QuantSeq Library Prep test-run

Katherine suggested I work through the library prep protocol with a few samples to practice and work out kinks. From her experience, the libraries she generated later in the game were of higher quaity. I’m generating 8 test libraries - this was her recommendation based on the heavy use of 8-channel pipettes.

Following the QuantSeq manual and tips from Katherine.

Quick reference for Libary Generation


Notes from this test run:

Started on 8/20/2019

Prior to beginning, I created programs on the PTC-200 DNA Engine Cycler. Programs are labeled “Quantseq-#”, with the # corresponding to step number in the QuantSeq manual.

first strand cDNA synthesis

  • Based on Katherine’s suggestions, I will generate 40 libraries at one, in 5 rows of 8 on a PCR plate. When adding solutions to each well, rather than using a single channel pipette and transferring solutions to individual wells, I’m going to use PCR strips, load enough of each solution into 8 wells (with a bit excess), and use a multichannel to distribute. This way I can hopefuly reduce time.
  • Between each step I need to quickly “spin down” my PCR plate. Our benchtop centrifuge has a minimum run time of 1 minute, and with the time it takes to accelerate and decelerate total time is ~4 minutes. The “quickly spin down” instructions do not define a centrifuge speed to use, so I set it to 3 rcf based on whatt is specified in the equipmentt list - “Benchtop centrifuge (3,000 x g, rotor compatible with 96-well plates”. To reduce the amount of time to spin down plates, I should either set the speed lower, OR simply start, then stop the centrifuge manually.

RNA removal

  • No notes.

second strand cDNA synthesis

  • SS1 and USS are indeed viscous; hold pipette in place when pulling volumes.

Placed in -20C overnight.


(this is 1 of 2 purifications, dubbed “pre-PCR”)

  • As Katherine hinted, it’s important to have a magnetic plate that fits the PCR plates used. The plates we have just have rods - no wells for plates to hold plates in place, which doesn’t work. I ordered a magnet/plate from ebay to hopefully improve the process.

Placed in -20 on 8/21/19 until next step (qPCR assay).

qPCR assay for optimal # cycles

Performed on 9/3/2019

  • Created a custom qPCR protocol with Sam’s help -
qPCR Assay Mastermis Calcs      
Item per rxn (uL) all rxns (inc. NTC) (uL) all rxns * 1.1
Number of samples 1 9  
cDNA, diluted to 19uL 1.7 15.3 16.8
PCR mix (PCR) 7 63 69.3
P7 Primer (7000) 5 45 49.5
Enzyme mix (E) 1 9 9.9
2.5x SYBR Green I nucleic acid dye 1.2 10.8 11.9
Elution Buffer (EB) 14.1 126.9 139.6
Mastermix total vol 28.3 254.7 280.2
SYBR Green Calcs Volumes    
Stock concentration 100    
Desired concentratino 2.5    
Total volume needed 2.5x 11.9    
Volume 100x 0.2970    
Volume DMSO 11.58    
Dilution ratio (should be 1:40) 0.025    
Final concentration 2.5    

Results: Located on Owl, https://owl.fish.washington.edu/scaphapoda/qPCR_data/cfx_connect_data/ with today’s date.

No amplification :/ max RFU should be ~10x10^12, and my no-template control (NTC) has same curve as samples.

2019-09-03_ qPCR-assay-test-run

First troubleshooting step is to see if I actually synthesized cDNA - I believe I can use the Qubit for that. If yes, then I messed up the qPCR somehow (wouldn’t surprise me). Perhaps there is an issue with bubbles? Maybe BioRad settings needed be adjusted for sybr green?

I ordered a trial QuantSeq kit (n=4) for trouble shooting, and as a result spoke with the WA respresentative. Notes from our call:

  • If cDNA synthesis did not occur, then I most likely had contamination with organics or salts, which can inhibit 1st strand synthesis and cause cDNA to degrade when trying to degrade RNA.
  • The TurboDNase method might be an issue, since it did not include a column.
  • Using a cleaner column on existing RNA may do the trick. I did order one box of Zymo Cleaner-Concentrator (n=50), so could run all my samples through this column.
  • 500 ng of input RNA is not necesary (she said that “no one uses that much”). 100 ng should be adequate!

I will receive the trial kit tomorrow, and Kristy is connecting me with the tech support guy.

Update on qPCR Assay, 9/5/2019

Well, I’m an idiot, but in this case it’s a good thing. When preparing my plate for the qPCR assay I pulled my “samples” from the incorrect well, so loaded blank buffer (i.e. all NTCs) instead of samples - not surprising that I didn’t get the expected results - i.e. amplification at about 15 cycles. It is weird that there was any amplification at all, suggesting possible contamination? I re-did the qPCR assay step today. Before realizing my qPCR mistake, I also tried to quantify my ds cDNA libraries.

  • Added 3uL elution buffer to all wells
  • Quantified ds cDNA using the Qubit 1x dsDNA assay, and 1 uL from each of my samples.
    • Results for all samples - TOO LOW. Not sure if this is expected…
  • Prepared mastermix for qPCR assay just like before (calcs above), with 2 exceptsion:
    • Prepared twice the amount of sybr green working solutiom - 1 uL sybr green stock + 39 uL DMSO.
    • Loaded PCR plate in the AirClean PCR workstation to minimize airborn contamination.
    • Spun PCR plate down after preparing to remove bubbles. This was harder than anticipated - the ~3,000 rcf speed did not remove bubbles, so I went as high as the benchtop centrifuge would allow - 4,000 for 3 minutes. According to the centrifuge manual it should go as high as ~10,000 rcf, so not sure why I kept getting an error message at speeds above 4k.

Results: 7 out of 8 samples started to amplifly at the expected cycle! One did not, producing an amplification curve similar to the NTC.

Well Fluor Content Sample End RFU
A04 SYBR Unkn 296 3531
A05 SYBR Unkn 323 3829
A06 SYBR Unkn 341 3848
A07 SYBR Unkn 403 3850
B04 SYBR Unkn 472B 3118
B05 SYBR Unkn 490 3253
B06 SYBR Unkn 513 3735
B07 SYBR Unkn 531 3784


To calculate the optimal number of cycles for endpoint PCR when amplifying/adding adapters to my ds cDNA I do the following:

  • Determine endpoint RFU, aka RFU value at plateau
  • Calculate 50% of endpoint RFU
  • Identify cycle # at 50% endpoing RFU
  • Subtract 3 cycles

Katherine advises that I should round down if the calculated cycle number is a decimal, to avoid overcycling.


Cohort Treatment Tissue RNA Sample no. RFU @ endpoint 50% max No. Cycles @ 50% No. Cycles @ 50% minus 3 cycles Cycles, round down
Dabob Bay 6 Low High Ctenidia 296 3530 1,765 16.97 13.97 14
Oyster Bay 6 Ambient Ctenidia 323 3829 1,915 17.52 14.52 14
Fidalgo Bay 6 Ambient Ctenidia 341 3848 1,924 17.59 14.59 14
Dabob Bay 10 Ambient Larvae 403 3850 1,925 18.38 15.38 15
Fidalgo Bay 6 Low Larvae 472b 3118 1,559 17.27 14.27 14
Oyster Bay C1 10 Ambient Larvae 490 3253 1,627 18.81 15.81 15
Oyster Bay C1 6 Ambient Larvae 513 3735 1,868 29.17 26.17 NA
Oyster Bay C2 10 Ambient Larvae 531 3784 1,892 16.68 13.68 13
      NTC 3840 1,920 28.20 25.20 NA
Written on September 3, 2019