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Old 12-01-2015, 12:44 PM   #1
SunPenguin
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Default Double SPRI with Ampure XP for higher mw products

I'm been trying to sequence these PCR amplified products that are around 550-650 bp long. Extremely pesky to amplify. So far I've tried doing Ampure clean up (at 0.6x) and also gel purification (sybr safe, LMP agarose, etc.). Gel purification does clean up the product very well, but I always end up with just okay recovery.

The SPRI cleanup, on the other hand, gave me much better recovery, and actually cleaned out low mw products <300-350bp quite well. I do still end up with some higher mw minor products > 1k bp, however, that I don't want. Though I think for now, the high mw minor products shouldn't eat up too many of my reads, I would like to get rid of them if possible, without anymore gel purification.

I'm trying to figure out how I would do double spri to get rid of that product.

Something I noticed reading around is that it seems the SPRI bead protocol becomes weird when you try to go below 0.5X beadsroduct ratio; it seems the protocol misbehaves and doesn't have the mw cutoff you would expect (looking at the Broad material slide 61).

I was hoping I could do something like 0.4-0.3X SPRI to retain the 1kb products onto the beads, then take the supernatant and perform the regular 0.6X SPRI to get rid of lower mw products. I also realized, however, that the supernatant taken from the first SPRI cut into the second cut would still have leftover SPRI buffer/PEG. Would I have to change the ratio of the second cut to compensate for the difference?
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Old 12-01-2015, 01:17 PM   #2
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I think a 0.4X followed by a 0.6x should work. Your supernatant after the 0.4X will contain bead buffer and you will need to adjust the volume that you add for the 0.6X cleanup. If your sample is 50ul you would add 20 uL of beads for the 0.4X upper and then 10 uL of beads to the supernatant for the 0.6X lower.
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Old 12-01-2015, 01:55 PM   #3
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50uL would be exactly what I'm going for.

Could you go over the math here? is the volume here assuming i'm only taking over 50uL of supernatant from the first SPRI into the second SPRI?
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Old 12-01-2015, 02:34 PM   #4
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Originally Posted by SunPenguin View Post
50uL would be exactly what I'm going for.

Could you go over the math here? is the volume here assuming i'm only taking over 50uL of supernatant from the first SPRI into the second SPRI?
This is assuming that you are taking all of the clear supernatant over.
So...
1. Add 20 uL of beads to 50 uL of sample, incubate, magnetize, and transfer ~68-70 uL of clear supernatant. (20uL of this supernatant will be bead buffer)
2. Add 10 uL of beads to the supernatant, incubate, remove supernatant, magnetize, wash, resuspend, transfer clear sample.

Since the amount of buffer determines the selection and not the beads themselves, adding an additional 10uL of beads to the supernatant that is already has 20 uL of buffer should get you to around a 0.6X buffer to sample ratio (50uL of sample and 30uL total of bead buffer).

This second bead amount might need a little adjustment to get exactly what you want, but 10uL should be pretty close to what you're aiming for.

Last edited by jdk787; 12-01-2015 at 05:31 PM. Reason: Can't do math
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Old 12-01-2015, 03:23 PM   #5
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Here is a helpful diagram that I use when determining bead cleanup concentrations.
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Old 12-01-2015, 04:24 PM   #6
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Hmm, the graph again doesn't extend beyond 0.5, but the exponential (or polynomial) nature of the curves may explain that.
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Old 12-01-2015, 05:15 PM   #7
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That's quite helpful, Angelmass! Did you generate that yourself? I've been meaning to do a systematic test like that with AMPure XP beads.

SunPenguin, in my hands the recovery rate from SPRI double-sided size selection can be comparable to or less than what I get from a gel extraction. Highly dependent on your amplicons and their proportions, of course, but I try to avoid the Broad's method unless I have access to a liquid handler. Let us know if it works! Out of curiosity, can you potentially re-amplify your purified fragments now that your primers should be on-target?

