Electroporation-Based Delivery of Fluorescent Biomolecules into Bacterial Cells

Begin with a chilled cuvette containing a suspension of electrocompetent bacterial cells, which have increased susceptibility to membrane permeabilization.

The suspension also contains biomolecules tagged with organic fluorophores.

Apply a high-voltage electric pulse to create temporary membrane pores, enabling biomolecule entry into the cytoplasm.

Add a nutrient-rich medium and transfer the cells to a tube. Incubate to promote membrane repair.

Centrifuge the cells, then discard the supernatant containing non-internalized biomolecules.

Resuspend the cells in a high-salt buffer containing a detergent to detach surface-adhered biomolecules.

Centrifuge again, discard the supernatant, and resuspend the cells in the buffer.

Transfer the suspension onto a filter and centrifuge to discard the flow-through containing any non-internalized biomolecules. Then, rinse the cells with fresh buffer and resuspend them.

Mount the cells onto an agarose pad and place a coverslip. The fluorescently labeled bacteria are ready for visualization.

Starting with a stock of fluorescent biomolecules prepared in a low ionic strength buffer, transfer up to five microliters of the biomolecules into either a 20 microliter aliquot of competent bacteria or 50 microliters of competent yeast cells. The amount of fluorescent biomolecules incubated with the cells directly correlates with the amount of biomolecules internalized per cell after electroporation.

Incubate the mixture on ice for up to 10 minutes. At the same time, select an electroporation cuvette of appropriate electrode spacing and chill on ice. Transfer the cell mixture into the pre-chilled cuvette.

Gently tap the cuvette a few times to remove any bubbles from the solution. Remove any moisture from the cuvette with a paper towel and place the cuvette into the electroporator. Apply a high voltage pulse to the cells.

Just after this process, verify that the time constant displayed on the electroporator is between four and six milliseconds. Lower time constants are often due to the presence of excess salt or bubbles within the cuvette, and it may decrease or completely inhibit uptake of biomolecules into the cells. Higher voltage pulses improve the cell loading, but can also affect the cell viability.

Immediately after electroporation, add 500 microliters of rich media and transfer the cells into a fresh 1.5 milliliter tube. Let the cell membrane recover by incubating the bacteria at 37 degrees Celsius or yeast at 29 degrees Celsius for 2 to 10 minutes. After biomolecule incorporation and growth recovery, spin down the cells for one minute in a four degrees Celsius pre-chilled centrifuge and discard the supernatant.

Resuspend the pellet in 500 microliters of buffer and repeat the spin down, supernatant removal, and buffer resuspension processes three more times. These wash steps will remove all traces of noninternalized biomolecules from the cell suspension.

In the case of labeled protein internalization, an additional purification step is performed by pipetting the cell suspension into a 1.5 milliliter tube fitted with a 0.22 micron membrane filter on top. Separate the cells from the buffer by centrifugation while ensuring there's enough liquid remains above the filter to avoid the cells drying out, and discard the flow-through fraction. Below the filter, add 500 microliters of fresh PBS over the cells and repeat the combination of a spin down followed by fresh PBS addition two more times.

Perform a final spin down at 3,300 g for one minute to remove the previous PBS wash. Resuspend the cells with 150 microliters of PBS.

Finally, remove the upper cover slip from the previously made agarose pad. Load the cells onto the pad by pipetting 10 microliters of the cell suspension dropwise. After placing a new, cleaned cover slip on top of the agarose pad, the sandwiched cells are now ready for fluorescent microscopy.