Daniel:Notebook/PosSequencing/2017-6-21: Difference between revisions

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==Discussion==
==Discussion==


<math>&alpha;</math>
First I'll establish the math behind this. The equations are numbered for convenience.
 
[[Image:Equations-20170622.png]]
 
The important equations are '''3''', the idealized y=mx+b with y as number of cycles and x as log(input), and '''4''', which is '''3''' but corrected for the efficiency (e<sub>x</sub>) of reaction explicitly added. Note that efficiency should be between 0 and 1.
 
So using the above equations for sample and control cycling as a function of input.
 
From those suppositions and using the ''controls'', we can calculate the b' constant as 15.737 and m' as 11.138. From those, we can estimate the constants and efficiencies from the ''samples'' trendline:
 
[[Image:Equations2-20170622.png]]
 
 
So the '''m''' based calculations doesn't make any sense but '''b''' gives an answer of 0.497, which is reasonable. So maybe it's 50% efficient? If so, the following table will dictate how many cycles it would take to amplify given ammounts. Bolded samples are included but from data.
 
{| class="wikitable" <hiddentext>generated with [[:de:Wikipedia:Helferlein/VBA-Macro for EXCEL tableconversion]] V1.8</hiddentext>
|- style="background-color:#92CDDC;font-size:12pt;font-weight:bold" align="center"
| width="100" height="28" | Input
| width="110" | Cycles
 
|- style="font-size:12pt" align="center" valign="bottom"
| height="15" | '''1 pmol'''
| align="center" | '''92.5'''
 
|- style="background-color:#BFBFBF;font-size:12pt" align="center" valign="bottom"
| height="15" | '''500 nmol'''
| align="center" | '''87.5'''
 
|- style="font-size:12pt" align="center" valign="bottom"
| height="15" | '''100 nmol'''
| align="center" | '''111'''
 
|- style="background-color:#BFBFBF;font-size:12pt" align="center" valign="bottom"
| height="15" | 10 nmol
| align="center" | 114.0
 
|- style="font-size:12pt" align="center" valign="bottom"
| height="15" | 1 nmol
| align="center" | 119.7
 
|- style="background-color:#BFBFBF;font-size:12pt" align="center" valign="bottom"
| height="15" | 100 fmol
| align="center" | 125.4
 
|- style="font-size:12pt" align="center" valign="bottom"
| height="15" | 10 fmol
| align="center" | 131.1
 
|- style="background-color:#BFBFBF;font-size:12pt" align="center" valign="bottom"
| height="15" | 1 fmol
| align="center" | 136.8
 
|- style="font-size:12pt" align="center" valign="bottom"
| height="15" | 100 amol
| align="center" | 142.5
 
|}
 
[[Category:PosSeq]] [[Category:20170621]]

Latest revision as of 20:32, 22 June 2017

Bead qPCR Test[edit]

Back to Calendar

This experiment has 2 purposes: to determine the density of beads necessary to optically block qPCR detection (and hopefully some levels that don't) as well as the efficiency of bead-based qPCR vs aqueous. This will hopefully help me make the next experiment run smoother, specifically using digital qPCR to amplify the beads.

Protocol[edit]

  1. Template-Bead Binding; Make 4 samples and follow this protocol for each
    1. Suspend 5 uL (50 ug, ~50M beads) beads in 100 uL Dynabuffer
    2. Apply magnet for 30 sec and remove supernatant
    3. Suspend in 20 uL Dynabuffer (conc. 2.5 ug/uL)
    4. Add 5 uL 10 uM (50 pmol total, 2X excess) dual biotin oligo and 15 uL nfH2O per sample to bead solution; incubate at RT for 15 min
    5. Wash twice with 100 uL Dynabuffer
    6. Resuspend beads in 10 uL TE buffer
  2. Dilutions
    1. Make dilutions using 2 uL suspended beads with 18 uL TE for dilutions
    2. Make dilutions of 1:10, 1:100, 1:1000, and 1:10000
    3. Sample Matrix
      Beads in Sample Dilution (Power 10) Amount Added (uL) ~Amt BioU Primer (pmol)
      10 M 0 2 5 pmol
      2 M 1 4 1 pmol
      1 M 1 2 500 nmol
      0.2 M 2 4 100 nmol
      0.1 M 2 2 50 nmol
      20 K 3 4 10 nmol
      10 K 3 2 5 nmol
      1 K 4 2 1 nmol


  3. qPCR
    1. Make the following master mix
    2. Reagent 1X Added Master Mix (21.2X)
      Primer 2 (10 uM) 1 21.2
      Barcode Oligo 100 pM 1 21.2 (1)
      Beads 4 84.8
      Kapa 2X SYBR Fast Mix 20 424
      nfH2O 14 296.8 (155.4)
      Total 40 848
    3. Add 36 uL master mix to each lane
    4. Add n uL of sample and 4-n uL nfH2O to appropriate lanes according to plate layout
    5. For the "Supernatant" sample add 10 uL TE to original beads (0 dilution), mix to suspend, then pull down; take 4 uL supernatant for sample
    6. File:PlateLayout-BeadsqPCR-20170621.png
    7. Vortex and spin down after sealing wells
    8. Use the standard thermocycler program->40 cycles

Results[edit]

Discussion[edit]

First I'll establish the math behind this. The equations are numbered for convenience.

File:Equations-20170622.png

The important equations are 3, the idealized y=mx+b with y as number of cycles and x as log(input), and 4, which is 3 but corrected for the efficiency (ex) of reaction explicitly added. Note that efficiency should be between 0 and 1.

So using the above equations for sample and control cycling as a function of input.

From those suppositions and using the controls, we can calculate the b' constant as 15.737 and m' as 11.138. From those, we can estimate the constants and efficiencies from the samples trendline:

File:Equations2-20170622.png


So the m based calculations doesn't make any sense but b gives an answer of 0.497, which is reasonable. So maybe it's 50% efficient? If so, the following table will dictate how many cycles it would take to amplify given ammounts. Bolded samples are included but from data.

Input Cycles
1 pmol 92.5
500 nmol 87.5
100 nmol 111
10 nmol 114.0
1 nmol 119.7
100 fmol 125.4
10 fmol 131.1
1 fmol 136.8
100 amol 142.5