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Old 07-19-2016, 03:10 AM   #1
JBKri
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Default MinION protocols?

Is there a site where I can find detailed protocols for library prep for MinION? A few days ago I signed up for the community, but I still don't have access to the community website. I would like to know more about what consumables and equipment is necessary for various library types. The "Equipment and consumables" pdf is rather general and suspect it does not cover all types of libraries.
Jon
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Old 07-21-2016, 07:33 PM   #2
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If you're doing the 15-minute rapid (1D) sequencing, then you'll need a small Blunt/TA ligase kit from NEB (M0367), less than 1μl is used, so it'll last for ages for the rapid sequencing prep.

For the standard gDNA (and/or amplicon) protocol, you'll need a bit more:
  • MyOne C1 Streptavidin beads (or equivalent)
  • AMPure XP beads (or equivalent)
  • Nuclease free water
  • NEB Blunt / TA Ligase Master Mix (50μl used per library, so it's fairly expensive)
  • NEBNext Ultra II End-repair / dA-tailing Module
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Old 07-21-2016, 11:27 PM   #3
nanos
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maybe also add the NEBNext FFPE DNA Repair mix to repair the input DNA for the standard protocol. This was shown to increase sequence quality and length quite a bit.
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Old 07-22-2016, 07:51 AM   #4
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Is it correct that the g-TUBES are optional?
Is the Hula mixer really necessary; can't we just mix by hand?
Could you suggest an alternative to the MyOne C1 Streptavidin beads? The smallest bottle would cost us as much as the MinION starter pack!
Jon
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Old 07-22-2016, 12:45 PM   #5
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  1. Yes, g-Tubes are optional. You can fragment with a fine-bore pipette if you want something really cheap, or no fragmentation at all. The MinION will sequence reads over 50kbp, and just the act of pipetting will get reads down to that size unless you're really careful.
  2. A rotator works as well. I suppose you could mix gently by hand, but it's a 5-minute process -- do you really want to be slowly windmilling your hand for 5 minutes?
  3. Other beads are available, just make sure that you alter the initial added amount to correspond to the correct bead concentration.
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Old 07-26-2016, 09:33 AM   #6
JBKri
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Quote:
Originally Posted by gringer View Post
  1. Yes, g-Tubes are optional. You can fragment with a fine-bore pipette if you want something really cheap, or no fragmentation at all. The MinION will sequence reads over 50kbp, and just the act of pipetting will get reads down to that size unless you're really careful.
  2. A rotator works as well. I suppose you could mix gently by hand, but it's a 5-minute process -- do you really want to be slowly windmilling your hand for 5 minutes?
  3. Other beads are available, just make sure that you alter the initial added amount to correspond to the correct bead concentration.
Thanks, this is very useful information.

We do have a bottle of Dynal M270 streptavidin beads, but it expired in 2012. I think the bottle has only been opened once, and it has always been stored at 4 C. Do you think we could still use it? As far as I can tell, the major difference between C1 and M270 is that the C1 has twice the binding capacity, and they are smaller (1 um versus 2.7 um for the M270).
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Old 07-26-2016, 11:05 AM   #7
Markiyan
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Lightbulb My version of the 2D library protocol (2016/07/13).

Quote:
Originally Posted by gringer View Post
  1. Yes, g-Tubes are optional. You can fragment with a fine-bore pipette if you want something really cheap, or no fragmentation at all. The MinION will sequence reads over 50kbp, and just the act of pipetting will get reads down to that size unless you're really careful.
  2. A rotator works as well. I suppose you could mix gently by hand, but it's a 5-minute process -- do you really want to be slowly windmilling your hand for 5 minutes?
  3. Other beads are available, just make sure that you alter the initial added amount to correspond to the correct bead concentration.
Best is to fragment with syringe needle (if you have 4-10ug of DNA available per lib prep)
In my case (for my second nanopore run) I had done the following:

