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Old 08-06-2010, 05:40 AM   #1
KorNor
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Default GS Junior - bead issues.

Anyone else using a GS Junior?
I've set up ours and done a few runs - only moderately successful (user incompetence, ha!)

I am getting many more mixed beads than anticipated, even after quanting and using the calculator on the my454 site.

My bead counter always shows a level above 2mil at end stage.

Emulsions are not broken, etc etc

Anyone else had any problems like this with this platform? Any hints?

Last edited by KorNor; 08-06-2010 at 06:55 AM.
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Old 08-06-2010, 11:29 AM   #2
Baseless
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Hello KorNor,

I can only speak from experience from the FLX, but there mixed beads result from too much DNA fragments per beads during the emPCR (and this is probably the situation in the entire workflow i fear the most).
I just took a look at the emPCR protocol for the Junior, it clearly states that if you have a bead count above 2min (this equals a recovery of more than 20%) you should not continue.
Since they claim your optimum window is between 500kBeads and 2mBeads, the window looks very familiar from the FLX (5-20%).

Empty or mixed beads are a very sensitive matter and even if someone needs data fast, never make compromises here, this can fail entire runs.

I suggest that you a) requantify your library and retry and/or b) cross check what I just said with customer support.

If you have access to a bead counter, you could determine the exact recovery rate and win an insight how much too much it actually was. But be aware that below 2% and above ~35 % these counts are also no longer trustworthy.
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Old 08-06-2010, 01:59 PM   #3
KorNor
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Hi Baseless,

Yes, I understand that; the first time was during training run, and despite knowing we had over 2 mil beads, we pushed on, just to get the practice of setting up the run.
The next time, I reduced the template DNA by 1/3, still over 2 mil beads (I didn't put a seq run on with that). Repeated the emPCR again, using fresh library, again over 2 mil beads (again, I didn't put the seq run on).

I'll talk to tech support again next week.

Last edited by KorNor; 08-06-2010 at 02:02 PM.
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Old 08-26-2010, 11:57 AM   #4
Palecomic
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Hi KorNor,

Just doing my training on newly installed GS Junior at the moment, we had exactly the same problem. More than double the upper limit of the range of beads in the bead counter. After interrogating the trainer, it seems that they've removed the titration step that's in the FLX protocol and specified 2 molecules per bead as ideal (in this case for 16S amplicon sequencing), but it looks like it's probably not good in this case.

Will see whether it improves next time, probably going to go with 0.5 and see whether the enrichment is more acceptable.
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Old 08-26-2010, 03:43 PM   #5
KorNor
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Hi Palecomic,
Thanks for your reply; after speaking with tech support, I too have dropped the 2 molecules per bead in favour of a 0.6 - 0.8:1 ratio and that has been working well (if a little inconsistent). I think once you've done a few dozen runs, you'll probably have a better feel for it, but 2:1 is certainly much too high.
Otherwise, she's a good little breadmaker!
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Old 08-27-2010, 12:58 AM   #6
Palecomic
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That's really good to know, thanks. I might suggest to them that in the next update to the manual they change that example.

Whereabouts are you? I'm not sure that there are that many Juniors about at the moment, I've just started a postdoc at Imperial.

Cheers

Mike
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Old 09-18-2010, 05:58 PM   #7
plnagy
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Anybody using Agilent Sureselect routinely for capture for sequencing by the 454JR ?
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Old 10-18-2010, 10:08 PM   #8
James Haile
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Default Splitting GS Junior Emulsion Oil

Has anyone tried splitting the GS Junior emulsion oil? i.e. adding the A beads (and Live A mix) to half the volume of oil, and the B beads (and Live B mix) to the other half? This way, if you don't need your sequences read in both directions, you could use two different bead to template ratios. Cheers!
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Old 10-22-2010, 07:41 AM   #9
Palecomic
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plnagy: we haven't tried that system.

James: we haven't tried that either - I'm still getting to grips with the minutiae of the method, what would be the advantage of having both A and B beads in the same mix?

