PCR

Quick Reference
DNA concentrations lower than 100 ng/ul
Use the C1V1 = C2V2 calculation

• C1 = Concentration of DNA measured (ng/ul)
• V1 = The total amount of elution buffer (EB) you used
• C2 = 20 ng/ul this is the concentration we want
• V2 = The total volume we will have after diluting

• V2 = C1V1
• C2
• V2 = Total Volume
• V1 – V2 = the amount of liquid you will add **

 

** To find out how much molecular grade water we need to add we subtract the amount of EB we used from Final volume. There should be enough room to add the molecular grade water in the microfuge tube holding your DNA sample. If not use the calculation below.

For DNA concentrations higher than 100 ng/ul
Same calculation C1V1 = C2V2

• C1 = Concentration of DNA measured (ng/ul)
• V1 = 2 ul of Extracted DNA
• C2 = 20 ng/ul this is the concentration we want
• V2 = The total volume we will have after diluting

• V2 = C1V1
• C2
• V2 = Total Volume

 

This time to find out how much molecular grade water we need to add, we subtract the 2 ul of DNA template from the Final volume. In a new tube add 2 ul of your extracted DNA and the calculated amount of molecular grade water, label tube with lab ID and a capital D to indicate it is diluted.

Details
It is important to measure the quality and quantity of extracted DNA. This is achieved with a spectrophotometer. We use NanoDrop but there are other brands of spectrophotometers that can give us the measurements we need. Two important steps to help ensure you get the best measurements possible from NanoDrop, clean both pedestals well before using and make sure you blank with the same liquid the DNA is suspended in.

There are two important numbers we need to assess the quality of the extraction 260/280 nm and 260/230 nm ratio. The ideal 260/280 ratio for DNA is 1.8 nm and for RNA is 2.0 nm. If the value is less than the ideal numbers, there may be a phenol, protein, or another type of contaminate that absorbs at 280 nm. If you are measuring DNA and it is higher than 1.8 nm then you may have some RNA in your sample. The pH of your sample and the composition of base pairs can also affect the absorption.

The ideal 260/230 ratio is 2.0 but acceptable range may be 1.8 nm to 2.2 nm. Sometimes chemicals used in the extraction process are not completely removed or there are carbohydrates that escaped the purification process can lower this ratio.

Keep in mind, what you intend to do with the DNA that is extracted. You might still get a good enough quality PCR to begin the molecular identification process but not other good enough for other molecular methods. If the quantity of DNA is close to 1 ng/ul you might not get an accurate reading on some machines.

Once you have all three numbers and you are ready to move forward with running a PCR you will likely need to dilute your DNA. I find PCR is more successful when my DNA concentration is between 20 and 5 ng/ul.

Example Calculations
After using a spectrophotometer you find the concentration of two different samples are 30 ng/ul and 224 ng/ul. To dilute both samples to about 20 ng/ul we will use two different approaches.

C1 = 30 ng/ul (what we have)
V1 = 198 ul (current volume of DNA extraction after NanoDrop)
C2 = 20 ng/ul (target concentration)
V2 = How much molecular grade or sterile water do we need to add to dilute the sample to 20 ng/ul

• V2 = C1V1
• C2
• V2 = Total Volume

• V2 = 30 ng/ul (198 ul)
• 20 ng/ul
• V2 = 297 ul

 

Now we know the final volume will be 297 ul.
To find out how much water we need to add we subtract 198 ul from 297 ul.

•  297 ul
• -198 ul
•    99 ul of water to dilute our sample

 

This one was easy because we just needed to add 99 ul of sterile water to dilute our sample and the end amount does not exceed what the microfuge tube can hold.

Lets approach the next sample the same way
C1 = 224 ng/ul is what we have
V1 = 198 ul is the volume we have
C2 = 20 ng/ul is the concentration we want
V2 = How much water will it take?

• V2 = C1V1
• C2
• V2 = Total Volume

Solve for V2

• V2 = 224 ng/ul (198 ul)
• 20 ng/ul
• V2 = 2,216.6 ul

So we can see right away we are exceeding the volume of most microfuge tubes (1.7 ml). Therefore we need a different approach. We could pipette 10 ul of our DNA into a new tube and recalculate how much water we would need to dilute our DNA.

