Cell Line Information
● Cell line name: HeLa (human cervical cancer cells)
● Mutant gene: NUMA1
● Mutation sites: K1330R/K1335R
● Species: Human
● Cell morphology: epithelioid, adherent growth
● Complete medium composition: MEM (NEAA) + 10% FBS + 1% PS
● Passage ratio: 1:2 to 1:4
● Passage frequency: Passaged once every 2–3 days.
● Mycoplasma test: Negative

Gene Editing Information
● Construction method: The theoretical basis for constructing cellular point‑mutation models is homologous recombination repair. When the double strands of genomic DNA are cleaved by external factors, the cell’s DNA repair mechanisms are activated to repair the broken DNA ends; depending on the repair pathway, this can be classified as homologous recombination repair (HR) or non‑homologous end joining (NHEJ). Homologous recombination involves using a homologous DNA sequence as a template to achieve precise repair, whereas non‑homologous end joining entails the direct ligation of the broken DNA ends by cellular proteins, which may randomly introduce or delete a variable number of nucleotides. Based on this theoretical framework, the construction of point‑mutation cell lines proceeds in three steps:

1. Site-specific DNA cleavage ： During this process, the Cas9 protein is transfected into the target cells together with the sgRNA. Under the guidance of the sgRNA, the Cas9 protein cleaves the DNA at the target site, generating a double-strand break.
2. Homologous recombination repair of DNA ： After a double-strand break is induced in genomic DNA, an additional homologous repair template must be provided to the cell so that DNA can be repaired via homologous recombination and mutations can be introduced at the target site. This homologous template sequence is delivered into the cell by co-transfection with the sgRNA and Cas9 protein.
3. Positive cell screening ： For the point‑mutant cells generated by the aforementioned method, no additional reporter or resistance genes were introduced, thereby minimizing any impact on unedited genomic loci. Single‑clone cells were isolated from the edited cell pool and subjected to deep sequencing to obtain the expected point‑mutant clones.

● Application scenarios: Liman Bio’s point‑mutation cell lines precisely mimic single‑base substitutions, making them ideal for gene‑function analysis and disease‑mechanism studies. These platforms enable the generation of pathological cell models—such as those for cancer and genetic disorders—enabling drug‑target validation and resistance‑assessment assays, thereby providing a standardized, highly reliable in vitro experimental system for basic research and new‑drug development.

● Validation data:

Cell Culture Instructions

● Cell recovery

1) Remove the cryovial from the liquid nitrogen tank or the −80°C freezer, and promptly place it in a 37°C water bath, gently swirling it to facilitate rapid thawing. To minimize the risk of contamination, you may first transfer the cryovial into a clean, disposable PE glove before immersing it in the water bath.

2) Once the liquid in the cryovial has thawed, remove it immediately and wipe the surface of the cryovial with a 75% ethanol‑soaked gauze.

3) Place the cryovial in a biosafety cabinet, transfer the contents of the cryovial into a centrifuge tube containing approximately 5 mL of complete culture medium, centrifuge at 200 × g for 5–10 minutes, discard the supernatant, and proceed.

4) Resuspend the cells in an appropriate volume of complete culture medium, transfer them to a new culture flask, and incubate the cells in a 5% CO2, 37°C incubator under static conditions.

● Cell passage

When cell density reaches 70%–90%, subculture can be performed at a ratio of 1:2 to 1:4, with subculturing every 2–3 days. The specific procedure is as follows:

1) Aspirate and discard the culture medium from the flask, add an appropriate volume of calcium- and magnesium-free PBS, gently swirl to rinse the cells 1–2 times, then aspirate the PBS.

2) Add approximately 1.0 mL of 0.25% trypsin (use this volume for a T25 flask; for a T75 flask, you may increase the amount slightly). Gently rock the flask to ensure all cells are evenly covered, then place it in the incubator and allow digestion for 3–4 minutes. Under the microscope, observe that the cells have rounded up; gently tap the flask wall and, when about 80% of the cells have detached, add approximately 3–5 mL of complete culture medium to neutralize the trypsin. Carefully pipette up and down to detach the cells from the flask surface, dispersing them into a single-cell suspension. (Adjust the volumes of trypsin and neutralizing medium according to the flask size.)

3) Transfer the cell suspension after digestion into a clean 15 mL centrifuge tube, centrifuge at 1000 rpm (approximately 250–300 × g) for 5 minutes, and discard the supernatant.

4) Resuspend the cells in an appropriate volume of complete culture medium, transfer them to new culture flasks according to the passage ratio, replenish with fresh complete culture medium to the desired final volume, and incubate the cells in a 5% CO₂, 37°C incubator under static conditions.

● Cell cryopreservation

Cell cryopreservation can be performed when cell confluency reaches 80%–90% and the cells are in good growth condition. The homemade cryopreservation solution consists of 90% serum and 10% DMSO. To ensure high post-thaw viability, it is best to prepare the cryopreservation solution in advance and pre‑cool it at 4°C. The recommended cell density for cryopreservation is 5 × 10⁶ to 1 × 10⁷ cells/mL.

The specific procedures for cell cryopreservation are as follows:

1) Aspirate and discard the culture medium from the flask, add an appropriate volume of PBS to wash once, and then carefully remove all the PBS.

2) Add approximately 1.0 mL of 0.25% trypsin (use this volume for a T25 flask; for a T75 flask, you may increase the amount slightly). Gently rock the flask to ensure the enzyme evenly covers the cells, then place the flask in the incubator and allow it to digest for 3–4 minutes. Under the microscope, observe until the cells become rounded. Gently tap the flask wall; when about 80% of the cells have detached from the surface, add approximately 3–5 mL of complete culture medium to neutralize the trypsin. Gently pipette up and down to detach the cells from the flask surface, dispersing them into a single-cell suspension. (Adjust the volumes of trypsin and neutralizing medium according to the flask size.)

3) Transfer the digested cell suspension to a clean 15 mL centrifuge tube, centrifuge at 1,000 rpm (approximately 250–300 × g) for 5 minutes, and discard the supernatant. Cell counting can then be performed to facilitate adjustment of the cryopreservation cell density.

4) Add an appropriate volume of pre‑prepared, pre‑cooled cryopreservation solution based on the cell pellet size (recommended: 1–2 mL per T25 flask; 3 mL per T75 flask), and gently pipette up and down to ensure uniform cell dispersion.

5) Transfer the cell suspension into cryovials, 1 mL per vial. Label the cryovials in advance or affix labels. The label should include the cell name, passage number, cryopreservation date, and the operator’s name, among other information.

6) Place the cryovials in a cell‑freezing box, and then transfer the box to a −80°C freezer overnight. After overnight storage, transfer the vials to a liquid nitrogen tank for long‑term preservation; following transfer, it is recommended to thaw one vial to verify the viability of the cryopreserved cells.

Click the link below to view detailed information.

Product Manual | LIMAN Easy Point Mutation Cell Line Construction Custom Services and Products

Service Guide | Q&A on Leman Point-Mutant Cell Line Construction Services [Version 2411]

Service Guide | Comprehensive Q&A on Site‑Directed Mutagenesis–Based Stable Cell Line Construction: From Concepts to Experimental Details—All in One Article! [Version 2512]