Do you mean you could set up A beads with 0.6 molecules per bead, B beads with 2 molecules and then capture them separately, sequencing whichever one had the level of enrichment that you wanted? It would avoid the issue of losing an emPCR kit every now and then to the wrong ratio.

I've only used the Lib-L kit so far, is separating the beads efficient and straight-forward?
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Old 10-28-2010, 11:15 AM   #10
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Hi KorNor et al.,

We've actually had a problem going the other way: despite our bead recovery being JUST at the 500,000 mark (i.e. barely recovering enough beads), we lost more than 98% of our reads to the mixed filter on two shotgun runs on different organisms. Emulsions aren't broken, and the sequence that does manage to pass the filter is good quality and maps well to our reference (suggesting that there isn't anything inherently wrong in our sample preps? Your thoughts? One of the library preps was set up under Roche supervision, the other exactly following the protocol...so unless their Junior library prep protocol has a whole in it?).

Control beads produce good sequence, and running Roche's canned E. coli library works fine (so there are no problems with our emuls-PCR technique or sequencing set-up...). We've tried reducing our input library by 1/3, but that has also reduced the amount of beads we recover to below the 500,000 magic limit. It is like we are getting overenrichment of beads, without overenriching! Tech support is thinking there might be a sample-specific problem in the emuls-PCR: anyone got any ideas? We are fresh out of them, and not looking forward to being told just to "run it again".

Any help would be super-greatly appreciated...
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Old 01-03-2011, 06:29 PM   #11
Jacky76
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Default Hello

In my experience, you should get less enriched beads if you decreased the concentration (cpb). But if you are getting 2 mil. beads continuously, there must be some problem. Amplicons tends to be amplified during the emPCR more than shotgun. And if you are doing amplicon, you should check with PicoGreen. The assay is more accurate than Agilent calculation. And if you are doing shotgun, you don't have to leave the library and the standard longer than 6 hours after you finish the library prep. It is also recommended to measure mol/ul using fluorometer as soon as you can for using the calculator on my454.
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Old 03-01-2011, 06:59 AM   #12
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hi, Ive had many cases with overenrichment with FLX and most were due to broken emulsions, detectable by settled beads in the wells after ePCR. However i did have one or two samples with continuous overenrichment and the tech support told me that it may be due to the nature of the sample. I would re-quantify my library and measure using picogreen or other similar measures like HS Qubit and calculate your molecules/ul. Maybe run a titration with 0.1cpb and 0.5cpb and make sure you wash as long as it takes until no white beads are coming off. Also, i have had a problem with some MV oils as it was resulting in broken emulsions ( detectable by settled beads in the wells after ePCR, whether or not there was a phase separation in the wells) which Roche replaced...
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Old 04-17-2011, 09:08 AM   #13
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My experience with GS Jr, perhaps, is the most illogical case of all. I am working with ancient DNA extracted from bone, not as old as Neanderthal, but quite old. Amounts of DNA extracted are at the detection limit of the Picogreen assay, so library quantification was likely unreliable. Based on this estimate using an about 50% of calculated recommended library volume, I got right amount of enriched beads, between 5e5 and 2e6. However, 98% did not pass the mixed bead filter. Next, I used 2 times less for emPCR. This time I got about 7% of valid reads, a tiny bit better. Yet, when I took 20 times less I got down only to 83% of lost reads. Make a little sense, huh?
Blaming library quantitation, I used SYBR Green qPCR with emPCR primers to requantitate the library.The estimate by SYBR Green was 100x higher than qPCR data! So it lloked like I was actually working with even less library than I thought, yet I was getting most of the reads lost to mixed bead filer. Makes even less sense , huh?
Anyway, I used this new concentration to load emPCR based on recommendd calculations. I got a way over 2e6 enriched beads. I reduced amount of library 5 fold, got a bit less packed enriched beads, but still a way over 2e6. Then I took 25 times less library, but I got even more packed enriched beads than the first time! This is as weird as it gets. I did not run any of these samples, but I am tempted to since the entire story makes no sense.
Tech support suggested that library denaturation may not be sufficient before loading. I do denturation step as recommended in the manual, except instead of letting thermocycler to chill to 4C, I stick the tube in ice to minimize annealing.
Another problem may be broken emulsions. In new emPCR kits, there is a yellow insert saying that oil formulation was changed and a top clear layer after PCR is normal, and does not indicate broken emulsion. However, that is exactly what shown in broken emulsion example in the manual, so it is very confusing now, since I always see this clear layer.
Does anyone experienced similar lack of logic?
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Old 04-18-2011, 02:42 AM   #14
Palecomic
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Hi Yaximik,

With your emulsions, do you see a small clearing at the very bottom of the tubes?