1. Diluted my freshly preped DNA with NFW (you can use TE) to total of 400uL,
2. Sheared it by passing through 21G 4' needle attached to 1ml syringe for 20 times
(or use 26G one for 3-5 times),
3. Than performed an Ampure XP wash at 0.45X ratio (+180 uL of beads). Incubate on the rotator for 10 min. Pellet at 60 degrees angle (help gravity and magnetic forse to work in the same direction for ~5min.
4. Wash beads 3 times with 1mL of 70% ethanol.
5. End repair/A-tailing times were set as per NEB recommendation (30 min each instead of 5). All library handling after this step was done with the large bore tips.
6. Intermediate ampure XP washes were done on 0.45X ratio (instead of 1X),
7. The first steps of the ligation had been done for 20 min on the RT and for 30 min in the fridge (ligating adapter with a huge helicase attached to it to a long (25-50kb) DNA fragment requires cool and quiet conditions for quite a while).
8. The tether ligation was done for 20 min at the RT.
9. Dynabeads streptavidine M280 beads were washed with the ONT kit's bead wash solution, leave to bind for 10 min.
10. Pellet at angle ~60 degrees (to have gravity and magnetic field working in the same direction).
11. Resuspend with extreme care - like cosmid library ligation tube flicking.
12. Pellet again at an angle, and repeat the wash. Elute as per protocol.
13. Make sure to degas the fuel mix and water used to prime the flowcell (otherwise there would be a lot of tiny bubbles over the ASIC array after a few hours @34 degrees).
I had used an Eppendorf Concentrator Plus set to V-AQ mode for 90-180 sec @ 20 degrees.
14. Entire library had been loaded.
15. The run had started at -185 mV bias voltage, and by the 48h the voltage had gone to -250mV.

The modal RAW read length had been 8.5kb, and there was a long tail with a good fraction of 20kb, 30kb, even quite a few 60kb raw reads. Total RAW events had been ~450M.

PS: it looks like the manufacturers adapter ligation conditions had been optimised for the previous version of the kit, when there was no motor protein attached to the adapter during a ligation step. The attachment of the motor changes the adapter's molecular dynamics quite a bit (by increasing it's mass by ~2 orders of magnitude), which increases the time required for efficient ligation to the long DNA fragment - have a read of the BAC/YAC library construction protocols, ligation conditions section, to give you some more background info.

So the results had been a raw yiels of ~446M events, with the following raw reads distribution (see attachment):
M160713_48h_raw_reads_histogram.png

For metrichor basecalling results see attachments: M160713_metrichor_48h_1D_res_c2.png and M160713_metrichor_48h_2D_res_c2.png
Attached Images
File Type: png M160713_48h_raw_reads_histogram.png (37.0 KB, 79 views)
File Type: png M160713_metrichor_48h_1D_res_c2.png (77.0 KB, 70 views)
File Type: png M160713_metrichor_48h_2D_res_c2.png (63.4 KB, 41 views)

Last edited by Markiyan; 07-27-2016 at 07:39 AM. Reason: clarifications/+results.
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Old 07-26-2016, 11:19 AM   #8
Markiyan
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Exclamation Take care with old streptavidine beads -there are quite a few things that can grow in

Quote:
Originally Posted by JBKri View Post
Thanks, this is very useful information.

We do have a bottle of Dynal M270 streptavidin beads, but it expired in 2012. I think the bottle has only been opened once, and it has always been stored at 4 C. Do you think we could still use it? As far as I can tell, the major difference between C1 and M270 is that the C1 has twice the binding capacity, and they are smaller (1 um versus 2.7 um for the M270).
I would be extremely wary of using it - check under the good microscope that there is nothing growing there (liquid is clear after standing in a fridge),
PS: For larger fragments I think that the Dynabeads M270 would be a bit better (they have to swim with quite a long tails).

PPS: do not do pipette shearing - can generate quite a few short fragments/strabd breaks (my first go at nanopore). better use syringe needle fragmentation with freshly prepped DNA.
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Old 07-27-2016, 12:58 AM   #9
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@Markiyan, if I may ask, how do you degas these buffers?
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Old 07-27-2016, 02:51 AM   #10
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Lightbulb Just by using some slight vacuum.

Quote:
Originally Posted by nanos View Post
@Markiyan, if I may ask, how do you degas these buffers?
Just by using some slight vacuum without heat, so the dissolved gasses escape first. This was done using eppendorf concentrator for 90 sec (you do not want to actually to concentrate your buffers much, just get rid of the dissolved gasses).
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Old 07-29-2016, 05:48 AM   #11
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Quote:
Originally Posted by Markiyan View Post
I would be extremely wary of using it - check under the good microscope that there is nothing growing there (liquid is clear after standing in a fridge), ...
.
Well, at 200x magnification, the liquid loooks clean and clear, and the beads look good...
Jon
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Old 08-10-2016, 12:39 PM   #12
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Quote:
Originally Posted by Markiyan View Post
Best is to fragment with syringe needle (if you have 4-10ug of DNA available per lib prep)
What would you change if you were doing a library prep with only 300-500ng after shearing?

Do you have any ideas on modifications for the low input protocol (20ng)?
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Old 08-10-2016, 01:17 PM   #13
Markiyan
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Lightbulb

If one would to shear smaller ammounts of DNA - than get yourself the Hydroshear with custom assembly or megaruptor (so you can fragment in
smaller volume).

