Drawing of metabolic cages

I want to analyze my RT-qPCR experiments. I want to compare the expression of genes in cell type A, B and C and I have this cell types from the individuals 1, 2 and 3 (this are my biological replicates n=3). I have a reference gene (housekeeping gene, GAPDH), but in this case I dont have a control sample. For example I could normalize all values for each individual 1, 2 or 3 on cell type A from each individual. But this feels wrong because the standard deviation for cell type A will be nearly 0 and a statistical comparison with this cell type will not be possible.

Would it be also possible to choose the lowest value I measured in all samples from all three biological replicates and then normalize every value to it? Or to apply the deltadelta ct method and just normalize the values to the reference gene (housekeeping gene) without normalizing it to a control sample?

Hi,

I've been currently having issues with creating consistent Western Blots, and it's driving me insane figuring out why. The issues I've been having are the concentration of my proteins of interest are never consistent. For instance, one trial I have extremely saturated tubulin bands and almost no GFP bands, while in another trial I have weak tubulin bands and very strong GFP bands. A third trial then shows tubulin showing up relatively strong while GFP not so strong. The point is the results aren't consistent, and I think it may have to do when incubate my membrane with proteins. See I know the antibodies I am using have the greatest affinity for my proteins of interest, as my PI has used them in the past with good results. The problem is then the solution I use. I've been putting 1 microliter of each antibody in 2 mL of PBST buffer, and I get the feeling I'm not pipetting them correctly. I understand you have to dip the pipet into buffer before dispensing, but when I do it for such small volumes and push to the second stop, I get a little bubble that pops out at the tip and then I become unsure whether or not the antibody actually dispensed or not. Can anyone give a little insight into what I may be doing wrong here? Thanks!

Hi everyone! I have stained embryos with JC1 and now I am trying to quantify with the ImageJ software. I got values like area, mean, mode, integrated density and raw integrated density. My question is which value (integrated or raw integrated density) should I consider for quantification? And after considering any of these parameters (integrated or raw integrated density), do I need to measure corrected total cell fluorescence or I can directly use it as fluorescence intensity? Thanks in advance.

I can't find their components online but would like to know the precise inhibitors they use.

I have been having trouble with my siRNA transfections where it seems as though my target gene is not being knocked down by the siRNA. I am using a silencer select pre-designed siRNA from Thermofisher (Ambion), along with Lipofectamine 3000 (Invitrogen). I am using a 6-well plate and my cells (adherent) are approximately 80% confluent when I transfect them for 24 hours and then terminate the well. My siRNA concentration is 2ug. Should I:

1) increase the time they are exposed to the siRNA (to 48 hours?)

or 2) increase siRNA concentration? or 3) Reduce confluency of my cells to about 60 to 70%?

Also, on a side note, whenever I perform western blots on cell lysates as opposed to tissue lysates (from mice), I am getting significantly more nonspecific bands with my cell lysates. I am blocking with 5% milk in TBST for 1 hour. I wash 4x 5min ea with TBST before my primary ab. I incubate my blot in primary ab overnight 4deg. and then I wash 6x 5min. each after the primary.  Is there a way to reduce this background noise on my westerns? 

Recently, I had some problem with my WB.

I used 10% gel, run at 100V for 2 hours. 

Like the following picture: the bands were not concentrated and there were uneven band size. The band is really ugly. Any suggestions ?! Thank you!!

I was performing a western blot today and I made a mistake that after protein transfer, I used skim milk to perform the blocking but I suddenly decided to use Ponceau S to  perform a quick check on my membrane, so I took the nitrocellulose membrane out and stained with Ponceau S. The Ponceau S bonds to the membrane in a great tensity. Although the observable bands imply that the protein transfer was successful, the Ponceau S could not be washed off using distilled water. I flushed that membrane for about 10 min but it was not working, so I used PBS-TWEEN 20 (my wash buffer) to wash the membrane. Unsuprisingly, the membrane was easily cleaned up. I was worried if the proteins transferred from the gel would also be washed off by PBS-TWEEN 20, so I stained the membrane again with Ponceau S and the bands were still observable. Therefore I assume the transferred proteins were still on the memrbane. Here are my questions:

1. How does the protein bind to the nitrocellulose membrane? via what interaction?

2. Why is wash buffer not ablt to wash off the transffered proteins from the membrane?

3. How does skim milk work to block the membrane since the wash buffer can easily wash it off?

Thanks for your help.