Serum (FBS), 10% Dimethyl Sulfoxide (DMSO). Sterile filter through a 0.2 μm filter unit if FBS or DMSO are shared reagents.

2.2. Plate Coating for Endothelial Cell Seeding

1. Corning^® Transwell^® polyester membrane cell culture inserts.

2. Coating solution: Mix in sterile distilled water the protein stocks to reach a final concentration of 400 μg/mL of collagen IV from human placenta and 100 μg/mL of fibronectin from human placenta. Add 250 μL of coating solution to each well of a 24-well plate or a 1.12 cm² transwell. Incubate overnight at 37 °C, 5% CO₂ for best results in cell attachment and growth.

2.3. TEER Measurements

1. EVOM2 voltohmmeter (World Precision Instruments).

2. STX2 electrode (World Precision Instruments).

2.4. Sodium Fluorescein Assay

1. Fluorescein salt solution: Dilute Fluorescein Sodium salt in fresh and sterile filtered EC medium to achieve a concentration of 100 ng/mL.

2. Spectrophotometer.

3. 96-well plates compatible with spectrophotometer plate reader.

2.5. Poly-D-Lysine Coating for T-75 Flasks

Coating solution: Use 10 mL of sterile water to dilute poly-D-lysine stock to a concentration of 0.1 mg/mL. For a T-75 flask use a coating volume of 10 mL and incubate for minimum of 1 h or preferably overnight at 37 °C, 5% CO₂.

2.6. Fixing and Permeabilization of Cells

1. Fixing solution: 4% paraformaldehyde.

2. PBT buffer for permeabilization of cells: Phosphate-Buffered Saline (PBS) with 0.1% Tween 20.

2.7. PBTG Blocking Solution

1. PBTG: Add 5% Normal Goat Serum to PBT solution and sterile filter through a 0.2 μm filter unit. For long-term storage, place in 4 °C fridge.

3. Methods

3.1. General Recommendations to Prevent Contamination During the Differentiation Process

1. During the process of stem cell culture and differentiation, it is crucial to maintain the cells free of bacterial and fungal contamination. Use strict aseptic technique in the cell culture hood. Routinely screen stem cell stocks for hard to detect bacteria such as Mycoplasma, as contamination with this micro-organism can lead to decreased cell proliferation.

2. Before and after working in the sterile hood, use 70% ethanol to decontaminate reagents containers, gloves, and the general work surface. The work area should contain only the reagents required for the specific procedure that is about to be performed.

3. Date and label the reagents pertinent for cell culture and differentiation procedures to track and ensure sterility. Avoid leaving reagents uncapped and unattended. Sterile filter media stocks and divide in ready to use aliquots using sterile pipettes.

3.2. Maintenance of Stem Cell Culture

3.2.1. Preparation of Matrigel-Coated Flasks

1. Thaw the aliquots of Matrigel in cold (4 °C) DMEM/Ham’s F12 to achieve a concentration of 0.2 mg/mL of Matrigel. For a 24-well plate, use a volume of 200–400 μL to coat the wells.

2. Coat flasks or plates the day before using. Incubate at 37 °C, 5% CO₂ overnight (see Notes 1–2).

3.2.2. Thawing Stem Cell Stocks

1. Before thawing the cells, prepare the flasks that will be used for culture. Retrieve the flasks from the incubator and aspirate the medium used for coating the flask. If using a T-75 flask, add 5 mL of mTESR1 medium and set aside with appropriate labels.

2. Prepare 15 mL falcon tubes to centrifuge the stem cell stocks by adding 5 mL of mTESR1 medium into each falcon tube. Remove cryovial from liquid nitrogen and use 70% ethanol to clean the outside of the cryovial, then thaw the cells by gently swirling the frozen cryovial in the 37 °C water bath until a small crystal remains inside.

3. Dry excess water and use 70% ethanol to sterilize the outside of the cryovial, especially around the cap, before placing it inside the cell culture hood. Briefly let the cryovial air dry for 20 s, then transfer the contents to the pre-prepared 15 mL falcon tube with 5 mL of culture medium. Centrifuge for 5 min at 200 × g. Aspirate medium and resuspend the cells gently in 12 mL mTESR1 supplemented with 10 μM ROCK inhibitor (see Notes 3–6).

4. After seeding the cells on the flask, incubate overnight at 37 °C, 5% CO₂ without disturbing the cells in the incubator for at least 10 h. Allow the cells to get to 70–80% confluency before splitting to seed into flasks to start the differentiation process.