Make sure to us FPE repair mix if you DNA is not fresh or could have been damaged during prep.

If one is has 300-500 ng of DNA after shearing - I would keep 0.45X - 0.6X ampure beads ratio for 2D protocol (provided your sheared DNA is >2kb, and you want to loose small fragments). One may think about halving the ammount of adapter used in the ligation compared to 1ug input and extending ligation times.

For even lower ammounts of DNA:

One option can be using low input expansion kit (whole genome amplification) if your input is in the region of 20ng. Modify the ampure beads ratios and wash ethanol volumes as per main protocol.

Personally I deal with cultured samples, so it is not a problem to grow more if needed, but if you have a precious one, than practice on a not very important one, before moving to the precious ones.

PS: If you get any nice results using my modifications - please post the statistics (raw reads length distribution and 1D/2D basecalling quality reports), so the other users can have some idea about the impact.

If you had made any futher improvements to the protocol - please report them too, so others can benefit.
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Old 08-18-2016, 08:23 AM   #14
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Thanks for your suggestions @Markiyan.

I have modified the amount of ampure in the low input protocol in the past. The results have always been the same. This is probably due to low amounts of DNA in the entire solution and adjusting the beads amount just showed that I was wasting beads by doing a 1:1. I would probably have to go to a lower ratio of ampure to size select low input….but do I really want to…

Secondly, I have been running metagenomic extractions, so DNA has been sheared and beaten by beads. When I performed FFPE repair on low input in the past it showed no difference in quality or read length.

I did try extending ligation in hope that longer fragments in my low input were not getting enough attention. Unfortunately, I had an overwhelming amount of intermolecular ligation; my chimeric reads were very high. More dilute conditions probably would mitigate it and the right pmol amount of adapters for each run but then again beads beaten low input DNA doesn’t really have a size disruption.

ONT also released their views on low input protocol being sub optimal and I should turn to PCR DNA to get higher quality reads.

What extraction method do you use for cultured samples? I know people have used isolate II genomic DNA kit but your quality/quantity look great.

I will update when I run more sample test.
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Old 09-11-2016, 08:07 PM   #15
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From the equip for Nanopore Sequencing Kit gDNA protocol, one of the listed consumable is 10 mM Tris-HCl pH 8.5. Anyone knows which step would use this buffer? Could 10 mM Tris-HCl pH 8.0 be used instead? Thanks.
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Old 09-12-2016, 02:02 PM   #16
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The Tris-HCl is used for the elution from Ampure beads. As far as I know the higher pH facilitates the resolubilization of high molecular weight DNA.
You can definitely use pH8, but you might loose more DNA from the long fraction because it does not come off the beads.
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Old 11-03-2016, 12:19 PM   #17
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This 48 hr run generated ~ 300 Mb data, is this a usual case?

We tested a 2 hr run, and only generated 29 Mb data. But the protocol said 48 hr should generate 20 Gb. Are there any tricks to improve output yield?

The Y axis of MinKNOW histogram is labeled as "total # reads". Does this "reads" mean base or DNA fragment? We tested a 2-hr run, the MinKNOW histogram peak is more than 10,000,000. But we only got ~ 4000 reads passed.




Quote:
Originally Posted by Markiyan View Post
Best is to fragment with syringe needle (if you have 4-10ug of DNA available per lib prep)
In my case (for my second nanopore run) I had done the following:

1. Diluted my freshly preped DNA with NFW (you can use TE) to total of 400uL,
2. Sheared it by passing through 21G 4' needle attached to 1ml syringe for 20 times
(or use 26G one for 3-5 times),
3. Than performed an Ampure XP wash at 0.45X ratio (+180 uL of beads). Incubate on the rotator for 10 min. Pellet at 60 degrees angle (help gravity and magnetic forse to work in the same direction for ~5min.
4. Wash beads 3 times with 1mL of 70% ethanol.
5. End repair/A-tailing times were set as per NEB recommendation (30 min each instead of 5). All library handling after this step was done with the large bore tips.
6. Intermediate ampure XP washes were done on 0.45X ratio (instead of 1X),
7. The first steps of the ligation had been done for 20 min on the RT and for 30 min in the fridge (ligating adapter with a huge helicase attached to it to a long (25-50kb) DNA fragment requires cool and quiet conditions for quite a while).
8. The tether ligation was done for 20 min at the RT.
9. Dynabeads streptavidine M280 beads were washed with the ONT kit's bead wash solution, leave to bind for 10 min.
10. Pellet at angle ~60 degrees (to have gravity and magnetic field working in the same direction).
11. Resuspend with extreme care - like cosmid library ligation tube flicking.
12. Pellet again at an angle, and repeat the wash. Elute as per protocol.
13. Make sure to degas the fuel mix and water used to prime the flowcell (otherwise there would be a lot of tiny bubbles over the ASIC array after a few hours @34 degrees).
I had used an Eppendorf Concentrator Plus set to V-AQ mode for 90-180 sec @ 20 degrees.
14. Entire library had been loaded.
15. The run had started at -185 mV bias voltage, and by the 48h the voltage had gone to -250mV.

