Dear all,

I am currently performing CRISPR/Cas9-mediated knockout of the SLC35B4 gene in HT29, Caco2, and HepG2 cell lines using the Santa Cruz Biotechnology (SCBT) CRISPR/Cas9 KO double nickase plasmid and the UltraCruz® transfection reagent.

We are following the SCBT protocol that is originally designed for 6-well plates, and we adapted it to a 96-well format by dividing all reagent and DNA quantities by 15, as recommended.

Our Current Protocol in 96-Well Plate:

• Cells were seeded at 30,000 cells/well in antibiotic-free complete medium (DMEM + 10% FBS) one day before transfection.
• We used 0.1–0.2 µg plasmid DNA per well.
• UltraCruz transfection reagent was used in the range of 0.6–1.0 µL per well.
• DNA and reagent were diluted separately in SCBT’s plasmid transfection medium, then mixed and incubated for 20 minutes at room temperature before being added (30 µL/well) to the cells.
• After 24–48 hours, transfection was evaluated via GFP expression, and puromycin selection was started at 48–72 hours using pre-determined lethal doses for each cell line.

Despite careful optimization within the recommended ranges, we observed low transfection efficiency, particularly in Caco2 and HepG2 cells, with weak GFP expression and limited survival post-selection.

I would truly appreciate your input on the following:

Any protocols, tips, or shared experiences would be incredibly helpful as we try to troubleshoot and optimize our system.

Thank you so much in advance for your support!