3.2.3. Maintenance and Subculture of Stem Cells

1. During stem cell culture, perform daily media changes adding 12 mL of fresh mTESR1 to the flask. After the flasks have reached the desired confluency (between 70% and 80%) to passage the cells, aspirate the culture medium and add 3–5 mL of ReLeSR to the T-75 flask, incubate for 5 min at 37 °C.

2. After the incubation period, resuspend the cells with mTESR1 without ROCK inhibitor to obtain a final volume of 6 mL. Count the cells to seed approximately 1 × 10⁶ to 1.5 × 10⁶ cells in a new T-75 flask with a final volume of 12 mL of mTESR1.

3. Incubate overnight at 37 °C, 5% CO₂ and check for attachment and cell viability the next day before starting the differentiation process.

3.3. Differentiation Protocol

3.3.1. Differentiation Protocol Setup

1. The day after seeding the appropriate number of stem cells and culturing for 1–2 days in mTeSR1, switch the stem cell culture medium to growth medium containing knockout serum replacer to initiate differentiation [8] (see Notes 7–8). Cells will proliferate quickly and go through morphological changes from a stem cell state to a mixed culture of endothelial and neural cells by day 4 of medium exposure. Replace medium daily with fresh growth medium during this phase.

2. After day 5 (Fig. 2), switch the growth medium to EC medium supplemented with 20 ng/mL bFGF and 10 μM RA [9].

3. After 48 h of incubation at 37 °C, dissociate the cell monolayer with Accutase (see Note 9). Use an appropriate volume to cover the cell monolayer and incubate for 30 min to 1 h at 37 °C. Use 5 mL of Accutase for a T-75 flask.

4. Resuspend the dissociated monolayer by dispensing fresh EC medium lacking RA and bFGF onto the flask. Use a cell scraper to gently resuspend sections that are hard to reach with the serological pipettes if culturing in flasks. Collect in 15 mL falcon tubes and centrifuge the cells for 5 min at 200 × g. Aspirate the medium and resuspend the cells in EC medium without RA and bFGF but supplemented with 10 μM of ROCK inhibitor (see Note 10).

5. Use Trypan blue to evaluate viability after centrifugation and resuspension, use the live cell count for seeding calculations.

6. Plate cells onto 24-well tissue culture plates or 1.12 cm² Transwell-Clear permeable inserts (0.4–8 μm pore size) coated with 250 μL of a 4:1 mixture of human collagen IV and fibronectin. Seed between 5 × 10⁵–8 × 10⁵ cells/well. Incubate the seeded cells overnight and change the medium (without RA and bFGF) the next day.

7. The next day after seeding onto collagen IV and fibronectin coated plates, measure the TEER values on the Transwell plates using a STX2 chopstick electrode and an EVOM2 voltohm-meter. TEER measurements can be monitored in a timely manner relevant to the experiment but for maintenance culture purposes, measure every 24 h.

3.4. Freezing Down Differentiated Endothelial Cells

1. Freeze the endothelial cell stocks before the final selection on collagen IV and fibronectin coated culture vessels. After 30 min–1-h incubation with Accutase on day 8 (Fig. 2), collect and centrifuge the cells for 5 min at 200 × g.

2. Aspirate medium and resuspend the cells with the endothelial cell freezing medium. Make stocks of 8 × 10⁶ cells/mL making sure to break down the cell aggregates into small or single cell masses.

3. Aliquot in prelabeled cryovials and store at −80 °C overnight inside an isopropanol freezer container to allow the cells to freeze slowly. The next day, for extended storage, store cell vials in liquid nitrogen. Frozen cell stocks can be used within a month from the date of storage and yield a barrier with similar characteristics as non-frozen endothelial cell monolayers.

3.5. Co-culture of Stem Cell Derived Brain Microvascular Endothelial Cells with Primary Astrocytes

1. To set up the co-culture during the experiments, seed primary human astrocytes on poly-D-lysine coated plates at a final concentration of 2 μg/cm².

2. A day after the establishment of the astrocyte culture from cryopreserved cell stocks, replace medium with fresh astrocyte complete medium. For regular maintenance, change medium every 3 days and monitor to passage the cells when the culture reaches 80–90% confluency.

3. To passage and seed the astrocyte cells for the co-culture experiments, aspirate culture medium, wash cell monolayer using 5 mL of DPBS, and incubate the cells in 1 mL of 0.05% trypsin/EDTA solution for 5 min at 37 °C. After incubation, confirm under a microscope the change in cell morphology and detachment from the culture vessel.

4. Gently tap the side of the flask to help release the cells from the culture surface and transfer the trypsin and cell solution into a pre-prepared 15 mL falcon tubes containing 5 mL astrocyte complete medium. Rinse the flask with an additional 5 mL of astrocyte complete medium to collect any remaining cells. Centrifuge the falcon tubes containing the cells for 5 min at 200 × g.