The modal RAW read length had been 8.5kb, and there was a long tail with a good fraction of 20kb, 30kb, even quite a few 60kb raw reads. Total RAW events had been ~450M.

PS: it looks like the manufacturers adapter ligation conditions had been optimised for the previous version of the kit, when there was no motor protein attached to the adapter during a ligation step. The attachment of the motor changes the adapter's molecular dynamics quite a bit (by increasing it's mass by ~2 orders of magnitude), which increases the time required for efficient ligation to the long DNA fragment - have a read of the BAC/YAC library construction protocols, ligation conditions section, to give you some more background info.

So the results had been a raw yiels of ~446M events, with the following raw reads distribution (see attachment):
M160713_48h_raw_reads_histogram.png

For metrichor basecalling results see attachments: M160713_metrichor_48h_1D_res_c2.png and M160713_metrichor_48h_2D_res_c2.png
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Old 11-03-2016, 12:33 PM   #18
gringer
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Quote:
Originally Posted by mkdir View Post
This 48 hr run generated ~ 300 Mb data, is this a usual case?
That depends on what flow cell you're running. If it's an R9 flow cell, then 300Mb of basecalled data is reasonable, although with good sample prep and good, fresh flow cells, it can get to a couple of Gb.

Quote:
Originally Posted by mkdir View Post
We tested a 2 hr run, and only generated 29 Mb data. But the protocol said 48 hr should generate 20 Gb. Are there any tricks to improve output yield?
What protocol? Clive Brown has recently achieved an in-lab yield of 10Gb basecalled with 1D ligation kit + R9.4 flow cell (i.e. the most recent and fastest available kits and flow cells). It would be weird for any protocol to suggest a yield over twice that.

Without knowing anything more about your sample prep, it's difficult to say. If you're going for maximum yield, fragment FFPE-repaired (or PCR amplified) dsDNA into pieces of about 3-8kbp, and make sure you're loading library onto the sequencer that has a hairpin-containing dsDNA concentration (as quantified by a fluorescence-based method) of at least 4 ng/μl. Also, choose a flow cell that has >450 active channels at initial QC, and doesn't have SPDS (sudden pore die-off syndrome).

Quote:
Originally Posted by mkdir View Post
The Y axis of MinKNOW histogram is labeled as "total # reads". Does this "reads" mean base or DNA fragment? We tested a 2-hr run, the MinKNOW histogram peak is more than 10,000,000. But we only got ~ 4000 reads passed.
It's actually the predicted number of called bases. If there are 1000 reads with a read length of 8kb, then the histogram value would be 8,000,000.
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Old 11-03-2016, 12:49 PM   #19
mkdir
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Hi gringer, thank you very much for the information. The nanopore website suggested 21Gb (Product specifications comparison, https://nanoporetech.com/products#comparison). But you have answered all my questions. Thanks a lot.



Quote:
Originally Posted by gringer View Post
That depends on what flow cell you're running. If it's an R9 flow cell, then 300Mb of basecalled data is reasonable, although with good sample prep and good, fresh flow cells, it can get to a couple of Gb.



What protocol? Clive Brown has recently achieved an in-lab yield of 10Gb basecalled with 1D ligation kit + R9.4 flow cell (i.e. the most recent and fastest available kits and flow cells). It would be weird for any protocol to suggest a yield over twice that.

Without knowing anything more about your sample prep, it's difficult to say. If you're going for maximum yield, fragment FFPE-repaired (or PCR amplified) dsDNA into pieces of about 3-8kbp, and make sure you're loading library onto the sequencer that has a hairpin-containing dsDNA concentration (as quantified by a fluorescence-based method) of at least 4 ng/μl. Also, choose a flow cell that has >450 active channels at initial QC, and doesn't have SPDS (sudden pore die-off syndrome).



It's actually the predicted number of called bases. If there are 1000 reads with a read length of 8kb, then the histogram value would be 8,000,000.
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Old 11-03-2016, 01:16 PM   #20
gringer
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Wow! I wasn't aware there were marketing people at ONT who were more optimistic than Clive Brown. Those numbers are technically true, but Clive has said that the current flow cell performance is only about 10% efficient (a bit more for in-lab flow cell testing).
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