C1 = 30 ng/ul
V1 = 10 ul
C2 = 20 ng/ul
V2 = The total volume we will have after diluting

Solve for V2

• V2 = 30 ng/ul(10 ul)
• 20 ng/ul
• V2 = 15 ul total volume

•  15 ul
• -10 ul
•    5 ul of water to dilute our sample

Quick Reference

• Spin tubes 30 – 60 sec
• Multiply primer’s nmol by 10 ul
• Hydrate with sterile 1x TAE (or molecular grade water)
• Incubate at RT for minimum of 10 mins. Mix well before use

Steps
Before hydrating primers, spin tubes in centrifuge for 30 to 60 seconds to draw all the powder down to the bottom of the tube. During shipping the powdered primers spread all over the inside of the tube and if you do not centrifuge the tubes you risk losing some and therefore end up with an unknown concentration.

Now that you are ready to hydrate, you need to find the concentration of the powdered primers. You can find the primers the nanomole (nmol) concentration of the powder listed on the paperwork that came with your primers or you can find it on the tube’s label. To make a 100x Master Stock multiply the primer’s nmol by 10 ul of molecular grade water or sterilized 1x TAE solution. Let the liquid soak in for a minimum of 10 mins then vortex to mix well. I prefer to let newly hydrated primers sit overnight when possible.

1x TAE solution is best to use because it will protect your primers from shearing caused by thaw and freeze cycles your tubes go through every time you use your master stock. However, some find water more convenient and either use the master stock quickly or do not mind re-ordering primers when they become less efficient.

For example
Primer concentration is listed as 23.4 nmol. When multiply 23.4 by 10 the answer is 234. Therefore add 234 ul of sterile water or 1x TAE to hydrate primers.

Quick Reference

• Vortex primer master stock for 30 sec
• Dilute 10:1 (90 ul of water to 10 ul of Master stock)
• Vortex working stock for 30 sec before using

Steps
Vortex to mix the 100x primer master stock before making the 10x working stock. This is a 1:10 dilution. Simply breaks down to 90 ul of molecular grade water to 10 ul of master stock. Properly label your working stock with primer and concentration.

Quick Reference
1x Master Mix

• 22.5 ul PCR Super Mix
• 0.5 ul Forward primer
• 0.5 ul Reverse primer

To PCR tube add

• 1.5 ul Template DNA
• 23.5 ul Master Mix

Steps
Using Supermix from Thermo Fisher Scientific catalog number 10572014. This kit is designed for a total volume of 25 ul per PCR tube.

Before starting, measure your DNA using nanodrop. Your DNA template should be somewhere between 20 – 5 ng/ul. If you have trouble remembering how to dilute your DNA, see the dilution protocol.

If your sample is at 5 ul or slightly less then you should adjust your master mix to have less Super mix, primers and increase the amount of template DNA. Just be sure these adjustments still give a total volume of 25ul per tube.

When you are calculating how much master mix to make don’t forget to add one for control and one for variations in pipetting.

Quick Reference
Big gels (Owl, 20 wells) reagents
1.2 g Agarose
120 ml TAE 1x
5 ul SYBR safe

Small gels (5-10 wells)
0.6 g Agarose
60 ml TAE 1x
2.5 ul SYBR safe

Loading samples into the gel
5 ul PCR product
1 ul Loading dye

Steps
To 250 ml Erlenmeyer flask add TAE 1x and agarose
Microwave for 30 – 40 seconds, stop before boiling occurs (glass will be hot)
Swirl to mix, you want the agarose to melt completely into solution
Once in solution, let cool to touch then add SYBR safe, swirl to mix well then pour into casting tray.

SYBR safe is safer than Ethidium bromide but is mixed with DMSO which helps chemicals crossover membranes – you should wear gloves when handling SYBR safe. SYBR safe is light sensitive therefore be quick and efficient when pipetting from tube. Close the reagent cap right away and do not leave on the counter.

After gel has solidified, rotate gel tray and add running buffer (TAE 1x). Buffer should cover the top of the gel completely. Remember, DNA is a negative molecule therefore your samples should be on the negative end so they travel down the gel towards the positive end.

