We bought customized pLenti-shRNA-GFP with anti-blasticidin selection gene. I tried the MOI from 1 to 55 previously. After antibiotic selection for 6 days, all non-transduced cells were dead. I kept culturing the viable cells (very few) in blasticidin-contained media, but around a week later, they all died. I tried this time a higher MOI from 55-100, but when checking the transduction efficiency with fluorescent microscope 24 hours post transduction, there was no fluorescence at all. It is recommended that the fluorescence should be checked 48-72 hours post transduction, but I would assume that 24 hours is enough for the virus to integrate into the host cells and even though the efficiency is low, there should not be no fluorescence at all. I used polybrene 8 ug/ml as recommended and my cells are not sensitive to polybrene as tested before. Right before checking under fluorescent microscope I replaced the virus contained media into normal culture media. And normally the day after, I would start antibiotic selection. Does any of you have any idea why such things happen? Should I run PCR to test the knockdown efficiency before blasticidin selection by comparing transduced versus non-transduced cells?
There are two things that may be the cause of your problem: 1. the titer of your lentivirus is lower than you think it is/the virus non-infectious, 2. the concentration of antibiotic you are using is too high and is killing your cells even though they have been transduced (have you done a kill curve on your cells to know the concentration where all the cells are dead?). Also, you will not see fluorescence until at minimum 48 hours but more like 72-96 hours. Have you looked up what MOI is typically used for your particular cells? Too high of an MOI will kill cells as well.
Hi Tom, thanks to your response. I have done the killing curve and it worked well. For my cell line the MOI has not been reported yet since this is a chicken line developed in our department. I am also wondering if the titer is lower than it is said. Would you like to give me some suggestion on deciding the titer? I have very limited amount of virus and do not want to waste too much on this titer test.
Hi Yang,
Since your virus has GFP in it you can easily titer it on HEK293s and then either count GFP positive cells or use flow cytometry. I usually set up my titers in a 6-well dish using 1x105 cells and varying amounts of virus (i.e. 1ul, 0.5ul, 10-1, 10-2, etc). You can do the tranduction while the cells are in suspension (add cells, virus, polybrene, media to a tube, mix and plate). By doing the transduction in suspension you know exactly how many cells are being transduced. Change media after 48 hours and then look for fluorescence at 96 hours. You want the percent cells tranduced to be between 5-30% such that you can calculate the titer accurately.
Are there MOIs reported for any chicken cell lines? That may be a good place to start. Also, be sure you are not putting your virus through multiple freeze-thaw cycles since that will result in a loss of in titer of the virus.
Hi Tom,
Sorry for the late reply. Thank you very much for your kind suggestions! So I would like to confirm with you the principle behind titer determination. You suggested to use HEK293s cell line to determine the titer. Is it because this is a stably established cell line that can replicate infinitively, so that when I put the virus particles (e.g. 1 uL) in 5*10^4 cells for example, though HEK293s does not have my target gene sequence, the virus still should integrate into the genome and be reproduced with cell proliferation? Let's assume that the transduction efficiency is 30% after 96 hours detected by flow cytometry. Does it mean that the titer should be (5*10^4 (cells/mL)*30%)/(1*10^-3 (mL of virus))=1.5*10^7 LP/mL? Or when calculating, the cell number should be that at 96 hours post-transduction? And if this is the principle, does it mean that I can test the titer with any stable fast proliferating cell lines, such as cancer cells and embryonic stem cells?
Last question I want to ask is that if the titer is indeed very low, is there a way to re-condense it so that I don't need to put too much virus-contained media (to let virus and target cell contact better) for transduction?
Thank you so much!!
Hi Yang,
The reason to titer the virus on HEKs is because they are very easily tranducible and yes the virus will integrate and express GFP which can be measured and used to calculate the titer. The titer you calculate is based on the initial number of cells you use for transduction (i.e. 5x104 cells). So the formula you wrote is how you calculate the titer. HEKs are really the best cells to use since the virus is typically made in this line. Other cell lines might not be as easily transducible. Once you know what the titer is then you can add virus at whatever MOI you want (i.e. 5x104 cells with 2.5x105 virus for an MOI of 5).
To concentrate the virus you can centrifuge the supernatant through a 20% sucrose cushion at 20,000xg for 4 hours at 4 degrees C which pellets the virus and you then remove the supernatant and resuspend the viral pellet in a smaller volume. However, after you concentrate you will have to titer the virus so you know what the concentration is.
One thing you should be aware of is that lentivirus typically has the VSV glycoprotein on its surface and uses that to bind to the LDL receptor on the surface of cells. So if your target cells do not express a lot of LDL receptor then no matter how high an MOI you use, you will not get good transduction efficiency. I had this problem when trying to transduce rat stem cells. I went up to 40 and 80 MOI and could not get higher than ~75% transduction efficiency plus that much virus causes a lot of cell death. Conversely, I can get almost 100% transduction efficiency on human stem cells with an MOI of 5. So its possible you may see something similar with your chicken line.
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