Nextera XT on HiSeq

[Christine]

Hello all,

I'm wondering if anyone has sequenced Nextera XT libraries on the HiSeq and what modifications were made to get the cluster density in range?

I've been sequencing the XT libraries on the MiSeq so far and am very pleased with the performance (and the normalization!). We're scaling up to a HiSeq run next, with up to 96 libraries per lane. Needless to say, I was looking forward to skipping all those individual library quantifications, but Illumina has advised me not to use the normalization beads when prepping for the HiSeq run. Their reasoning was that the concentration of the library pools will be too low to get a decent cluster density.

The yield from the beads is not sufficient for optimal cluster density on the HiSeq. This is why we recommend to skip the normalization step if you plan to sequence Nextera XT libraries on the HiSeq. At this step you can quantify your libraries using qPCR. It is a bit more work, but it is the only way to successfully sequence Nextera XT libraries on the HiSeq.

Surely there's a simple workaround. Can I concentrate the pools on some columns, followed by 8 qPCRs instead of ~800? The libraries will be single stranded, so the binding efficiency might not be great and the qPCR will need a correction factor, but I think it's worth a try. What do you think, seqanswers?

[pmiguel]

Yeah, that is disappointing. Zymo has ssDNA/RNA clean-up/concentration columns. My guess is that they would work.

--
Phillip

[genomeseeker]

We're starting a HiSeq run next week on 2 pools of Nextera XT libraries. We quantified the final pools by qPCR (KAPA kit) and plan to treat them like all other libraries and load them 11pM.

Here's the instructions we received from tech support:
As long as you are not interested in following the Amplicon workflow, you can, in theory, sequence your Nextera XT libraries on a HiSeq. Currently Nextera XT is only supported on the MiSeq. If you want to sequence them on the Hiseq, here are some recommendations and guidelines. Please note that this is NOT supported or guaranteed.

1. Stop the Nextera XT protocol after PCR clean-up and don't proceed to Library Normalization. At this point, your library is already made.
2. Quantify each library using a dsDNA-specific fluorescent dye method, such as Qubit or picogreen.
3. Normalize and pool the libraries as needed.
4. Cluster on a cBot and use sequencing primers from the TruSeq Dual Index Sequencing Primer Box, Single Read or Paired End, depending on the type of the run.

[koadman]

Hi all, I've been looking at this as well. Reading through the Illumina protocol pdf, I noticed the following:

- Add 576 μl of HT1 to the DAL tube.
- Transfer 24 μl of PAL to the DAL tube containing HT1. Using the same tip, pipette up and down 3–5 times to rinse the tip and ensure complete transfer.


The recommended volumes for diluting PAL with HT1 represents a 25-fold
dilution. This dilution ratio was established by using the recommended
equipment (e.g. plate shaker calibrated for shaking speed) and following the
normalization procedure strictly under typical laboratory conditions (e.g. 20°–
25°C). If cluster density is found to be too high or too low, you may change
this dilution ratio to better suit the equipment, temperature, and user handling
in your laboratory after validation.

Which suggests that one might be able to play with the relative amounts of hybridization buffer and bead-normalized sample material. Would anybody care to speculate on whether this would or would not work for getting a library at the right loading concentration for the HiSeq? My understanding is that HiSeq loading concentration should be about 1.5x the miseq. The protocol calls for 24uL library in 600uL. What about 36uL library in 600uL?

[pmiguel]

Hi all, I've been looking at this as well. Reading through the Illumina protocol pdf, I noticed the following:



Which suggests that one might be able to play with the relative amounts of hybridization buffer and bead-normalized sample material. Would anybody care to speculate on whether this would or would not work for getting a library at the right loading concentration for the HiSeq? My understanding is that HiSeq loading concentration should be about 1.5x the miseq. The protocol calls for 24uL library in 600uL. What about 36uL library in 600uL?

cBot wants 120 ul of library/lane. Note that HiSeq lanes have a much larger surface area than a MiSeq lane.

A few labs reported results from clustering at a given concentration on their HiSeq in another thread. We had results consistent with those obtained by most of the labs posting in that thread. That is 16 pM resulted in about 800-850 Kclusters/mm^2. 16 pM is about 10 million amplicons/ul. So 1.2 billion amplicons were consumed per lane to yield about 240 million raw clusters. 20 % efficiency -- not bad. (But can either strand of an amplicon anneal to the flow cell oligos? That would be a 2x factor to consider.)

