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Old 11-17-2014, 12:52 AM   #11
bunce
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Hi nucacidhunter, I guess we will agree to disagree on some of these points. I can't conceive of a way in which a 2-step protocol can be 'cleaner' or less susceptible to artefacts than a 1-step set-up. It may be cheaper, arguably more efficient? - but not cleaner. When it comes to NGS, PCR is a necessary evil it should be minimised in anyway possible.

Quote:
Originally Posted by nucacidhunter View Post
Chimeric reads actually would be more prevalent in one step PCR rather than two step, because two major reasons for Chimera formation during PCR are incomplete extension and strand invasion. The condition for both are favoured in one step PCR where the reagents are most likely to deplete and concentration of amplicons increase to a critical point favouring those reactions.
Your logic is lost on me here as the 2-step method also goes though a 1st round PCR that is just is susceptible to Chimeras (I think we agree that PCR cycles should be kept to a minimum). In amplicon sequencing some people work indexes into their 1st round PCRs then pool and amplify (as a pool) to get p5-rd1/rd2-p7 onto the products. In these situations there has been a number of reports of 'jumping' indexes presumably the result of incomplete extension. This "can't" happen (i.e. index jumping) in a 1-step workflow as the forward and reverse indexes are the only ones in tube.

Quote:
Originally Posted by nucacidhunter View Post
By using indexed primers for second step from a plate sealed with a plastic film and frozen, thawed and resealed multiple times, I have got 1 in 10,000 reads that had indices not used in the reactions. By using individual tubes containing an indexed primer and opening them one at a time such as recommended in Nextera protocol, no read was detected that had unused indexed primer.
If a 1/10,000 contamination rate suits your application then that is good - it won't suit everyone. People not as adept at removing those pesky films may report a higher rate????

Quote:
Originally Posted by nucacidhunter View Post
One obvious advantage using two step PCR with dual indexing (such as one described in Illumina 16S sequencing protocol, see link in #3) is that identifying and eliminating cross contamination post-sequencing (caused either by physical contamination during library prep or image analysis error with higher cluster densities) would be easy as chance of both primers being contaminated is reduced.
This is is a way to identify contamination but is not an "obvious advantage " over a 1-step library generation. A 1-step workflow that integrates indexes and adapters at the 1st PCR so contamination is just as easy to spot.


Quote:
Originally Posted by nucacidhunter View Post
Cluster generation should not encourage chimera formation because bridge amplification takes place in solid state and unlike in solution PCR, fragments are not free to interact with each other. Fragments that are in close proximity or hybridize to each other and may fuse and form a cluster, will not produce pure signal either because of presence of mix template amplification or multiple primer binding sites and will fail filter so they cannot contribute to chimera reads.
Agreed, chimera's are not really an issue at cluster stage. But it is an amplification stage so one contaminatiing template molecule could initiate that cluster.


Quote:
Originally Posted by nucacidhunter View Post
It is not clear to me what sort of contamination would built over time, if one follows standard molecular biology lab hygiene such as using disposable gloves, aerosol free tips and cleaning bench and pipette surfaces.
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Aerosols build in labs over time. In a post PCR area you can use ART tips, UV and gloves but this minimises contamination it won't remove it. A good PCR once 'opened' will generate aerosols with 10^5-10^9 copies of DNA that can travel through HEPA filters and build in a lab.

But to bring this banter back to esherman's question - there are good ways to generate amplicon libraries using both 1-step and 2-step methods there are strengths and drawbacks to both approaches. How you tackle this is very much dependent on budget, sensitivity, contamination concerns and the application you intend for the data.
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