Running the Gel
110 V for about 45 – 30 mins depending on buffer and gel size.
Always be sure you see tiny bubbles from the tungsten wire inside and that your dye is traveling in the correct direction.

Quick Reference
0.5 M EDTA stock (500 ml)

• 93.05 g of Disodium EDTA
• 500 ml DI water

Dissolve disodium EDTA in 400 ml DI water, adjust to pH 8.0 with NaOH. Top off solution to 500 ml with DI water.

Note EDTA is not soluble until titrated to pH 8.0. Depending on the concentration you are preparing, it may take quite a bit of NaOH to titrate EDTA into solubility. Ensure your additions don’t overshoot the required final volume!

50X TAE Stock

• 242.0 g Tris Base
• 57.1 ml Glacial Acetic Acid
• 100 ml 0.5 M Disodium EDTA (pH 8.0)
• Bring to 1 L with DI water

Add Tris Base and Disodium EDTA to 700 ml of DI water, stir until dissolved. Add Acetic Acid and then bring to 1 L with DI water. The pH but does not need to be adjusted.

1x TAE Buffer (1 liter)

• 20 ml of 50x TAE stock
• 980 ml of DI water

10x TBE Stock

• 108.0 Tris Base
• 55.0 g Boric Acid
• 40 ml 0.5 M Disodium EDTA (pH 8.0)
• Bring to 1 L with DI water

Add Tris and Boric Acid to 800 ml of DI water, stir until dissolved. Add EDTA solution then bring volume to 1 L with DI water.

1x TBE Buffer (1 liter)

• 100 ml of 10x TBE stock
• 900 ml of DI water

10x TPE Stock

• 108.0 g Tris Base
• 15.5 ml 85% Phosphoric Acid
• 40 ml 0.5 M Disodium EDTA (pH 8.0)
• Bring to 1 L with DI water

Add Tris and Phosphoric Acid to 800 ml of DI water, stir until dissolved. Add EDTA solution then bring volume to 1 L with DI water.

50X Sodium Borate (SB)

• 20 g NaOH
• 112.5 g Boric acid
• Adjust to pH 8.0
• Bring to 1 L with warm DI water

Heat 800 ml water to near boiling then add boric acid adjust pH with NaOH. Once in solution, bring to 1 L.

Note: This will precipitate over time, reheat to bring back to solution

When to use each buffer
TAE is best to use when DNA is larger than 12 kb – acetate improves the separation of larger DNA fragments. This is also a great buffer if you are going to recover DNA from the gel. TAE has a lower buffering capacity but double stranded DNA fragments run faster in it than TBE.

TBE gives the best resolution of smaller DNA fragments 2 kb or less. It is more conductive than TAE and the gel stays cooler on longer runs. Has a high buffering capacity.

TPE very similar to TAE with the ability to give good separation of slightly smaller fragments than TAE. Its high buffering capacity makes it good for long runs. TPE is used in analysis of single stranded DNA. You can recovering DNA but possible interference with phosphate sensitive reactions may change your mind.

SB is unique in allowing for high voltage (300 V) fast runs (six to ten minutes) but this is only for DNA fragments 500 bp or smaller.

Further information
During electrophoresis, protons are generated at the anode and hydroxyl ions at the cathode. Thus a pH buffered system is critical to maintain the pH of the system. At a neutral pH range the buffers ensure the phosphate groups of DNA and RNA are charged and will migrate towards the anode.

T = Tris or Trizma is a buffer to maintain pH of the solution.
A = Acetic acid
B = Boric acid
P = Phosphoric acid
E = Ethylenediaminetetraacetic acid (EDTA) is a chelator of divalent cations. Specifically Fe2+, Ca2+ and Mg2+. It is used to remove such metals from solution which are important for the activity of most DNA involved enzyme reactions and limit metal-induced oxidation during electrophoresis. In other words, it inhibits DNase and RNase activity.

DNA Gel-loading Dye (10 ml)

• 0.025 g Bromophenol blue
• 0.025 g Xylene cyanol
• 200 ul 0.5 M EDTA
• 500 ul 10% (w/v) SDS
• 3.9 ml Glycerol

Bring to total volume of 10 ml with DI water