DNasing RNA, Round I
According to the QuantSeq library prep protocol, I need to ensure no DNA contamination in my RNA samples. Sam advises that all RNAzol-processed samples will definitely have some DNA contamination. So, I am using the Turbo DNase kit to clean my RNA.
July 31st - DNased RNA, qPCR to identify DNA contamination, batch 1 (n=30)
I ran a first batch of reactions (n=30) using the Turbo DNase kit. The kits we had in the -20 freezer were old (from 2014), so I walked over to the BioSciences stock room (J wing) and purchased 2 new kits (50 reactions per; I’ll need a couple more kits). NOTE: each kit is ~$145.
Turbo DNA Protocol
All reactions used 50 uL RNA, as per the manual’s example. Here were my steps:
- Labeled 2 sets of 0.5 mL microcentrifuge tubes with RNA sample names
- Transfered 1 uL of DNase into one set of tubes
- Transfered 5 uL of Turbo DNase Buffer into each tube
- Transfered 50 uL of RNA into tubes
- Mixed gently - used low setting on the vortexer.
- Incubated at 37C - used the thermocycler in FTR 209 - for 20 minutes
- Removed samples from thermocycler, added 5 uL inactivation reagent solution. Prior to pipetting this inactivation reagent, I vortexted it thoroughly.
- Incubated at room temperature for 5 minutes. Vortexed each sample briefly twice during this incubation time to keep solution mixed.
- Centrifuged samples for 90 seconds at 10,000 rcf
- Carefully transferred supernatant to fresh, labeled tubes.
- Held DNased RNA on ice, while I learned how to run qPCR
qPCR to assess DNA contamination
I will use the SsoFast enzyme, a DNA polymerase technology that performs a super fast reaction, and the BIORAD CFX Connect qPCR machine.
Protocol:
- Created mastermix for PCR reactions for a total of 64 wells. The following table shows volumes needed for 1 pCCR reaction, then volumes needed for a mastermix for 64 reactions:
per reaction | 7/31/19 | |
---|---|---|
# Samples | 1 | 30 |
# Reactions | 1.0 | 62 |
Template (RNA sample) (uL) | 1.0 | 62.0 |
Sso Fast (uL) | 10.0 | 682.0 |
Pf, 10 uM (uL) | 0.5 | 34.1 |
Pr, 10 uM (uL) | 0.5 | 34.1 |
DEPC-treated water (uL) | 8.0 | 545.6 |
Total Volume in mastermix (uL) | 20.0 | 1,357.8 |
Following Sam’s lab notebook entry, I used elongation factor primers to check for DNA contamination:
- EF1_qPCR_5’ (SRID 309) (Forward primer, Pf, SRID = 310)
- EF1_qPCR_3’ (SRID 310) (Reverse primer, Pr, SRID = 309)
These primer stocks are stored in a small fridge in FTR 213, and can be found using the primer database. The stock concentrations are 100 micromolar. I need to use a working stock at 10 uM, so I melted the stocks and diluted 15 uL of each stock in 135 uL DEPC-treated ultrapure water (150 uL total volume).
The total volume per reaction is 20 uL. After creating the mastermix, I pipetted 19 uL of mastermix into a qPCR well plate. NOTE: the type of plate is specific - it’s a white plate, and “low profile” - which is specified on the qPCR software. I then pipetted 1 uL of each sample (i.e. template), in duplicate, to the well plate. I loaded samples horizontally (A1, A2, A3 … etc.) for ease of reading data downstream. In addition to including a control sample, which has been processed alongside the other samples since homogenization (sample 571), I included a No Template Control (NTC, 1uL water added instead of a sample), and DNA isolated from Oly larvae back in March 2018 (sample 69a, RNA sample 8a) as a positive control. To seal the plate I used the clear tape-like cover, rather than the clear plastic caps. I did not vortex the well plate prior to the qPCR reaction.
