SEQanswers

Go Back   SEQanswers > Applications Forums > Sample Prep / Library Generation



Similar Threads
Thread Thread Starter Forum Replies Last Post
SMART Seq v2: RNA amount and cycles for preampfification candida utilis Sample Prep / Library Generation 0 05-07-2017 11:56 PM
Smart-Seq v4 ultra low RNA kit seqgirl123 Sample Prep / Library Generation 4 03-16-2017 02:39 PM
2x150 cycles vs 2x250 cycles: coverage, quality, aligning and cost issues seqask Illumina/Solexa 3 08-31-2013 07:27 AM
Has any one here used 200 cycles SBS kit to perform reads shorter than 50 cycles? Lanlan Illumina/Solexa 2 02-27-2012 08:54 PM

Reply
 
Thread Tools
Old 05-08-2017, 12:18 AM   #1
candida utilis
Junior Member
 
Location: Taiwan

Join Date: May 2017
Posts: 5
Default SMART Seq v2: RNA amount and cycles for preampfification

Hi all,
We aim to compare differential mRNA expression between 2 different cell types. We sorted 2 populations of cells by FACS, extract RNA by column based method, and SMARTSEQ v2 for pre-amplificaiton of cDNA.

Between those cell types, A has very low cell number (150-500 cells). In contrast, B has higher cell number (50,000 cells) but its RNA amount is less. So, I only do RNA quantification for B, because RNA amount in A is too low to be detected by Ribogreen.

For Pre amplification of cDNA, I have some questions:

(1) If the starting RNA amount used for SMARTSeq V2 is different between samples, will it affect the RNA Seq results, if we aim to compare the differential mRNA expression between those samples?

(2) For the pre-amplification of cDNA, will the number of PCR cycles for A and B need to be the same, if I would like to compare the differential mRNA expression between those cell types? If sample A need 20 cycles to get sufficient cDNA, while sample B only need 15 cycles, what should I do? Thank you
candida utilis is offline   Reply With Quote
Old 06-02-2017, 08:21 AM   #2
Simone78
Senior Member
 
Location: Basel (Switzerland)

Join Date: Oct 2010
Posts: 205
Default

Quote:
Originally Posted by candida utilis View Post
Hi all,
We aim to compare differential mRNA expression between 2 different cell types. We sorted 2 populations of cells by FACS, extract RNA by column based method, and SMARTSEQ v2 for pre-amplificaiton of cDNA.

Between those cell types, A has very low cell number (150-500 cells). In contrast, B has higher cell number (50,000 cells) but its RNA amount is less. So, I only do RNA quantification for B, because RNA amount in A is too low to be detected by Ribogreen.

For Pre amplification of cDNA, I have some questions:

(1) If the starting RNA amount used for SMARTSeq V2 is different between samples, will it affect the RNA Seq results, if we aim to compare the differential mRNA expression between those samples?

(2) For the pre-amplification of cDNA, will the number of PCR cycles for A and B need to be the same, if I would like to compare the differential mRNA expression between those cell types? If sample A need 20 cycles to get sufficient cDNA, while sample B only need 15 cycles, what should I do? Thank you

Hi,
I have a different concept of what is low number of cells!
To me, 50.000 seems awfully a lot even if the amount of RNA per cell is tiny.
Running the Smart-seq2 protocol as it is might be a problem with so many cells. You will eventually run out of some reagents during the PCR (dNTPs, primers, etc), so you might need some adjustments.
Anyway, my answers to your questions are the following.
1- if you pool A and B in the same seq lane you will get different number of reads. The more "unbalanced" is your mix, the more reads you will get from one of the 2 samples compared to the other. You might end up sequencing one sample to saturation while the other will get very shallow sequencing (but thatīs an extreme case). Smaller differences are not a problem. Even relatively large differences can be managed bioinformatically, i.e. downsampling.
2- no, it doesnīt need to be the same. I personally believe that 15 or 20 cycles doesnīt matter that much in terms of the bias you introduce. A lot of transcripts, especially the rare ones, are probably lost in the first cycles anyway due to poor processivity of the polymerase, secondary structure that delay or block the polymerase, etc. My suggestion is to amplify your cDNA in order to get enough cDNA for the following steps and not more. Over-amplification increases only the PCR duplicates and doesnīt give you more in terms of total number of trascripts.

I hope I answred your questions!
/Simone
Simone78 is offline   Reply With Quote
Old 06-02-2017, 09:35 AM   #3
candida utilis
Junior Member
 
Location: Taiwan

Join Date: May 2017
Posts: 5
Default

Hi Simone, thanks for reply. But I think my words mislead you. I didn't use all RNA for both samples. Instead, for 50,000 cells, I only use 3ng RNA with 15 pre-amplification cycles, no matter how much RNA I obtain. For 150-500 cells, I use all RNA with 25 pre-amplification cycles. After this step, for both samples, I put equal cDNA amount for library preparation and pool equal amount of library in a lane for sequencing. But, I am wondering, if our aim is to compare differential expression of "certain novel transcripts" between 2 samples, will it be meaningful if starting with different RNA input and different pre-amplification cycles ?? (eg for certain novel transcript, in sample B, it is amplified with 15 cycles, while in sample A, it is amplified with 25 cycles). Thank you.