We're doing amplicons and have carefully optimised our cpb ratio so that we get 5 % enrichment every time. We recently had an extra training session with some new staff, and the training kit included the newer oil along with yellow slip. The emulsions had a larger clear layer on the top as we were warned of, but also a very thin clear layer at the bottom. There was no sign of the classic banding and no pellet of beads that had dropped out of the emulsion. The emPCR gave us 20 % enrichment, much more enriched than expected, and the run failed due to over 50 % of the reads being mixed.

Another feature of the run was that of the reads that did make it past the filter, there were significant errors in the primer sequence (50 % again of the remaining reads). Roche confirmed that this was also the case with the control beads in the run, so perhaps it's worth checking this with the few reads that you do get?

I'm rerunning the same samples today with another kit, it does have the more turbid oil, but doesn't seem to include the yellow slip. I'll have to check, but I think it's a different lot number. Will let you know how it goes, but I've informed Roche and they are following up, so hopefully if this is another case of dodgy oil they'll track this through.

Over all, I've found that we've had a lot of run variability, and that if we sequence anything over 5 % enrichment, it's never a good run (20 % is definitely bad). Our pump has been replaced (as has the pump of one of the other guys with a Jr in the UK) and these are the first runs after that, making this trickier to troubleshoot, as we can't control all the other variables.

Good luck!
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Old 04-19-2011, 02:30 AM   #15
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OK - to update, we have the same problem again. A very small volume of clearing at the bottom of the emPCR, no classic banding of the emulsion and this time it is possible to see a small pellet of beads.

I will be setting up the same library yet again with an older kit to double check that it's not other aspects of our work flow.

In the meantime if anyone comes across a similar problem to that detailed by myself or Yaximik it would be really useful if you could mention it here as well as to Roche. The lot number of oil I have a suspicion of is 93807320.

Thanks!
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Old 04-19-2011, 03:56 AM   #16
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Zippy, did you use rapid libraries for your shotgun? If so, were you sure you did the denaturation step in the emPCR set-up? A low enrichment, plus such a high proportion of mixed reads would point to non-denatured rapid library in the emPCR. You would have dsDNA going into the emPCR with each strand being in opposite directions, hence all reads would come out as being mixed.