5. After centrifugation resuspend the cell pellet in astrocyte complete medium for plating into new coated culture plates. For maintenance culture, seed approximately 5000 cells/cm². When setting up the co-culture transwell experiments, seed 20,000 cells per well in a 12-well plate. When the astrocyte culture has reached 70% confluency in the wells, start the co-culture by transferring transwell inserts already seeded with endothelial cells into the astrocyte plate, one insert per well.

3.6. Immunofluorescence Staining of Brain Microvascular Endothelial Cells Grown on Transwell Membrane

1. To perform immunostaining of the endothelial cells grown on the transwell membrane inserts, fix cells for 15 min with appropriate volume of 4% Paraformaldehyde (PFA) at room temperature. For a 1.12cm² transwell size use 200–250 μL of the fixing solution. After the incubation period, wash once with 1× PBS, aspirate, add 200 μL 1× PBT, and incubate for another 5 min at room temperature.

2. After incubation, aspirate 1× PBT and wash once with 1× PBS, aspirate, add 200 μL of PBTG per well, and incubate for 1 h at room temperature.

3. Make primary antibody solution diluting primary antibodies in PBTG according to the concentration recommended by the supplier. After incubation period, aspirate PBTG and add 200 μL of primary antibody solution per well. Incubate the primary antibody 1 h at room temperature or overnight at 4 °C. After incubation, wash 3 times with 1× PBS for 5 min at room temperature.

4. Aspirate 1× PBS and add 200 μL of secondary antibody solution (secondary antibodies diluted in PBTG according to the supplier instructions) and incubate at room temperature for 1 h. Aspirate and wash 3 times with 1× PBS.

5. After the final wash, aspirate 1× PBS and cut the transwell membrane carefully using a scalpel or razor blade. Place scalpel on the border of the membrane from the basolateral side of the transwell and turn the transwell to release the membrane. Use tweezers to hold the membrane to avoid damaging the cell monolayer, dab excess 1× PBS on the membrane using a paper towel and mount the cells on coverslips then place on microscope slides. Make sure to mount the membrane cell-size down onto the coverslip with mounting medium. After placing the coverslips on the microscope slides, seal the edges with clear nail polish to preserve the slide.

3.7. Measurement of TEER for Barrier Tightness Assessment

1. Take TEER measurements using STX2 electrodes attached to an EVOM voltohmmeter. Place the STX2 electrodes within the transwell and the containing well (see Note 11, Fig. 3A).

2. Measure the resistance value in each transwell 5 times, including a cell-free transwell to use as control for membrane resistance.

3. Adjust the average electrical resistance value (Ω) with the surface growth area (cm²) of the transwell to obtain the barrier TEER (Ω .cm²) value of each transwell.

Fig. 3.

Barrier integrity evaluation methods. (a) Transendothelial electrical resistance measurement of the endothelial cell monolayer with chopstick electrodes. Electrode placement between the transwell containing endothelial cells and plate well containing astrocytes. The electrical resistance is measured in Ohms (Ω) and adjusted to the growth surface area of the transwell (cm²). (b) Sodium fluorescein assay schematics. Permeability to sodium fluorescein is measured based on the salt concentration that diffuses through the monolayer of endothelial cells into the basolateral side of the transwell chamber over a time

3.8. Sodium Fluorescein Assay

1. Aspirate medium from the apical chamber of the transwell plate and replace it with sodium fluorescein (Na-F) in fresh EC medium (Fig. 3b). Every 30 min, during the following 2-h window, remove 500 μL of medium from the basolateral chamber of each transwell. Immediately, replace the volume on the basolateral chamber with 500 μL of fresh medium. In a 96-well plate, transfer 100 μL of the previously removed 500 μL medium to measure the concentration of Na- F flowing through the membrane.

2. Use a cell-free transwell coated with 4:1 collagen IV and fibronectin solution to control for the effect of the transwell membrane. Measure the fluorescence of the collected samples in a microplate reader compatible with your spectrophotometer using Ex(λ) 485 ± 10 nm and Em(λ) 530 ± 12.5 nm [18, 19]. Perform this assay with technical triplicates for the fluorescent label reading.

3. Calculate the permeability coefficient (Pe) value by obtaining the clearance of Na-F flowing from the apical to the basolateral side of the chamber. Calculate clearance (μL) from the initial 100 ng/mL sodium fluorescein (Na-F) diluted in fresh EC medium added to the apical side of the chamber and the final concentration of Na-F in the basolateral side as: Clearance (μL) = C_A × V_A/C_i. In this case, C_A is the basolateral Na-F concentration, V_A is the volume of the basolateral chamber, and C_i is the initial Na-F concentration used (100 ng/mL).

