Figure out your OD. You could easily calculate "mg" of cells from this number.
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How are you transfecting your cells? Are you doing a transient transfection, waiting two days and then transfecting again?
Do you linearize your plasmids when you transfect your cell with multiple plasmids. If not, I would at least linearize the LacO plasmid. That guy is huge and the cell may have some difficulty taking it up.
I would start by transfecting your lowest transfection efficiency plasmid, selecting for it and then proceeding with the others. How are you selecting for your plasmids? Does each plasmid harbor a different mammalian resistance or are you doing downstream assays to check the expression of each plasmid?
It would be better if you could provide a detailed description of what you are doing. There are tons of cloning resources on this website that you might consider browsing through. My most common cause of ligation failure is proper digestion of my PCR fragments. You usually get good digestion with a vector due to its size and shape, but PCR fragments can be problematic depending on the size.
I usually like to test the efficiency of my ligation by running a simple PCR with a primers that recognize both the vector and PCR fragment. Amplification is a good indication that some ligated products are in your sample.
What are your JM109 cell controls like? Do you observe a good transformation efficiency?
You just shocked the system. When I add antibiotic to my tissue culture media (DMEM), I see an immediate color change. After letting the media sit for ~12hrs, the color is restored. You just need to keep in mind that pH indicators are present in the media and when a solution is added, it needs to fully equilibrate. I wouldn't worry as long as you followed the protocol.
Try gently mixing the solution or placing it in a water bath for about 30mins.
Go review Glycolysis and the Krebs cycle and look for any enzyme with the work "dehyrogenase" in it. This will show you where the protons are being generated. I am not sure if I understand the second question. My only knowledge on that would be that protons are released into the "actual media" due to symport and antiport for the use of bringing in the nutrients that are present in your DMEM (or other media). This is the energy that is required for specific uptake of some of your nutrients.
This is as much as I can elaborate without reviewing the relevant literature.
Hope this helps.
As with any living tissue, cellular respiration (glycolysis and oxidative metabolism) generates protons that can reduce the pH of cell media. It is the same as a respiring tissue within a living organism; however, a circulatory system is absent to replenish the media.
*On a side note. Pour bleach into your media and this will demonstrate that H+ ions are responsible for your pH shift. I do not know why I get such a thrill by doing that.
Do you have multiple cell counts at different time points or are you just given one number? Go to ATCC and find the growth curve of your cells and from here you can determine what phase your cells were in.
Double check your cut site to make sure that you are getting the correct dropout size. Salt concentration can have an effect on the migration of DNA in gels, but from your post I would assume that the size difference is significant.
Do you see the 50bp product in your controls without primer? You have used this primer before with no additional bands or is this a recently designed primer? Does the appearance of the additional band only occur with this sample?
Are you trying to compare the fold increase of your cytokine at time point zero vs. a later time point?
There are a couple of different ways you can approach your quantification depending on what you are trying to show. Does the treatment of your sample suddenly switch on your cytokine or is your cytokine usually differentially expressed? Are you trying to compare your cytokine expression to the expression of a common ubiquitous cytokine?
I am really confused. You designed this to test whether your deletion was present in a large portion of cells. You are looking at relative fluorescence to see how many cells have the deletion, as compared to a non-deletion containing control? What you see is that your "deletion containing cells" do not have the relative fluorescence that you were hoping for. So, you are looking for an easier method to determine the ratio of cells that contain the deletion?
I usually use 2ug of RNA and get good, consistent results. The amount of RNA will not affect the qPCR, because your cDNA protocol should require the degradation of your RNA products following Reverse Transcription.
Edit: The ending amount of RNA will not affect your qPCR data.
1) I never add water. Are you adding water to get your DNA dye down to 1X? When I set up my PCR I add a consistent concentration of DNA so I can quantify my amplicons using the DNA ladder.
2) You probably don't add water here because you would overload the well and your sample mix would spill out.
3) Use one lane if the well can accommodate the entire sample. This will reduce the total amount of agarose an reduce the volume of reagents that will be required for gel purification.
4) No. Review the methodology behind electrophoresis on this website or wiki.
I have used SYBR Green. I usually used a total reaction volume of 12uL. The final primer concentration would have been 2-10uM (I played around with this and the results stayed consistent - trying to determine the minimal amount of primer I could use - check SYBR green protocol). I would add 0.5uL of a 2ug cDNAtemplate. I would add 6uL of the SYBR Green master mix.
My thermal cycling program:
Doublets of the reaction mixture were loaded and cycled once at 50ºC (2mins), and 95ºC (10mins); 40 cycles at 95ºC (15secs), 58ºC (30secs), and 72ºC (30secs); and a final cycle at 95ºC (15secs), 60ºC (20secs), 95ºC (15secs), and 60ºC (15secs).
The 58C will be unique to your primer Tm's. I would always try to design your primer to a specific Tm. This will allow you to perform very diverse RT-PCR's simultaneously (saves plates and optical film).