Centrifuge for single-layer renal cell culture

Tissue culture involves removing tissues or cells from an organism and culturing them artificially in vitro to enable their survival, growth, and reproduction outside the body. Tissue culture can be categorized into organ culture, tissue block culture, and cell culture. Currently, cell culture is the most widely used technique in practical applications. It is commonly employed for the diagnosis of viral diseases and for the production of biological products (such as vaccines and diagnostic antigens).

There are many methods of tissue culture, broadly categorized into suspension culture and immobilized culture. In suspension culture, minced tissue pieces are separated by centrifugation using a centrifuge. The dispersed cells obtained through low-speed centrifugation are then placed in an appropriate culture medium, where they float freely in the liquid for cultivation, after which viruses are inoculated.

In addition, there is the fixed-culture method, in which tissue blocks are dispersed by low-speed centrifugation and then allowed to adhere to the container walls for culture. Once the cells have grown well, a virus is inoculated. The primary indicators of viral replication in tissue-cultured cells include cytopathic effects (such as necrosis, disintegration, and sloughing), cell fusion or inclusion-body formation, and the ability of the virus to adsorb onto red blood cells.

The following provides a brief introduction to the general operational techniques for “single-layer renal cell culture” using a low-speed centrifuge.

1. Kidney Collection: Using sterile techniques, remove the kidneys from young animals around weaning age and place them in a sterile petri dish. Trim off any remaining renal capsule and fat, then longitudinally cut each kidney into two halves. Remove the renal capsule and fat again, and further longitudinally split each half into two pieces. Excise the renal pelvis and medullary region. Wash the kidneys thoroughly with Hank’s solution containing double antibiotics (i.e., penicillin and streptomycin; the same applies hereafter), transfer them to a small beaker, and mince them into a milky paste. Then wash the minced tissue 2–3 more times with Hank’s solution until the liquid becomes clear and free of blood cells.

2. Digestion: Add 10 times the volume of 0.125% trypsin solution for digestion. There are two methods: hot digestion (in a 37℃ water bath) and cold digestion (in a 4℃ refrigerator). The duration of digestion varies depending on the type of tissue cells and the activity of the trypsin used, until the kidney tissue, which initially forms flocculent clumps, becomes uniformly dispersed again. Remove the digested tissue and discard the trypsin solution; wash the tissue several times with Hank’s balanced salt solution. Then, add a small amount of Dulbecco’s Modified Eagle’s Medium (DMEM), and use a pipette to gently pipette up and down (i.e., aspirate and expel repeatedly) 10–20 times to dissociate the tissue into individual cells. Filter the resulting cell suspension through 2–3 layers of sterilized gauze. Transfer the filtrate to a low-speed centrifuge and centrifuge at 1500 rpm for 10 minutes. Discard the supernatant, resuspend the pellet in an appropriate volume of DMEM, and centrifuge again at low speed. Finally, add more DMEM to suspend the cells completely, and collect the cell suspension into a sterile Erlenmeyer flask.

3. Cell Counting and Aliquoting: Take the prepared cell suspension and count the cells using a method for counting white blood cells. Based on the counting results, dilute the suspension with nutrient medium (pH 6.8–7.0) to achieve a concentration of 500,000–600,000 cells per milliliter. Aliquot the suspension into small flat bottles (approximately 5 ml of suspension per bottle with a capacity of about 50 ml), or alternatively, aliquot it into streptomycin vials.

4. Culture: Incubate at 37℃; after 3–4 days, cells will adhere to the flask wall and form a monolayer. Thereafter, replace the maintenance medium every 3–4 days.

5. Virus inoculation: Cut the diseased tissue into small pieces, add Hank’s solution containing double antibodies at a ratio of 1:4, and grind thoroughly until a milky emulsion is formed. Subject the mixture to freeze-thaw cycles to lyse the cells and release the virus fully. Then, using a low-speed centrifuge, adjust the speed to around 3,000 rpm and centrifuge for 15 minutes. Transfer the supernatant into culture flasks already seeded with a monolayer of cells, allow it to adsorb onto the cells at 37°C for 30–60 minutes, and then add maintenance medium to continue culturing.

6. Harvesting the virus: Observe cell cytopathic effects daily or every other day under a low-magnification microscope. When 50–75% of the cells exhibit cytopathic changes, collect the culture medium from the flask and store it in a refrigerator (-20℃). This constitutes the virus stock for propagation.

7. Precautions: The cleanliness of petri dishes and other utensils, the proper use of low-speed centrifuges, water quality, pH levels, and aseptic techniques—all are critical factors that determine the success or failure of tissue culture. These must be carried out strictly in accordance with established requirements.