4. Plot the average clearance vs time for each timepoint (every 30-min measurement of the removed 500 μL from the baso-lateral side of the transwell set up). Use linear regression analysis to obtain the slope of the clearance curve. The slope (P) of the clearance curve for the monolayer is the permeability by the surface area of the transwell (μL/min). Denote the permeability of the control cell-free transwell membrane as Pc and the permeability for the experimental transwell culture as Pt. Then calculate the value for Pt using the equation: 1/P = 1/Pc + 1/ Pt. Divide Pt by the surface area of the transwell to obtain Pe of each transwell in cm/min [18, 20].

4. Notes

1. It is recommended to coat the Matrigel plates prior to the day of seeding the stem cells. If planning to thaw and monitor multiple batches of stem cells in the same week, it is possible to coat multiple flasks with Matrigel. Be sure to use these flasks within 5–7 days after coating to avoid the DMEM/Ham’s F12 medium from evaporating. Monitor humidity inside the incubator to prevent the flasks or Matrigel-coated plates from drying out faster than expected. If there are sections of evaporated medium on the Matrigel plates, do not use them as the dried patches compromise the coating quality and limits the attachment of the stem cells in the culture vessels.

2. To aliquot Matrigel stocks and to dissolve into DMEM/Ham’s F12 for coating flasks, it is recommended to use chilled equipment (for example, tips and medium) to slowly dissolve the Matrigel into the medium and immediately apply the coating solution to the culture vessel. Avoid applying body heat from handler’s fingertips directly on the Matrigel aliquot by holding the centrifuge tube away from the frozen Matrigel pellet.

3. When thawing the stem cells, do not release the contents of the cryovial against the wall of the falcon tubes, aim to slowly release the cells directly into the 5 mL medium to avoid extra shear stress applied to the cells.

4. To resuspend the stem cells, Accutase can also be used; however, it tends to break the stem cell colonies into small colonies or even single cell cultures. This can cause spontaneous differentiation or lower the attachment efficiency of the stem cells. We recommend using ReLeSR instead, as incubation period is short (~5 min) and the culture is maintained in small to medium colonies that attach well when re-seeded in new Matrigel-coated flasks, maintaining a high yield of differentiated endothelial cells.

5. It is recommended to resuspend stem cells so that they form small to medium sized colonies when plated. To achieve this, resuspend the cells in mTESR1 medium with a 5 mL pipette 2–4 times. Do not force the cells against the wall of the falcon tube or flask as the excess shear force will reduce attachment and viability.

6. Ideally, keep stem cell cultures that will be used to differentiate into endothelial cells under passage 40. Thaw and culture the stem cells until the cells reach splitting confluency (70–80% confluency on a T-75 flask). Split stem cells in a 1:6 ratio into new T-75 flasks to start the differentiation process. Check for spontaneous differentiation in the stem cell culture: if the flask contains over 20% differentiated colonies, it is not recommended to use the flask for differentiation.

7. Change growth medium every day and increase medium volume from 12 to 15 mL during the growth phase as the cell number increases and the flask gets confluent (day 4 and 5). It is recommended to prepare fresh batches of medium stocks that will be used within 1 week and keep them stored at 4 °C.

8. During day 6–8 it is recommended not to change the endothelial cell culture medium; however, cell numbers might be high, and nutrients could be used up quickly during this 48 h incubation period. To avoid nutrient starvation and drastic low pH in the culture, increase culture medium volume to 20 mL and monitor the pH at 24 h. If the medium pH reaches 6.5–6.7, it is preferable to change medium to freshly made endothelial cell medium than inducing further stress in the culture.

9. After day 8 of differentiation, to resuspend the cells for seeding on coated Transwells, plates, or slides, incubate at 37 °C for 30 min to 1 h with Accutase. Do not let the incubation period go longer than 1 h as it will decrease cell viability. After 30 min of incubation, monitor the cell monolayer looking for signs of lifted cells in the culture flask. Rock the flasks back and forth to avoid drying out the cells while the Accutase is incubating. After the cells have lifted, use 5 mL of endothelial cell medium to resuspend the cells and break up into small clumps. A cell scraper could be used to lift any remaining cells and help collect all of the differentiated cells.

10. After centrifugation of the differentiated cells at day 8, it is recommended to break the pellet into single cells by pipetting up and down several times in small volume using a p1000 micropipette. This is recommended for either seeding or freezing down the endothelial cell stocks.

11. When measuring the TEER values, place the electrodes straight over the transwell culture (Fig. 3), avoid placing the electrodes with inclination or touching the monolayer on the apical side of the transwell so the monolayer is not disturbed.