There is a theoretical limit for how many beads you can get back from emPCR, so it could be that you're well beyond that limit. Reducing the input may not result in the same proportional drop in enriched beads. For our 16s studies, we've found that using the Bioanlyser sizing, plus a Qubit/Picogreen quantification gives us hightly reproducible results on our FLX. We pretty much get 10% enrichment +/- 2% every time with 0.5cpb.
If in any doubt, use qPCR (such as the Kapa kit) but be aware that we've found different library types require different emPCR input.
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Old 04-20-2011, 10:26 AM   #17
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Many emPCR reactions can be started with dsDNA, the oligo bound to the bead is usually the B' or A'. Starting emPCR with ssDNA just avoids too much solution phase PCR. I'm not sure how you could get reads in all directions as described.
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Old 04-21-2011, 06:00 AM   #18
yaximik
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Well, I could not stand the lack of logic in my experience described in the previous post and made a seq run using beads that by the protocol must have been considered failed because I got about 4-6 million enriched beads. Guess what? I got the best result so far with this ancient DNA, with 23% passed filters, while with all these diluted samples before the best was 17%. The distribution of reads was quite intersting though, with two maximums, one around 450-550 bases and another around 100 bases, with minimum around 250 bases. The total number of reads was about 50% of what I got with the worst case (1% usable reads).
In fact, after reading the original paper and the emPCR protocol I realized that it is quite possible to get over 2 million enriched beads, contrary to what is stated in the protocol. The original paper says they got about 30% enrichment. The kit does not say how many capture beads are supplied, but assuming that the formula provided in the manual gives a real example, one gets 10 million beads with the kit. So, simple math gives you 3 million beads of normal enrichment output.
I also feel that amount of beads one gets is quite dependent on how harsh the washing steps are performed. The enrichment beads are likely 0.8 micron streptavidin magnetic beads (I scoped them), much smaller than ~30 micron capture beads, meaning that strength of binding to a magnet will depend on how many brown enrichment beads are attached to the white capture beads. Instead discarding unbound beads right away as recommended, put these in another tube and stick into a magnetic separator - you will be surprised how many almost white beads will keep sticking to the magnet, meaning these also bear amplified library fragments! The harsher wash the more of these weak-binding beads will go down the drain.
I feel the lack of logic in my experience has something to do with small fragments present in library that distorts library quantification. Perhaps, sizing did not work the way it is supposed to work. I found one reference, which uses double-step sizing, so I want to try it to see if I get more logical results.
I do not think that I have broken emulsions. I get very consistent picture every time - white sediment and clear oil layer on top.
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Old 04-22-2011, 07:35 PM   #19
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OK, again another curious result. I used a totally different sample library this time, made from DNA extracted from an about 100-old tooth, as compared to previous excercises with ~1000-old bones/teeth. Library concentration was estimated to be at ~1.5e7 mol/ul, so I used 1 ul. From the enrichment step I got about 3 million beads (by GS Jr bead counter), which means my emPCR supposed to be failed again. Nope. I did a seq run with recommended 0.5 million of these beads - 37% reads passed all filters. Out of 173.5K reads (E.coli traning generated 240.5K reads), the largest loss was not after the mixed filter but after the dot filter. Overall distribution was 45.7K lost at the dot filter, 21.8K lost at the mixed filer, and 25.5K lost at the short quality filter. Read lenghts peaked at 500 nt, with a minimum around 300 nt and a substantial number of reads of 50-250 nt. The longest read was 1038 nt, not bad at all. This likely means that at about 30% enrichment the picotiter plate was still not loaded to its full capacity (in terms of beads carrying amplified fragments), many wells had beads with either too short fragments or produced no signal. Which means that the manual has a very narrow applicability and considering the variability of samples everyone is basically on his/her own to get the best from his/her libraries. Do not throw away your purportedly failed beads! Question authorities...

I keep y'all posted on furter developments.
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Old 05-03-2011, 03:09 AM   #20
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epistatic, it was my understanding that the Y adapters contain both the A and the B sequences, annealed at one end. When ligated to an adenylated dsDNA library, this would cause both strands to have the A and B adapters attached, as such:

A-LIBRARY-B
B-LIBRARY-A

If you didn't denature, then both strands would go into the micro reactor, anneal to the same capture bead and then amplify. Sequencing using the A primer, would then give one read: LIBRARY and another read YRARBIL on the same bead, which would be mixed signal.
It's different for amplicon libraries as only one strand of the amplicon would contain the A and B sequences in the correct orientation. ssDNA libraries only have one strand anyway, so there's no issue with mixed reads.

yaximik, we did a run recently where we had large numbers of small fragments in our library. We used the Bioanalyser/Qubit to estimate concentration, using the largest peak as our average size.
Our enrichments were much higher than normal, (approaching 30%). We believe that this was because our quantification was wrong (smaller library lengths mean a higher molecule/ÁL per ng of DNA). We sequenced anyway and got much higher dot fails than normal.
Roche said, and I quote "These shorter fragments will lead to brighter signals, ending up in elevated dot filtration".
I think, you may need to be more stringent in your method of size selection. Ampure XP beads may not be removing all the small library fragments. Some of your read lengths do look a bit on the small size. Perhaps you should look at a gel-cut option for a run and see if you can improve your selection.
Can I ask how you are estimating your library concentration and counting your beads? Do you get a stable raw read output from all your runs?
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