I carried the well plate over to the qPCR room, loaded it onto the CFX Connect, and opened the MAESTRO software on the adjacent computer. I used the Wizard to help configure the run. Here are the steps to execute the run:
Select “User Defined”
Select Protocol: “CFX_2StepAmp_EVAGreen+Melt.prcl”
Select plate file: “QuickPlate_96 wells_sybr_white.pltd” - this ensures that all wells are measured. We don’t assign sample names prior to running, but can edit the data file after completion.
Select “Next”
Select “Save” - it will automatically save the file to Owl and filename will include the run date.
This is a screenshot immediately upon protocol initiation
I downloaded MAESTRO to my computer (Mac version), and edited the plate setup to include sample names, and color coded melt curves by sample type: GREEN is positive control (n=2); RED is NTC (n=1), and PINK is the homogenization/isolation/DNase control (n=1); BLUE are the samples.
Data and report are saved on github in the O.lurida_Stress repo.
The melt curve doesn’t look like Sam’s recent run. However, I realize that the primers were not O. lurida, but were C. gigas. I didn’t think to ask whether the primers needed to be O. lurida specific, but I’m guessing yes. I will plan to move forward with the next batch of Turbo DNase-ing, and will figure out which primers are optimal. Interestingly I did see some DNA, and a melt temperature, for the positive controls, but the fluorescence was not as high as Sam’s example. Also interesting is that my homogenization/isolation control (pink) had a weird peak, suggesting some contamination.
Here ares ome qPCR notes from Sam’s instructions:
- Keep RNA on ice while working with them, and store in -80 always.
- There are 2x SsoFast aliquots in the fridge, and also in the freezer in the “PCR supplies” box in the -20 (both in FTR 209).
- Mastermixes should be used the same day they are prepared, but can sit on ice for a few hours.
- qPCR plates can be prepared, then sealed and held in the fridge for a bit. For example, I could prepare one qPCR plate, then while it is running I can prepare another and hold it in the fridge until the machine is ready again.
- Always use the button to open/close the BioRAD CFX Connect lid - don’t manually close the lid
Quantified DNased RNA
Used Qubit HS RNA to measure RNA concentration in DNased samples. Approximate volume remaining for DNased RNA is 50 uL. I find it odd that some of my samples have more concentrated RNA after the DNasing. I will look in to that.
Date larvae collected | Cohort | Treatment | TISSUE SAMPLE # | Homo./RNA TUBE # | VOL RNAzol (mL) | MASS TISSUE (mg) | [RNA] ng/uL | Volume for DNase treatment | Amount of RNA in Dnase treatment (ug), max is 10 ug | Date Turbo Dnase treatment | [RNA] after Turbo Dnase treatment |