Last edited by candida utilis; 06-02-2017 at 09:58 AM.
candida utilis is offline   Reply With Quote
Old 06-02-2017, 09:43 AM   #4
minukk
Junior Member
 
Location: Arizona

Join Date: Jun 2015
Posts: 2
Default

Hi,
Is it not possible to use same cell numbers for both? Approx.~ 400-500 cells? You should be able to get RNAseq data from single cells so I would assume that 500 cells is plenty for Smartseq2 protocol.
minukk is offline   Reply With Quote
Old 06-02-2017, 01:35 PM   #5
Simone78
Senior Member
 
Location: Basel (Switzerland)

Join Date: Oct 2010
Posts: 205
Default

I agree with minukk. Since you have a lot from one of the 2 samples why not taking comparable amounts? Even with single immune cells (T, B, ILC, etc) we never have to do more than 22-23 cycles and even then we get out several ng of cDNA. Isnīt 25 cycles a bit too much for 150-500 cells?
But I agree, a difference of 10 cycles between the 2 samples might distort the whole picture and create artefacts. So I wouldnīt do it.
Simone78 is offline   Reply With Quote
Old 06-10-2017, 08:20 PM   #6
candida utilis
Junior Member
 
Location: Taiwan

Join Date: May 2017
Posts: 5
Default

Hi Minukk, thanks for your reply. We didn't use Minukk's strategy because the RNA amount between cell types is quiet different. But we can think about this strategy.

Hi Simone78, thanks for your reply. But I am still not very clear.

(1) If we aim to compare expression of novel transcripts between samples, does the pre-amplification cycles between samples need to be the same? May I know why, in the first time, you said 15 and 20 cycles doesn't matter much, but then you said 10 cycles between 2 samples might create artifacts? How many cycles differences do you think to be acceptable without introducing too much bias? Could you explain a bit more for the reasons?

(2) I have two RNA samples: One is pure RNA extracted by Picopure; the other is cell lysate in lysis buffere without purification). If we do SMART Seq2 for those samples, will the RNA Seq results be comparable? ( We aim to compare expression level of novel transcripts between 2 samples)

Thanks again!
candida utilis is offline   Reply With Quote
Old 06-11-2017, 05:01 AM   #7
Simone78
Senior Member
 
Location: Basel (Switzerland)

Join Date: Oct 2010
Posts: 205
Smile

Quote:
Originally Posted by candida utilis View Post
Hi Minukk, thanks for your reply. We didn't use Minukk's strategy because the RNA amount between cell types is quiet different. But we can think about this strategy.

Hi Simone78, thanks for your reply. But I am still not very clear.

(1) If we aim to compare expression of novel transcripts between samples, does the pre-amplification cycles between samples need to be the same? May I know why, in the first time, you said 15 and 20 cycles doesn't matter much, but then you said 10 cycles between 2 samples might create artifacts? How many cycles differences do you think to be acceptable without introducing too much bias? Could you explain a bit more for the reasons?

(2) I have two RNA samples: One is pure RNA extracted by Picopure; the other is cell lysate in lysis buffere without purification). If we do SMART Seq2 for those samples, will the RNA Seq results be comparable? ( We aim to compare expression level of novel transcripts between 2 samples)

Thanks again!
Hi, I agree my answers might have been a bit confusing!
I donīt think anybody know for sure, simply because I believe nobody has done a systematic test with one sample and try to increase the number of PCR cycles more and more. What we noticed is that, if one starts from single cells, the more PCR cycles we did and the more transcripts we "lost". Or, put it another way, with 6 or 8 cycles of PCR we could detect more genes than 18, for example. Again, I donīt know what would happen if you start doing 28 cycles when only 18 would be needed (just an example). Based on our small test, I would say that the largest bias has already been introduced in the first cycles, so it shouldnīt matter. On the other hand, I would think that over-amplifying A LOT (and here I mean 10 cycles or more) is not good either. Unfortunately I donīt have data to support my claims.

Regarding question 2, there will be a difference for sure due to the fact that the Rt is taking place in a sub-optimal environment when performed on a cell lysate. How much this will affect rare transcript is a thing that you have to test. I donīt think nobody know for sure for the simple reason that a rare transcript might not be observed for 3 reasons:
- it was not there in the first place, since the cell didn't express that specific gene at that specific time point (differences in cell cycle, stress, etc).
- it was there but the sub-otpimal conditions of RT caused its loss.
- it was not there but, although conditions were optimal (cell lysate, for example) it was lost due to the intrinsic limitations of the enzyme used. remember that the sensitivity of STRT or SMART-seq2 is about 35-40%, which means that most of the transcripts are actually lost in the beginning.
In my opinion, since there 3 factors are at play simultaneously there is no way to tell them apart. I believe that the samples might be comparable but I canīt be entirely sure.
Sorry for not being more helpful!
/Simone
Simone78 is offline   Reply With Quote
Reply

Thread Tools

Posting Rules
You may not post new threads
You may not post replies
You may not post attachments
You may not edit your posts

BB code is On
Smilies are On
[IMG] code is On
HTML code is Off




All times are GMT -8. The time now is 11:38 AM.


Powered by vBulletin® Version 3.8.9
Copyright ©2000 - 2020, vBulletin Solutions, Inc.
Single Sign On provided by vBSSO