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Quantitating Newly Ordered DNA Strands

Provided by Damien Woods

This is a 2-step protocol where we first try to set the DNA concentration to be very roughly 100 μM, and then adjust to bring it close to 50 μM. For a faster 1-step protocol you could easily adjust the calculations and go straight for 50 μM.

The protocol assumes we will dissolve in 1x TE, some people use Milli Q H20 instead.

1. Switch on biophotometer. Prepare bench, lay out the things you'll need (for consistency, do this the same way every time you do the protocol).

2. Spin down lyophilized (dried) DNA to make sure there are no flakes on/near lid.

3. First, shoot for 200 μM: If IDT claim to give us x nanomoles, we add 5x μL 1x TE, to the dried DNA, to get roughly 200μM, explained as follows:

 \frac{x \times 10^{-9}}{5x \times 10^{-6}} \quad \frac{moles}{litre}  \quad
=\quad \frac{2x \times 10^{-9}}{x \times 10^{-5}} \quad  \frac{moles}{litre}
=\quad 2\times 10^{-4} \quad \frac{moles}{litre}

\quad=\quad 200 \times 10^{-6} \quad \frac{moles}{litre}
=\quad 200 \quad \frac{micro moles}{litre} \quad = \quad 200 \, \mu M

4. Vortex (mix) for a few minutes. Spin down using microcentrifuge to remove any droplets form the lid/top of test tube. Vortex again. Spin. It is important to get the DNA completely into solution, so mix well. Finish by spinning down.

5. Put 98 μL of solvent (1X TE buffer) into a plastic cuvette. Place in biophotometer with (upwards pointing) triangle facing you. Push down in a specific pattern (top-left-bottom-right, then bottom-left-top-right, then top left). (Insert the cuvette in exactly same direction, and using the same technique, every time.) Set large pipettor to one side (do not let the tip touch anything, I leave it over the side of the bench).

6. Hit "blank".

7. Add 2 μL of DNA. Mix with small pipettor (suck up and down). Mix with large pipettor (suck up and down very carefully). Try not to introduce bubbles, nor splash on side of pipettor.

8. Insert cuvette, using the same technique as before. Hit "sample", and record the value.

9. Mix with large pipettor (stir, while in biophotometer), record value.

10. Mix with large pipettor (remove from biophotometer, suck up and down), record value. Discard cuvette.

11. If the 3 values are close to each other (within a few percent), then record the first one. If not, try again.

12. Make a note of the fact you have removed 2 μL from the DNA.

13. Repeat at least 3 times, by going back to step 5 on each iteration. Usually I do 3 to 5 samples. If the results are not consistent, re-mix, and do more. Let A260 be the average of all samples. (If there is one outlier I ignore it.)

14. Use the following formula to find the concentration: []μM = dilution-factor \times \frac{A_{260}}{\epsilon} \times 10^6

Here dilution-factor = 50 (i.e. you put 2 \, \muL of DNA into 98 \, \muL 1x TE), A260 is the absorbance value, and ε is the extinction coefficient of your oligo usually given in /M/cm (supplied by IDT with your oligo, or else use the in-house DNA group code for double stranded complexes). This comes from: A_{260} = C \times L \times \epsilon where C is the concentration, L is the path length (1 cm for the Biophotometer). And this in turn comes from the Beer-Lambert Law. See IDT's Page

15. Shoot for 100 μM. We will calculate how much 1x TE to add in order to get as close as possible to 100 μM. To get the current volume of DNA in solution, subtract 2y \, \muL from the initial volume (5x \, \muL), where y is the number of DNA samples you took. Then apply the following formula to find out how much 1x TE to add to get the desired concentration (100 μM in our case):

 \textrm{volume\_to\_add} \, (\mu L) = \frac{\textrm{current\_concentration} \, (\mu M) \times \textrm{current\_volume} \, (\mu L)  }{\textrm{desired\_concentration} \, (\mu M)}  \,\, - \,\, \textrm{current\_volume} \, (\mu L) 

16. Add  \textrm{volume\_to\_add} \, (\mu \textrm{L}) to the solution. Mix, spin, mix spin.

17. Do steps 4-14 with the new sample. If everything went well, you should get a value close to 100 μM.

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