Since your single-color compensation controls have to be acquired and analyzed in order to generate the compensation matrix, they are not themselves compensated at the time of acquisition (i.e., you can't apply a matrix that you haven't already generated!). I do this routinely with a 10-color panel. It will absolutely work to collect your compensation controls and samples without compensation applied and then generate and apply a compensation matrix in FlowJo post-hoc. I was wondering if anyone knows what would be the best way to represent this data as overlayed histograms, or whether it is more correct to actually represent them separately, side-by-side, with their actual y-axis counts, and percentage of the population per gate.
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In flowJo, if you overlay the two histograms, it automatically scales the counts' y-axis if selecting the option "relative to mode" this only changes the y-axis so to be a max of 100 if you select manual however, it plots the actual counts on the y-axis (which are different between my samples).
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The differences are very clear by looking at the percentages (frequency of parent) of each cell cycle stage (different gates), but I'm quite concerned on how to represent the histograms overlayed.
The problem was that in certain cases, the number of treated cells was rather low, so I wasn't able to acquire as many cells as in the untreated group. I have data from an experiment in which I'm comparing cell cycle profiles (PI staining) in treated vs untreated cells. This is also backed up by the fact that I didn't have a high percentage of dead cells or low number of total cells for the conditions in which there was no necrotic spleen tissue (negative controls and Ova+CpG). I suspect that I may have lost a lot of my live cells when I removed this pellet, as my collaborator mentioned that often the live cells will stick to the dead cells. I did notice that there was an insoluble pellet of dead spleen tissue that I had to remove from the tubes in most of the experimental conditions post-ACK lysis. We then did an ACK lysis step to remove RBCs, after which we counted the cells. These 5 organs were placed onto a 40uM filter and mashed up with the rubber end of a 1mL syringe, followed by 4 successive washes with media (we pooled the lymphocytes and splenocytes together, again to try and maximize our chances of seeing our Ova specific T cells). Does anybody know how long after a booster injection I should look at the T cells?įor obtaining lymphocytes and splenocytes, we removed spleens and both inguinal and popliteal lymph nodes from each mouse. Is it possible that I waited too long after the booster injection to analyze the T cells? What I'm thinking is that by 7 days after the boost, most of my Ag-specific T cell population has died off, perhaps due to lack of persistent antigenic stimulation. Additionally, gating on the remaining dead cell population in FlowJo revealed that these cells were primarily CD8+ Tetramer+. I believe I lost a lot of my Ag-specific T cells in this dead cell population - mostly due to the fact that despite the macroscopic evidence of an immune response I wasn't able to see an increased percentage of antigen specific T cells. The lymph nodes in the experimental conditions were quite enlarged and inflamed however, there was also a huge amount of cell death in these conditions. I analyzed the Ova-specific splenocytes and lymphocytes on D14 with MHC tetramers. The regimen is a primary injection on D0 followed by a booster injection on D7.
I'm performing an In vivo experiment in which I inject mice with a DC vaccine containing ovalbumin.
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It's also a question of who will use the software in the lab - if it is only you and you are already a FlowJo expert you may stick with it, but if there are people less adept with it I think Cytof is easier to learn, and since there is less flexibility analyses from other users are easier to understand and to review. But if you need something different, there is no way to change it. On the other hand, Flowjo also has some features which at least I have not found yet in Cytobank (overlay of scatterplots for example), and it allows for more fine tuning, customizing and parameters to be defined by the user (like setting gates for compensation manually etc.), whereas Cytobank sometimes appears a bit like an Apple product - very good, very intelligent, perfect for 90% of all situations. So, when you do Cytof or more than 15 channel FACS and you want to use tools like Spade or Visne there is probably no way around it. Cytobank has very nice and modern looking graphs, a very intuitive and intelligent work flow and user-interface and is very good with high-throughput and especially multi-channel analyses. In my opinion it depends a lot on what you plan to do with it and what kind of "style" you prefer.
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