Stewart Laboratory Standard Operating Procedure
Creation and use of your infectious vector:
Plate 5 x 105 293T cells in 6 cm2 dishes containing 5 ml of media. (This can be scaled up if desired).
The following day set up (use polypropylene tubes for this; polystyrene tubes DO NOT work!).
1 ug retroviral DNA encoding gene X
1 ug packaging plasmid (such as pCLEco, pCLampho, pUMVC, pHR’CVM8.2 deltaR etc.). If you are using a 3 plasmid system then:
for lenti: 1 μg packaging (pHR’8.2deltaR) at a 8:1 ratio with the envelope plasmid (pCMV-VSV-G).
for MuLV: the packaging plasmid (pUMVC3) at a 8:1 ratio with the envelope plasmid (pCVM-VSV-G)-a total of 1ug.
DME without serum to 94μl
6 μl Fugene
Mix and wait 15 to 30 minutes at room temperature
Add to 293T cells without touching the sides of the dish (DO NOT CHANGE MEDIA).
If you are using amphotropic virus then move immediately to BL2+ in a secondary container, which has an absorbent material. (This does not mean a couple of hours; it means Immediately!). The rest of this protocol is the same for all virus---the BL2+ safety practices are in place if you are using amphotropic viruses.
The following day change the media to whatever media you wish to use when infecting target cells. 293T cells are easily detached so remember not to put the media directly onto to cells but rather “run” it down the side of the dish. Remember that you will get the highest titer virus when your cells are “happy.” Plate out your target cells.
The following day. Remove the medium from the 293T cells and use a 0.45 u syringe filter to remove any 293T cells. DO NOT use the 0.2 u filter, as it is likely to shear the envelope from your virus making it noninfectious.
Note: After filtering, the filter should be removed and placed in the biohazard bag in the hood and the syringe rinsed with bleach and decontaminated for a minimum of 20 minutes. It is useful to place, in advance, a plastic beaker with bleach in the hood for this.
Add 8 to 10 μg/ml of polybrene (Hexadimethrine bromide) or protamine sulfate to the virus and add to the target cells.
Carry out infection for 1 to 4 hours. Remove virus and replace with fresh media.
Note: If you wish to do a second infection the following day, it is important to put fresh medium on the cells and not let the virus remain on the cells overnight. The medium contains huge amounts of envelope both associated and unassociated with viral particles that will bind all the cell surface receptors required for virus adsorption, causing their down-regulation. Hence, if you don’t change the medium after the initial infection, very few receptors will be available for the next round of infection. In addition, very few cells tolerate the presence of that much envelope for extended periods of time (i.e. a lot of your cells may die).
Allow the cells to recover and begin to express the virus-encoded genes. The cells usually require 48 hours for this to occur.
Add drug if you are scoring for the presence of a vector that carries the appropriate drug resistance marker. Prior to this step it is advisable each time you do an infection to titrate the drug to be used for selection in order to know precisely how much to add. In addition, it is necessary to bring an extra plate of uninfected cells that are often referred to as “canaries.” Add drug to both plates. When your canaries are dead, you can remove the drug.
Testing for Horizontal Transfer (When using the 3 or 4 plasmid system you are only required to do this every couple of months on a randomly chosen virus prep. When using the two plasmid system or packaging cell lines you must do this EVERY time you make a new virus stock!!)
Once you have cells that emerged from selection and are growing, you can test for horizontal transfer, i.e. for the inadvertent generation of replication-competent virus, which may have occurred during the course of your experiment. Please note that the vector-infected cells, prepared as described above, must be growing and “happy.” You should essentially treat them as if they were 293T cells. When they are 50 to 75% confluent, remove medium from these cells, filter it, add polybrene, and infect your new target cells.
The question is often raised concerning which cells should be used as infectable target cells in order to test for the horizontal transfer resulting from the inadvertent creation of infectious retrovirus. The best choice is a cell type that is easily infectable with the specific viral vector you happen to be using. Useful cells here are those that have a relatively rapid cell cycle and are known to be susceptible to infection by the retroviral vector (and therefore to the envelope glycoprotein) that you are using. At present (6/01), we have found that C3H10T1/2 cells are especially useful here.
Wait 48 hours and add selection drug to these cells AND to a set of canary cells. Both sets of cells should die at the same time. If the canaries die but the cells infected by the viral vector don’t do so with identical kinetics, then you have good evidence that horizontal transfer resulting from the inadvertent generation of an infectious retrovirus has occurred, and all cells and media resulting from this experiment should be destroyed by the addition of bleach and subsequent autoclaving.
Many have asked what to do if they have used green fluorescent protein (GFP) instead of a selectable drug marker in the vector. In this case, you proceed exactly as outlined above. 48 hours later after infection of test target cells, instead of adding drug you lift the cells and the canaries and resuspend them in 2% formaldehyde or paraformaldehyde and run FACS to determine if any of the cells are green. You should run at least 10,000 cells to be sure. If you don’t know how to run flow speak with someone who does. Also, both formaldehyde and paraformaldehyde autofluoresce in the green channel, so it is important that you have a set of infected cells canary here as well.
Selection (drug concentrations to be used with CH310T1/2 cells):
Neomycin 1 mg/ml
Hygromycin 300 μg/ml
Puromycin 2 μg/ml
Zeocin 1.2 mg/ml
Blastocidin 15 μg/ml