To get about the same cluster density you would need 600 ul of ~11 pM. Is that right, 4 billion amplicons? What does that get you using v2 hardware? 12 million clusters? 0.3% efficiency?

Please check my logic/math. Seems pretty outlandish that you would use more amplicons to yield orders of magnitude lower numbers of reads on the MiSeq.

If true, that suggests, in principle, the normalization beads should yield enough amplicons to cluster a HiSeq lane with.

A couple of cautions, though:

(1) Not sure about the flow cell oligos on a HiSeq flow cell being able to capture either strand of an amplicon. Since the bead-based normalization of NexteraXT presumably removes one of the strands, it would be critical that there be oligos to capture the strand that is retained.

(2) NexteraXT calls for a heat denaturation prior to clustering. Whereas all HiSeq protocols call for base denaturation. Ignoring dilution issues, will heat denatured templates even work on the cBot? NexteraXT calls for the run to be started as soon as possible after heat denaturation of templates. Could be the cBot has a pace to leisurely for this to work?

--
Phillip

[FWOS]

Cbot denaturation

I am pretty sure that the cbot relies more on heat denaturation than on the NaOH base denaturation. So the base is just a failsafe. The first step in cbot cluster gen is an incubation around 98C. So the material that actually clusters is always single stranded. The single strands should cluster without much bias as long as they have both p5 and 7 adapters.

[pmiguel]

I am pretty sure that the cbot relies more on heat denaturation than on the NaOH base denaturation. So the base is just a failsafe. The first step in cbot cluster gen is an incubation around 98C. So the material that actually clusters is always single stranded. The single strands should cluster without much bias as long as they have both p5 and 7 adapters.

What oligos are on the flow cell? A double stranded amplicon would be structured like:

5'->>>>P5>>>>>>>>>>>>>---------insert----->>>>P7-complement>>>>-3'
3'-<<<P5complement<<<---------insert-----<<<<P7<<<<<<<<<<<<<<<<-5'

The flow cells have to have both P5 and P7 oligos on them to allow cluster PCR to occur. Does that mean that each strand is a separate amplicon, or is one non-functional for some reason?

--
Phillip

[josdegraaf]

It is remarkable that the diversity between what people cluster is so big. We use 10 pM and some others 16. Something that I find hard to believe is machine dependent.

Anyway, each strand is a separate amplicon. One initially anneals through the p5, the other through p7, or do I misunderstand you Phillip?

FWOS, just forget the NaOH to check Illumina did not for nothing changed the sealing on the NaOH row of the cBot plate...

[pmiguel]

It is remarkable that the diversity between what people cluster is so big. We use 10 pM and some others 16. Something that I find hard to believe is machine dependent.

You mean in the other thread? Yes. Actually looks like all the results are fairly linear for pM to Kclusters/mm^2, except for one outlier.

Anyway, each strand is a separate amplicon. One initially anneals through the p5, the other through p7, or do I misunderstand you Phillip?

Thanks, that is what I meant.

--
Phillip

[Zuuhl]

FWOS, just forget the NaOH to check Illumina did not for nothing changed the sealing on the NaOH row of the cBot plate...

Ahh, but the NaOH on the cBot plate it not used to make the initial library single-stranded.
It is used during the Denaturation-Hybridization step at to get the clusters single-stranded before the seq. primer annealing.

Kim

[docbio]

Sorry I didn't see this post earlier. We've been running NexteraXT libraries on the HiSeq with good results. Following the recommendations of Illumina, we stop after library cleanup (no bead based balancing), then we quantitate the libraries with a high-sensitivity Qubit assay.

Our HiSeq core requires minimum 10nM pools in 25uL volume. Using the approximation of 2nM = 1ng/ul for 1kb fragments, we figure that's 125,000 pg of DNA in 25uL, and divide that between however many samples we're pooling (48-96) to figure out the number of picograms of each library to pool. The pool volume winds up being quite high, but we run it through a 30k mwco amicon concentrator column to get our final 25uL pool. I was nervous about the whole ordeal but the balancing has been acceptable and the clustering has been very good.

Another caveat about the bead based balancing and the MiSeq, once you come out of the BBB you've got single stranded libraries. They claim only a 10-20% loss in a freeze-thaw cycle but in practice we've seen clustering drop from 50-70% with a single freezethaw between MiSeq runs.

So they work, if you pay attention to the caveats and avoid the bead based balancing step.

