Improved culture methods for patient-derived tumor samples are critically important as the field moves away from the use of conventional cell lines. This protocol allows for the in vitro culture of patient-derived tumor cells in a platform that is amenable to high throughput and high content screening, unlike in vivo cancer models. The platform incorporating PDXs allows us to assess more heterogeneous systems reflective of native tumors.
This may yield insight into drug mechanisms and allow for faster drug screening. The method could be expanded through the incorporation of stroma, endothelium, and immune cells. These co-cultures could be even more reflective of the native tumor population and architecture.
Practice loading the microfluidic plates with easily-expanded cell lines to become highly proficient before shifting to more precious cells like PDXs. Alignment during plate dispensing greatly impacts success, and a visual demonstration can easily convey this concept. To begin, transfer tumor tissue to a pre-weighed sterile 50-milliliter conical tube.
Rinse six times with 30 milliliters of sterile PBS to remove blood and contaminants. Remove as much liquid as possible, and weigh the tumor tissue. Transfer the tumor tissue to a 60-millimeter round tissue culture dish, and use a sterile razor blade or a scalpel to mince it into one-millimeter pieces.
Add five milliliters of PDX processing medium to collect the tumor slurry. Add five milliliters of dissociation enzyme solution to collect the tumor slurry. Rinse the culture dish with another five milliliters of dissociation enzyme solution.
Then add five milliliters of PDX processing medium. Incubate 20 minutes at 37 degrees Celsius with gentle shaking. Halfway through the incubation time, swirl the tube gently.
After incubation, pipette up and down gently with a serological pipette to break up clumps. Place a 70-micron cell strainer over a new sterile 50-milliliter tube, and filter the cells. Then centrifuge at 1, 200 rpm for two minutes to pellet the cells.
Remove the supernatant and resuspend in two to three milliliters of PDX culture medium. Count the cells using a hemocytometer or automated cell counter. Use this table to estimate the required number of the associated PDX derived cells needed to achieve the desired cell density per chip.
Plate one to two times 10 to the six cells in five milliliters of PDX culture medium per well of a six-well tissue culture plate. Incubate for 48 hours at 37 degrees Celsius, 5%carbon dioxide, and 95%humidity with gentle shaking at 50 to 55 rpm to promote cluster formation. After the clusters have formed, proceed to centrifugation.
First prepare 20 milliliters of 100%density gradient solution by thoroughly mixing 18 milliliters of density gradient centrifugation solution with two milliliters of sterile 10X HBSS in a sterile 50-milliliter conical tube. Dilute this 100%solution with sterile 1X HBSS to make 10 milliliters each of 20%30%40%and 55%density gradient solutions. Add three milliliters of 55%density gradient solution to the bottom of a 15-milliliter conical tube.
Holding the tube at an angle, slowly dispense three milliliters of 40%density gradient solution onto the angled side of the tube and on top of the 55%layer. Repeat with the 30%density gradient solution. Then collect the supernatant of PDX rotation cultures with a five-milliliter serological pipette into a tube, rinsing the plate surface gently.
Centrifuge at 1, 200 rpm for two minutes to pellet cells. Remove the supernatant, and resuspend the cell pellet in three milliliters of 20%density gradient solution. Carefully layer the 20%density gradient solution with cells onto the top of the gradient in the 15-milliliter tube.
Cap the tubes and centrifuge in a swing-bucket rotor centrifuge 30 minutes at four degrees Celsius, 2, 000 times G, and zero break. After centrifugation, fractions are visible. Viable PDX cell cultures are typically found at the 40 to 55%density gradient solution interface.
Collect two to three milliliters of each fraction into fresh 15-milliliter tubes. Add three to four volumes of sterile 1X HBSS to each fraction, and invert to mix thoroughly. Centrifuge at 1, 000 times G for three minutes to pellet the cells.
Remove the supernatant. Resuspend the cell pellet in one to two milliliters of PDX processing medium. Transfer a small aliquot between 50 to 100 microliters into a tube, and add an equal volume of dissociation enzyme solution for redissociation.
