Myworkismyplay

Doing a simple alkaline digest of tail snips or ear punches provides plenty of gDNA and without all the inhibitory ingredients from PK digestion, and takes much less time, as well as saves tons of $$$.

Add 0.5mm tail or 1-2mm ear punch to 30-40 uL 10mM NaOH. Heat to 95C for about 30-45 minutes (shaking at 500rpm, or vortex every 5-10 minutes). Alternatively I've done 60C for 20 min before the 30 min 95C incubation to help in the solubilization of the DNA (not getting caught up in the insoluble proteins denaturing within the sample becoming insoluble). Vortex briefly to ensure sample tissue is sufficiently broken up. After the heat/mixing step, spin down & cool on ice. Add equal volume (30-40 uL) 10mM Tris, pH 7.5 to neutralize the pH (still leaving basic, for best solubility of DNA). Keep cold for subsequent protocols.

For PCR just use 0.5-2ul per 20ul reaction. You don't need to centrifuge the tail digest to remove impurities & pellet crap, just don't pipet up the blob for PCR. if it really bothers you you can centrifuge at 5-10 min at 10,000rpm (cold is better) & transfer the supernatantl to a new tube.

If your PCR reactions are optimized and primers work well, there shod be any issues for this to work for you.

We clip tails, digest, do PCR, and run our gels all in one day. It saves a ton of time and money. There's no need to purchase expensive DNA extraction kits/reagents for colony genotyping.

For those who just want purer DNA for whatever reason, throw in an ethanol precipitation/wash step, dry, & resuspend in TE.

Good luck!

I would suggest trying to do everything on ice if you are not already. What is your RNA extraction method post-IHC?

I assume the expected yield is from 1 million cells? Viability is the wrong term to use here. As soon as that cell pellet was frozen all of the cells died. You are referring to quantity and quality of genomic DNA from frozen, lysed cells.

The sample seems much too dilute to check integrity and your concentration is most likely way off judging by how low it is and your A260:A230 ratio. Even if you are using NanoDrop, you are really below the accurate threshold for that spec. A better & more accurate method is the Qubit (DNA kit) to tell you how much DNA is really there, or if your A260 I just the residual side peak of the absorbance spectrum dropping past 260nm on the way down from 230nm, where your contaminants & buffer/salt residual material would absorb.

Most likely what you have in that 200 uL is impure nucleic acid, of a very low concentration.

If you are determined to use this particular sample, I suggest a clean-up ethanol precipitation to first bring your volume down, as well as increase the concentration so your spec reading can be more reliable. This will also remove some of the impurities you see at the A230. If done properly, you should recover >95% of your sample.

I would not carelessly toss this "DNA" onto an array and assume the gDNA is of good enough quantity, and especially not of good enough quality to have meaningful, reliable results for your research project.

If you can get a fresh PBMC sample that's what I would start with-- to get quality sample, before wasting money and time with a bad sample. Do the kit extraction protocol according to procedure and see if your sample improves. It should be easy to get good gDNA from PBMCs.

Good luck!

Sometimes try a google search....most of this stuff is easy to find sufficient resources for.

These days vendors are even taking the time to post resources online to help choose the correct product & to understand how the product works.

From Pierce (Thermo Scientific), background on detergents...

- http://www.piercenet...55-4F3977738B63

This is just a small except from their web page:


"Membrane Disruption, Protein Binding and Solubilization

Generally, moderate concentrations of mild (i.e., nonionic) detergents compromise the integrity of cell membranes, thereby facilitating lysis of cells and extraction of soluble protein, often in native form. Using certain buffer conditions, various detergents effectively penetrate between the membrane bilayers at concentrations sufficient to form mixed micelles with isolated phospholipids and membrane proteins.

Denaturing detergents such as SDS bind to both membrane (hydrophobic)and nonmembrane (water-soluble, hydrophilic) proteins at concentrations below the CMC, i.e. as monomers. The reaction is equilibrium-driven until saturated. Therefore, the free concentration of monomers determines the detergent concentration. SDS binding is cooperative (the binding of one molecule of SDS increases the probability that another molecule of SDS will bind to that protein) and alters most proteins into rigid rods whose length is proportional to molecular weight.

Nondenaturing detergents such as Triton X-100 have rigid and bulky nonpolar heads that do not penetrate into water-soluble proteins; consequently, they generally do not disrupt native interactions and structures of water-soluble proteins and do not have cooperative binding properties. The main effect of nondenaturing detergents is to associate with hydrophobic parts of membrane proteins, thereby conferring miscibility to them.

At concentrations below the CMC, detergent monomers bind to water-soluble proteins. Above the CMC, binding of detergent to proteins competes with the self association of detergent molecules into micelles. Consequently, there is effectively no increase in protein-bound detergent monomers with increasing detergent concentration beyond the CMC.

Detergent monomers solubilize membrane proteins by partitioning into the membrane bilayer. With increasing amounts of detergents, membranes undergo various stages of solubilization. The initial stage is lysis or rupture of the membrane. At detergent:membrane lipid molar ratios of 0.1:1 through 1:1 the lipid bilayer usually remains intact but selective extraction of some membrane proteins occurs. Increasing the ratio to 2:1, solubilization of the membrane occurs resulting in mixed micelles. These include phospholipid-detergent micelles, detergent-protein micelles, and lipid-detergent-protein micelles. At a ratio of 10:1, all native membrane lipid:protein interactions are effectively exchanged for detergent:protein interactions.

The amount of detergent needed for optimal protein extraction depends on the CMC, aggregation number, temperature and nature of the membrane and the detergent. The solubilization buffer should contain sufficient detergent to provide greater than 1 micelle per membrane protein molecule to ensure that individual protein molecules are isolated in separate micelles."

I know plenty of people who only add supplements (like FBS, A/A, etc.) without removing any of the base media (DMEM, a-MEM, RPMI, etc). The correct way to do it would be to proceed like you are making a buffer, starting with 60-80% volume of base media (like with water for buffers), add any supplements like 5-20% FBS, antibiotics, antimycotics, etc., then q.s. to final volume with your base media.

The difference in the concentration of serum you are using is very minor, I would not worry about it. If you want to be absolutely sure you would have to repeat your experiments with the correct concentration, of course. The important thing right now for your experiments, if you are in the middle of one, is to be consistent -- make it the exact same way each time throughout the entire experiment so if it does have an effect, it won't be a variable for analyzing your experimental results.