Maybe run your DNA sequence with your FLAG tag though a secondary structure predictor to see if a weird hairpin develops that prevents proper translation.
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Is there any increase in mRNA levels? I wonder if this is prone to degradation, especially since you can see the GFP.
When you say that you cannot reproduce the results, are you saying that right after cDNA prep you can obtain a band and a week later you can't?
This may sound insulting, but are you handling your reagents properly. I have seen people hold their SYBR green master mix on a vortexer for <10seconds, ultimately destroying the polymerase.
It is indeed funky that your qPCR only works a handful of times. Have you let someone prepare your samples for qPCR?
Have you tried scrubbing your pipettes and bench? Maybe you have a potential DNase contamination.
That is weird. Do you add anything to your water to curb the growth of bacteria (i.e., SDS)? It sounds like your setting are off on your incubator. You shouldn't have an abundance of condensation like you are seeing.
Edit: What kind of system are you using? Air or water-jacketed?
Yeah. Not a very informative method, but it can tell you the purity of the DNA/RNA. Absorbances of 1.7-1.9 usually signify purer DNA. You can sometimes use this as a close estimate to the relative amount of DNA, as compared to RNA; however, it is all relative. You can't really base it on this number, especially since cDNA protocols contain a lot of contaminants.
If you can spare some of your sample, run it on a fluorometer. Fluorometers can usually detect small ng amounts of DNA, so you could possibly dilute a small amount (<0.25uL) of your sample.
Sorry I don't know have additional recomendations
shRNA can be tricky. The main thing is you do not know where the shRNA will intergrate into the genome. It is possible that the shRNA is integrated into inactive heterochromatin or recombination damaged the shRNA construct. This would maintain antibiotic resistance, but hinder shRNA expression. It is also possible that something is altering the formation of your shRNA. When you ordered the lentivirus, did they give you an information sheet with binding information? Sometimes this costs extra, but it would allow you to check possible binding sites of your shRNA.
Alot of things could be going wrong. It is hard to pinpoint just one.
Did you use a negative/positive control?
Did this help?
It is hard to say without using a DNA specific dye. You would be detecting both DNA and the degraded RNA. What is your 260/280 ratio? Can you spare 1uL and run your sample on a fluorometer? If we are just ballparking it I would just subtract 50ng and say you have an estimated concentration of 100-115ng/uL.
You could enzymatically deglycosylate your proteins. It wouldn't be 100%, but you should see an relative increase in your band intensities. You could convince your reviewers on the effectiveness with any RT-PCR data you have on the isoforms.
NEB enzyme people have used in the lab (I'm sure others exist).
Edit-You are probably already aware, but you may be able to get a free sample to convince your PI it works before purchasing.
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.
When you import your new ligand and your orignal protein dissappers, does it still list the original protein in the right side bar? It could be that you are entering a very small ligand (compared to your protein) and it blowing out your protein from view. I would try centering on your protein (easily done in protein drop down menu) and then moving your ligand to your protein.
Depending on the complexity of your ligand, you can easily draw it within Pymol. I have found that for some projects, this is much easier.
I think the issue is that when you upload your ligand, your protein gets pushed out of frame.
Let me know if that works.
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.
The two replies above are spot on. Depending on the amount of information on your protein, I use http://dbptm.mbc.nctu.edu.tw/. This will tell you all of the experimentally determined post-translational modification sites that are on your protein of interest. From here you can use inhibitors to see if they alleviate the more prominent band.
Are you waiting long enough in between each reading (mutiple readings on same sample)? It seems like you understand the issue with temperature and concentration reading. I would try using the standard with each machine and then taking a manufacturer provided plasmid and reading the concentration on each machine. This could tell you which machine is right, if both are wrong or if an undergrad left the standardizing solutions at 25C for a week.
Probably not the case, but open the Fluorometer lid and check and see if you see any dried solution on the spec's lenses. Dried liquid will disrupt the emitted wavelength and give you funky results.
If you have an additoinal spec laying around (nanodrop), verify the standard concentrations. Each standard will have a predetermined concentration in them (i.e., standard; 0-200ng/uL). This may confirm if someone has "double dipped" into your standards.
I never add the PI at that step, just because I will be denaturing it at 95c within 30mins. I have never had any issues.
Adding PI to the wash buffer following bead pulldown seems kind of excessive. I would try them side by side and determine if the results change. At least this way if anyone higher up has a problem with it, you have evidence supporting you.
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).
Same here. We autoclave regular tap water and supplement the water with SDS. Distilled and Milli-Q water will rust your incubator.
What is the purpose of adding SDS? As an anti-bacterial?
We have seen it reduce the growth of bacteria. It is especially helpful if someone accidently splashes media into the water tray.