Assess the cell number in the clustered cell suspension. Count the cells with a hemocytometer or automated cell counter. Reconstitute hyaluronic acid hydrogel solutions according to the manufacturer's instructions.
Using a multi-channel pipette, add 50 microliters of HBSS to all wells in observation window columns of a two-lane microfluidic plate to maintain culture humidity and optimal imaging conditions. Calculate the volume of cell suspension needed for 50 microliters of hydrogel at the desired cell density, for example, 5, 000 cells per microliter. For seeding one microfluidic plate, aliquot the calculated volume into each of four sterile 1.5-milliliter centrifuge tubes.
Adjust the pH of the HA thiol solution to eight with 1 normal sodium hydroxide immediately prior to use. Perform a test gelation by mixing 40 microliters of HA thiol with 10 microliters of PEGdA and monitoring gelation over time. Next, centrifuge the cell suspension aliquots for two minutes at 200 times G and room temperature to pellet the cells.
Carefully remove the supernatant, and resuspend the cells in 40 microliters of HA thiol for a 50-microliter final volume. Add 10 microliters of PEGdA to one aliquot of the cells of HA thiol. Mix well, and wait one to three minutes before seeding the microfluidic plate.
Affix a tip for dispensing 1.5 microliters to a single-channel repeating pipette, and load with the cells in HA hydrogel solution. Remember to keep the hydrogen aliquot well-mixed to ensure even cell distribution. To seed the microfluidic plate, align the pipette tip perpendicular to the blade while gently placing the tip in the center of the gel inlet to ensure contact but no pressure when dispensing 1.5 microliters of hydrogel solution.
Observe the fill status of the microfluidic channels by viewing from the top of the plate, bottom of the plate, or by microscope. Assess the loading using this figure as a guide. One minute after the loading, invert the plate while preparing for the next aliquot.
Repeat the seeding of the microfluidic plate for the remaining three aliquots of cells in HA solution. After all chips are filled, incubate the plate at 37 degrees Celsius in a humidified incubator 45 minutes until gelation is complete. After that, using the manual provided, ensure the perfusion rocker is installed in the cell culture incubator with the correct profusion setting at 14-degree angle and four-minute intervals.
Add 50 microliters of PDX cell culture medium to all medium inlets. Gently tap the plate against a surface to encourage the liquid to fill the microfluidic channels. Then flip the plate to check if the channels are filled properly.
Add 50 microliters of DMEM/F-12 for all media outlets. If any air bubbles are trapped in the perfusion channel, remove by gentle tapping of the plate against a surface. Using a microscope and plate layout form, record chip filling success.
Exclude improperly filled chips from further experimental use. Place the plate on a tilting rocker set to a 14-degree tilt and a four-minute cycle to begin perfusion. Every two days, replace the PDX culture medium, first 50 microliters in the inlets, then 50 microliters in the outlets.
In this study, a programmable perfusion rocker was installed in a standard water-jacketed cell culture incubator, and two-lane microfluidic blades were prepared in a standard biosafety cabinet for loading. 3D microfluidic PDX culture viability and morphology were evaluated in both unseparated and density-gradient centrifugation-separated conditions. On day one, those cultures which underwent the separation method exhibited tenfold fewer single dead cells compared to unseparated cultures.
Importantly, the separated clusters primarily consisted of live cells. No statistically significant difference was identified for the cluster size distribution. Cultures were further maintained in the microfluidic plate for seven days.
The total number of live cells remained consistent, and clusters retained approximately 80%viability over the life of the culture. Remember that each PDX line is unique, so the ease of tumor digestion, the phenotypic size and morphology of clusters, and the cluster's location within the gradient purification will vary. PDX cultures established using this method can be assayed with image-or plate-reader-based viability assays, fixed and immunofluorescently labeled, or dissociated to collect cell lysates for other protocols.
The method will allow researchers to recapitulate the tumor microenvironment in order to explore biology or drug response of tumor cells. Users should complete standard human subjects training and bloodborne pathogens training prior to working with human-derived tissues. Appropriate personal protective equipment should be worn.
