Simple Establishment of a Vascularized Osteogenic Bone Marrow Niche Using Pre-Cast Poly(ethylene Glycol) (PEG) Hydrogels in an Imaging Microplate

An in vitro model of the bone marrow vascular niche is established by seeding mesenchymal and endothelial cells onto pre-cast 3D PEG hydrogels. The endothelial networks, ECM components, and ALP activity of the niches vary depending on the growth factor used. The platform can be used for advanced cancer models.

Most in vitro avascularization studies use naturally derived matrices. While typically very conductive to biological processes, their inherent biological activity complicates the study of influences of single parameters.Here. we can selectively add our players of interest and assess how the system changes.

The sequential seeding of cells on our blank slate synthetic hydrogels enables the formation of very defined niches. Here, hBM-MSCs colonize the hydrogels and deposit their inherent extracellular matrix. This provides the substrate for the subsequently seeded endothelial cells.

This feature is not commonly found in other 3D in vitro models. The possibility to add different cell types sequentially allows for the creation of high-throughput compatible in vitro models with great complexity. These models can be adapted to generate more specific osteogenic vascularized niches and to investigate the functions of cellular and molecular players.

To begin, take a tube of hBM-MSCs and pellet them by centrifugation. Discard the supernatant and resuspend the pellet in an appropriate volume of basal medium. Then prepare 50 mL conical centrifuge tubes containing the required volume of basal medium, supplemented with the respective growth factors.

Finally, add the hBM-MSCs from the stock solution at a dilution of 1:66, 67 to achieve a concentration of 1.5 x 10^5 cells per mL. Remove the polypropylene adhesive film covering the 96-well hydrogel plate to seed the plate. Carefully aspirate the storage buffer covering the hydrogels by positioning the aspirator tip against the wall of the well, and slowly moving toward the edge of the inner well.

While seeding, mix the cell suspension periodically to keep the mixture homogenous and add 200 L of the cell suspension to each well of the plate. Then maintain the cultures at 37 C and 5%carbon dioxide in a humidified atmosphere. Alternatively, use an automated plate washer with the aspiration nozzles set at least 3.8 mm from the plate carrier and toward the edge of the well and aspirate the storage buffer.

Seed the plate by mixing the cells with a serological pipette and dispensing equal numbers of cells to the well of the plate. Monitor the development of the cultures by brightfield microscopy with a 5x objective and acquire reference images approximately 30 minutes after seeding to evaluate the number of cells added. To begin, take the pellet of the GFP-HUVECs grown to a confluence level of 80 100%and resuspend the cells in an appropriate volume of basal medium to achieve a concentration of 1 x 10^7 cells per mL of stock solution.

Prepare 50 mL conical centrifuge tubes containing the required volume of basal medium, supplemented with the respective growth factors. Add the GFP-HUVECs from the stock solution at a dilution of 1:66, 67 to achieve a concentration of 1.5 x 10^5 cells per L.Then aspirate the medium from the plate containing the stromal monoculture and add 200 L per well of the prepared GFP-HUVECs suspension. Incubate at 37 C in 5%carbon dioxide in a humidified atmosphere, changing the medium every three to four days.

Monitor the development of the culture by brightfield and fluorescence microscopy using a 5x objective. Differences between the cultures could be observed from the brightfield and fluorescence images showing only the GFP-HUVECs. The cultures appeared less developed without any growth factor.

However, faster development was observed in the presence of FGF-2. In contrast, the brightfield images showed the densest cultures in the presence of BMP-2. However, vascular-like networks formed in both growth factor containing conditions and the most extensive and interconnected networks were formed with FGF-2.

After establishing the stromal endothelial cell co-culture, acquire the GFP signal from the GFP-HUVECs using fluorescence microscopy with settings suitable for quantification. Pre-process all the images acquired on the same day to further enhance the contrast. If using Fiji or ImageJ, open all the enhanced GFP channel images at the same time point and open the brightness and contrast menu.

Select an image representing an intermediate condition and auto adjust the contrast by selecting auto. Click set and check propagate to all other open images. Visually assess whether the automatically selected range fits all the images of the current time point.

If needed, manually readjust and re-propagate the range to all the images and save the adjusted images as TIFF files. Next, apply a median blur filter to all the images. Reduce the size by binning and save them as grayscale RGB color TIFF files in a folder for quantification.

This could be done manually or in batch mode using macros. Analyze all the created images using the batch process mode in the Angiogenesis Analyzer for ImageJ. Then validate the quantification results by examining the overlays of the recognized structures and the original images.

Adjust pre-processing parameters, reanalyze original images, or exclude problematic areas if the algorithm detects artificial structures with few or no cells visible in the original image. Finally, normalize the obtained values to an area of 1 square mm by multiplying the values of each sample by the ratio of the analyzed area to 1 square mm. Quantified parameters of the GFP-HUVECs networks showed that the total network length was highest in the presence of FGF-2 and lowest in the absence of growth factors.

The number of junctions indicating branching points in the networks followed the same trend as the total length. Conversely, both growth factors featured significantly fewer isolated segments indicating higher interconnectivity than the condition without any growth factor. To begin, take the hBM-MSCs and GFP-HUVECs co-culture and wash once with 200 L per well of PBS.

Add the BCIP/NBT substrate solution and incubate the cultures at 37 C while periodically monitoring the color development. Once the color develops in non-osteogenic conditions, immediately wash with 200 L per well of PBS. After fixing the samples in 4%paraformaldehyde, acquire color images of the stained samples.

To quantify the ALP activity in cell lysates, add 200 L per well of 0.25%trypsin EDTA to the cultures washed with PBS and incubate at 37 C.Agitate the cultures every 10 minutes by vigorously pipetting up and down to facilitate digestion and monitor the culture morphology with a standard cell culture microscope. Then transfer the samples to 2 mL tubes and add 200 L of mesenchymal stem cell basal medium to inhibit the trypsin. After centrifuging and discarding the supernatant, freeze the pellets at 80 C or proceed directly to the next step.

Thaw the cell pellets at 500 L of lysis buffer and incubate for 30 minutes on ice. Next, centrifuge at 16, 100 G for 10 minutes at 4 C and keep the samples on ice. Without disturbing any pelleted debris, dispense 50 L of the supernatant into each well of a transparent tissue culture 96-well plate in duplicates.

Add 50 L of the prepared alkaline phosphatase reagent to the wells of the 96-well tissue culture plate. Shake the plate briefly and incubate it at 37 C for 10 minutes protected from light. Stop the reaction by adding 100 L of 1 molar sodium hydroxide using a multichannel pipette.

Using a plate reader, read the optical density at 410 nm. For DNA quantification, thaw the frozen samples, centrifuge them and place them on ice. Without disturbing any pelleted debris, dispense 50 L of the cell lysate supernatant into the wells of a black 96-well plate in duplicates.

Add the DNA standards in duplicates. Next, use a multi-channel pipette to add 50 L of the DNA staining agent, then incubate and read the fluorescence intensity according to the manufacturer's instructions. Utilize the standard curve values to determine and apply the conversion of the measured intensity values to the DNA concentrations.

Finally, normalize the alkaline phosphatase values by dividing them by the respective DNA concentration of each sample. Quantification of alkaline phosphatase activity to characterize the osteogenic potential of the cultures showed that the normalized alkaline phosphatase activity varied greatly across conditions, with a trend of increasing activity with higher concentrations of BMP-2 and a plateau at 100 ng per mL. The lowest activity levels were identified for the condition containing 50 ng per mL FGF-2.

More extensive and intense purple staining was observed in the presence of BMP-2 as compared to FGF-2.