[matth431]

Sorry I didn't see this post earlier. We've been running NexteraXT libraries on the HiSeq with good results. Following the recommendations of Illumina, we stop after library cleanup (no bead based balancing), then we quantitate the libraries with a high-sensitivity Qubit assay.

Our HiSeq core requires minimum 10nM pools in 25uL volume. Using the approximation of 2nM = 1ng/ul for 1kb fragments, we figure that's 125,000 pg of DNA in 25uL, and divide that between however many samples we're pooling (48-96) to figure out the number of picograms of each library to pool. The pool volume winds up being quite high, but we run it through a 30k mwco amicon concentrator column to get our final 25uL pool. I was nervous about the whole ordeal but the balancing has been acceptable and the clustering has been very good.

Another caveat about the bead based balancing and the MiSeq, once you come out of the BBB you've got single stranded libraries. They claim only a 10-20% loss in a freeze-thaw cycle but in practice we've seen clustering drop from 50-70% with a single freezethaw between MiSeq runs.

So they work, if you pay attention to the caveats and avoid the bead based balancing step.

Hi DocBio,

Could I check whether you ever perform mixed flow-cells with some lanes TruSeq and some Nextera (not both in same lane)? We carried out a run last year with six lanes of TruSeq and 2 lanes Nextera XT, following the caveats about using the Nextera Sequencing Primer kit for the cBot/PE read, etc. However, despite good clustering of the TruSeq samples (we load at 10pM), we only obtained ~8 million clusters per lane for the Nextera samples. Illumina were at something of a loss to explain this. We didn't use the normalisation beads following library prep - the samples were individually diluted to 10pM (quantified using Qubit which works well for loading Nextera libraries on our MiSeq). I should note that we typically see equivalent cluster densities for both library types at 8pM loading on the MiSeq.

Any help appreciated as we have a user wishing to do a similar run in the next week or so!

Matt

[docbio]

Matt,

Sorry I can't be of much help. I'm sending my samples to a core facility and they flat out refuse to mix chemistries on a flow cell. I had been led to believe that it wasn't even possible to do what you're describing due to limitations of the cBot, but it seems you've found a strategy Illumina endorses. Not sure what the problem would be, unless there is some inherent different in the hybridization affinities between the two adapter types or some difference in the insert sizes that resulted in differential clustering of the two library types due to different concentrations of ends at a given pM concentration. It's late so I'm not even sure if that makes sense...

I do know Illumina says the MiSeq clusters more efficiently than the HiSeq, but you probably know that already.

Sorry I can't be of much help... if you figure out a solution post an update because I'm curious.

Best,
DocBio

[koadman]

Following up on my earlier posting I want to warn others of some problems we encountered with dual-index sequencing on the hiseq and how we resolved them to ultimately arrive at a good result. Our first attempt with a single lane gave encouraging results, so we loaded up the better part of a flowcell with dual indexed samples. Unfortunately the 2nd index failed to read on that run (and yes it compromised much of the planned analysis). Most of the 2nd index reads were of extremely low quality and those that did read were inside the adapter downstream from the first barcode. After much discussion with Illumina tech support we discovered that the NaOH which is used to denature the DNA prior to the 2nd barcode read had apparently neutralized during the several days of the first read, and this resulted in failure of the 2nd barcode. We attempted another run, replacing the NaOH with fresh solution before the 2nd barcode read and indeed, this seems to have resolved the problem. Illumina tech support said this issue may be related to the particular environment at our facility (too much sunshine in Davis?) and that not all users will experience the problem.

So, for those who are attempting dual index reads, if the 2nd barcode read quality is suspect it might be as simple as adding fresh NaOH!

[pmiguel]

After much discussion with Illumina tech support we discovered that the NaOH which is used to denature the DNA prior to the 2nd barcode read had apparently neutralized during the several days of the first read, and this resulted in failure of the 2nd barcode. We attempted another run, replacing the NaOH with fresh solution before the 2nd barcode read and indeed, this seems to have resolved the problem. Illumina tech support said this issue may be related to the particular environment at our facility (too much sunshine in Davis?) and that not all users will experience the problem.

So, for those who are attempting dual index reads, if the 2nd barcode read quality is suspect it might be as simple as adding fresh NaOH!

Eh? Why did the first index work then?
I mean does that make sense to you? NaOH sits around for days in the instrument during read1. Then it successfully denatures the first read to allow annealing of the index1 primer. But after the additional several hours it takes to do 8 cycles then it doesn't work?

--
Phillip