---|---|---|---|---|---|---|---|---|---|---|---|
5/24/17 | Dabob Bay | 10 Ambient | 14-A | 401 | 1 | 100 | 52.0 | 50 | 2.60 | 7/31/19 | 93.4 |
5/31/17 | Dabob Bay | 10 Ambient | 31-A | 402 | 1 | 10 | 140.0 | 50 | 7.00 | 7/31/19 | 114.0 |
5/26/17 | Dabob Bay | 10 Low | 23-A | 411 | 1 | 10 | 57.2 | 50 | 2.86 | 7/31/19 | 72.6 |
5/27/17 | Dabob Bay | 10 Low | 27-A | 412 | 1 | 10 | 60.8 | 50 | 3.04 | 7/31/19 | 31.2 |
6/12/17 | Dabob Bay | 6 Ambient | 59-A | 421 | 1 | 10 | 43.0 | 50 | 2.15 | 7/31/19 | 57.6 |
6/7/17 | Dabob Bay | 6 Low | 51-A | 431b | 1 | 20 | 61.2 | 50 | 3.06 | 7/31/19 | 83.0 |
6/17/17 | Dabob Bay | 6 Low | 72-A | 432 | 1 | 50 | 47.6 | 50 | 2.38 | 7/31/19 | 74.0 |
5/25/17 | Fidalgo Bay | 10 Ambient | 20-A | 441 | 1 | 70 | 46.0 | 50 | 2.30 | 7/31/19 | 16.2 |
6/3/17 | Fidalgo Bay | 10 Ambient | 38-A | 442b | 1 | 80 | 56.2 | 50 | 2.81 | 7/31/19 | 69.8 |
5/24/17 | Fidalgo Bay | 10 Low | 16-A | 451 | 1 | 70 | 68.4 | 50 | 3.42 | 7/31/19 | 68.4 |
5/24/17 | Fidalgo Bay | 10 Low | 18-A | 452b | 1 | 80 | 48.4 | 50 | 2.42 | 7/31/19 | 97.2 |
5/26/17 | Fidalgo Bay | 6 Ambient | 22-A | 461b | 1 | 100 | 54.0 | 50 | 2.70 | 7/31/19 | 84.0 |
5/29/17 | Fidalgo Bay | 6 Ambient | 29-A | 462b | 1 | 60 | 69.8 | 50 | 3.49 | 7/31/19 | 106.0 |
5/25/17 | Fidalgo Bay | 6 Low | 19-A | 471b | 1 | 100 | 71.0 | 50 | 3.55 | 7/31/19 | 108.0 |
5/26/17 | Fidalgo Bay | 6 Low | 21-A | 472b | 1 | 70 | 64.0 | 50 | 3.20 | 7/31/19 | 97.0 |
5/20/17 | Oyster Bay C1 | 10 Ambient | 02-A | 481 | 1 | 40 | 64.4 | 50 | 3.22 | 7/31/19 | 89.2 |
5/20/17 | Oyster Bay C1 | 10 Ambient | 04-A | 482 | 1 | 60 | 67.2 | 50 | 3.36 | 7/31/19 | 22.2 |
5/23/17 | Oyster Bay C1 | 10 Ambient | 09-A | 484 | 1 | 40 | 66.2 | 50 | 3.31 | 7/31/19 | 58.4 |
6/15/17 | Oyster Bay C1 | 10 Ambient | 66-A | 491 | 1 | 20 | 126.0 | 50 | 6.30 | 7/31/19 | 58.4 |
6/14/17 | Oyster Bay C1 | 10 Low | 62-A | 506 | 1 | 80 | 63.8 | 50 | 3.19 | 7/31/19 | 29.2 |
6/5/17 | Oyster Bay C1 | 6 Ambient | 45-A | 513 | 30 | 156.0 | 50 | 7.80 | 7/31/19 | 142.0 | |
5/21/17 | Oyster Bay C1 | 6 Low | 01-A | 521 | 1 | 70 | 54.4 | 50 | 2.72 | 7/31/19 | 66.6 |
5/22/17 | Oyster Bay C1 | 6 Low | 07-A | 522 | 1 | 20 | 60.8 | 50 | 3.04 | 7/31/19 | 32.2 |
6/15/17 | Oyster Bay C1 | 6 Low | 68-A | 528 | 1 | 30 | 162.0 | 50 | 8.10 | 7/31/19 | 87.6 |
5/24/17 | Oyster Bay C2 | 10 Ambient | 17-A | 531 | 1 | 60 | 88.2 | 50 | 4.41 | 7/31/19 | 95.4 |
5/23/17 | Oyster Bay C2 | 10 Low | 12-A | 541 | 1 | 40 | 45.6 | 50 | 2.28 | 7/31/19 | 44.4 |
5/24/17 | Oyster Bay C2 | 10 Low | 13-A | 542 | 1 | 30 | 82.0 | 50 | 4.10 | 7/31/19 | 32.8 |
6/3/17 | Oyster Bay C2 | 6 Ambient | 41-A | 552b | 1 | 80 | 64.8 | 50 | 3.24 | 7/31/19 | 74.6 |
5/21/17 | Oyster Bay C2 | 6 Low | 05-A | 561 | 1 | 40 | 43.4 | 50 | 2.17 | 7/31/19 | 28.0 |
NA | RNA Control | RNA Control | 571 | 1 | 10 | LOW | 50 | LOW | 7/31/19 | LOW |