Construction of the plasmids using Gibson assembly

We used NEBuilder® Hifi DNA Assembly Master Mix for Gibson Assembly to insert two fragments into our plasmid backbone, which was digested with the restriction enzymes MluI and BbsI before assembly. For the Gibson Assembly to be successful, we added 25-base pair (bp) overhangs to both sides of each fragment. These overhangs were identical to the DNA regions where the assembly would occur, allowing the 5’ exonuclease to create single-stranded overhangs that were complementary, hereby facilitating the seamless assembly of our fragments into the plasmid backbone.

Transformation into competent cells using heat-shock

We used heat-shock to make competent E.coli DH5α able to take in our plasmids assembled by Gibson Assembly. The heat-shock makes the E.coli want to adapt to the sudden change in temperature and therefore more willing to take in our plasmids. After the heat-shock, the cells are plated onto selection plates with 50 µg/mL ampicillin.

Colony PCR

We used colony PCR to verify that the plasmids were correctly assembled. This was done by selecting some of the colonies from the selection plates and run a PCR with primers that bind to the flanking sequences of the insertion segment. If the Gibson Assembly was successful, a band with the same length as the insertion piece would be revealed on a gel. In our case the length of the insertion piece for the Control Plasmid was 3720 bp and the length for the Treatment Plasmid was 3627 bp.

Western blot

We used western blot to see if our antibodies bind specifically to the ERα and ERβ receptors. The process began by extracting proteins from our two cell lines, which were then denatured to ensure they unfold. These proteins were separated by size using SDS-PAGE, a form of gel electrophoresis. After separation, the proteins were transferred onto a membrane. To prevent nonspecific binding, the membrane is blocked with a protein solution. Next, the membrane was incubated with one of our antibodies that specifically binds to the target protein following a second antibody that binds to the other target. The antibodies were conjugated with a fluorescent dye allowing the visualization of the target proteins.

Media Change

The media was changed to ensure the cells had the proper nutrients needed for continued growth. The old media was removed from the cells, and new media was prewarmed at 37°C before it was added to the cells.

Subculturing

Cells were subcultured to provide them with enough space to grow. This was done when the confluency was around 70-90%. The old media was removed, and the cells were washed with prewarmed PBS. The cells were then treated with using 0.25% Trypsin to detach the cells from the flask. New media was added to the cells, which were then transferred to a Falcon tube. The cells were centrifuged at 120xg for 5 min, and the supernatant was discarded after which the cells were resuspended in culture media and split into a suitable ratio (1:3, 1:6, or 1:10).

Electroporation

Cells were harvested and resuspended in PBS and transferred to a chilled cuvette. The plasmid was added to a final concentration of 5 μM. The electroporator's parameters (voltage, pulse length, and number of pulses) were optimized for the specific cell type. The pulse button was pressed to activate the automatic charge and pulse sequence. Following electroporation, the cuvette was incubated for 2 minutes on ice, followed by 8 minutes at 37°C. After incubation, the cell suspension was carefully removed from the cuvette and transferred to 5 mL of culture medium in a T25 flask for cultivation.

Transient transfection

Cells were harvested at an optimal density (70-90% confluence), and the desired number of cells was transferred to an Eppendorf tube. The transfection reagent was mixed with culture media and then added to the cells, after which the mixture was incubated for approximately 24 hours at 37°C. After incubation, the transfection mixture was removed, and fresh media was added to the cells. After a recovery time of 24 to 48 hours, transfection efficiency was assessed by measuring fluorescence using flow cytometry.

Filtration and fixation (prep for flow cytometry)

To obtain optimal results from flow cytometry analysis, the cells needed to be in a single-cell suspension, which required filtration. This was done by cutting the end of a P1000 pipette tip, then placing a small piece of nylon filter on top of another P1000 pipette tip while pushing the cut pipette tip down into the filter. A third pipette tip was used with a pipette to add the sample to the filter. The pipette tip was then discarded, the pipette was attached to the filter, and gentle pressure was applied to expel the last bit of sample.

Once filtered, the cells needed to be fixed before flow cytometry analysis. This was done by adding 4% formaldehyde to the filtered cell sample and were directly after washed three times with PBS. The cells were then fixated and could be taken out of the Class II lab for flow cytometry.