Barrick Lab :: QPCRToQuantifyPlasmidCopyNumber

---+ Absolute QPCR for quantification of plasmid copy number in _E. coli_

This protocol is based on methods described in Lee _et al_ (2006), [[http://www.sciencedirect.com/science/article/pii/S0168165605007509][link to paper]].

---++ Designing primers for qPCR

Design your primers as outlined [[http://barricklab.org/twiki/bin/view/Lab/QPCR][here]]

---++ Overview

You will determine plasmid copy number by:

   * <span style="background-color: transparent">creating a standard curve for gDNA copy number (copy number vs cycle threshold) by loading known amounts of gDNA into a qPCR reaction.</span>
   * <span style="background-color: transparent">creating a standard curve for plasmid number (number of copies vs cycle threshold) by l</span>oading known amounts of plasmid DNA into a qPCR reaction.
   * determining the number of copies of plasmid and genome in your experimental samples, and consequently, the number of plasmids/genome ("copy number").

---++ Preparation of gDNA and plasmid DNA for creating standard curves.

   * Grow overnight cultures of:
      * Your experimental strain, harboring the plasmid the copy number of which you wish to determine.
      * Your experimental strain, however, without the plasmid.
   * Dilute your saturated cultures 1:100 into fresh media and let grow for 2-3 hours until the cells reach a mid-exponential phase (OD<sub>600</sub><span style="background-color: transparent"> = ~0.4-0.6)</span>
   * For each sample, pellet 1 ml of cells for 5 minutes at 3,000 RPM
   * For gDNA template; perform a genomic DNA extraction on the strains do NOT harbor your plasmid.
   * For plasmid DNA template: perform a mini prep on strains that DO harbor the plasmid.

---++ QPCR using SYBR Green I dye, Part 1: Setting up a plate

   1 Set up standard curves for your gDNA and plasmid samples, these will also help you calculate your primer efficiencies (should be between 0.8-1.1) 
      * To generate standard curves, dilute your gDNA and plasmid templates in ten-fold increments
      * A total of 7 dilutions is enough to make a good standard curve, your CT values should be between 5-30
   1 Normalize your sample templates to 2ng/µL 
      * Make sure to dilute these so the concentrations fall within the range of the standard curves
   1 Once your DNA templates have all been diluted, you can being to set up a 96- or 384- well plate to run your qPCR experiment 
      * For 96-well plates the reaction volumes are as follows:

%TABLE{dataalign="center"}%
| *Template* | *10µM F primer* | *10µM R primer* | *SYBR Green PCR Mix* | *ddH<sub>2</sub>O* |
| X | 0.75 | 0.75 | 7.5 | template - X |

   1 Run your qPCR plate using the following cycling conditions: 
      * Step 1 = Hold Stage 
         1 50°C - 02:00
         1 95°C - 10:00
      * Step 2 = PCR Stage 
         1 95°C - 00:15
         1 54°C - 01:00
         1 Go to step 2-1, 40X
      * Step 3 = Melt Curve 
         1 95°C - 00:15
         1 54°C - 01:00
         1 95°C - 00:15

---++ QPCR using SYBR Green I dye, Part 2: Analyzing your data

   1 Calculate the primer efficiencies by plotting CT vs DNA concentration for the gDNA and plasmid standards, and using [[https://www.thermofisher.com/us/en/home/brands/thermo-scientific/molecular-biology/molecular-biology-learning-center/molecular-biology-resource-library/thermo-scientific-web-tools/qpcr-efficiency-calculator.html][this website]]
   1 If your efficiencies are between 0.8-1.1, then calculate the ratio of plasmid:gDNA for each of your samples

-- Main.DaciaLeon - 15 Dec 2016
This topic: Lab > WebHome > ProtocolList > QPCRToQuantifyPlasmidCopyNumber
Topic revision: r5 - 2017-05-25 - SimonDAlton