Also, it sounds like your products might be right on the cusp for Zymo's Select-a-Size kit if your unwanted smaller fragments are consistently below 300bp. Maybe worth a shot?
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Old 12-01-2015, 05:16 PM   #8
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Quote:
Originally Posted by jdk787 View Post
This is assuming that you are taking all of the clear supernatant over.
So...
1. Add 30 uL of beads to 50 uL of sample, incubate, magnetize, and transfer ~78-80 uL of clear supernatant. (30uL of this supernatant will be bead buffer)
2. Add 10 uL of beads to the supernatant, incubate, remove supernatant, magnetize, wash, resuspend, transfer clear sample.

Since the amount of buffer determines the selection and not the beads themselves, adding an additional 10uL of beads to the supernatant that is already has 30 uL of buffer should get you to around a 0.6X buffer to sample ratio (50uL of sample and 30uL total of bead buffer).

This second bead amount might need a little adjustment to get exactly what you want, but 10uL should be pretty close to what you're aiming for.
I assume the 30uL of beads is a typo? It should be 20uL (20/50=0.4)? then assuming I take about ~70uL of the supernatant, then add 10uL of beads to make 80uL total, I should get the effects of a 0.6x SPRI ((10+20)/50=0.6)?
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Old 12-01-2015, 05:27 PM   #9
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Quote:
Originally Posted by adam.geber View Post
That's quite helpful, Angelmass! Did you generate that yourself? I've been meaning to do a systematic test like that with AMPure XP beads.

SunPenguin, in my hands the recovery rate from SPRI double-sided size selection can be comparable to or less than what I get from a gel extraction. Highly dependent on your amplicons and their proportions, of course, but I try to avoid the Broad's method unless I have access to a liquid handler. Let us know if it works! Out of curiosity, can you potentially re-amplify your purified fragments now that your primers should be on-target?

Also, it sounds like your products might be right on the cusp for Zymo's Select-a-Size kit if your unwanted smaller fragments are consistently below 300bp. Maybe worth a shot?
Hmm, that's interesting. I am in the process of trying to limit how many PCR cycles to use to avoid chimeric products, so while the back up plan is to take the product and amplify it by 10 more cycles (or maybe even less), it would be preferable to not do that (as would anyone, I suppose).

At this point, I generally get this low intensity smear of DNA just at the high mw level (900-1000bp all the way to 6000bp or higher). It actually shifts the baseline of bioA (I attached the trace here). I think since the products that I want are the shorter major product, I should be okay, but still, it would be cleaner if I could get rid of the unwanted stuff.
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Old 12-01-2015, 05:33 PM   #10
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Here's another less dramatic example.
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Old 12-01-2015, 05:33 PM   #11
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Quote:
Originally Posted by SunPenguin View Post
Hmm, that's interesting. I am in the process of trying to limit how many PCR cycles to use to avoid chimeric products, so while the back up plan is to take the product and amplify it by 10 more cycles (or maybe even less), it would be preferable to not do that (as would anyone, I suppose).

At this point, I generally get this low intensity smear of DNA just at the high mw level (900-1000bp all the way to 6000bp or higher). It actually shifts the baseline of bioA (I attached the trace here). I think since the products that I want are the shorter major product, I should be okay, but still, it would be cleaner if I could get rid of the unwanted stuff.
Oh, wow. That's definitely a shifted baseline! Is it possible you're already getting overamplification? The smaller hump at 900bp makes me think so. How many cycles are you doing to get your product? Edit: or possibly contaminating gDNA?
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Old 12-01-2015, 05:34 PM   #12
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Quote:
Originally Posted by SunPenguin View Post
I assume the 30uL of beads is a typo? It should be 20uL (20/50=0.4)? then assuming I take about ~70uL of the supernatant, then add 10uL of beads to make 80uL total, I should get the effects of a 0.6x SPRI ((10+20)/50=0.6)?
Sorry about that, I edited my post with the correct numbers.
What you have here is correct.
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Old 12-01-2015, 05:38 PM   #13
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I think that's definitely possible, but I don't have enough experience to know what that looks like.

The product goes through several round of PCR, with a SPRI/gel extraction between each step. The total number of cycles right now sit at 70 (it's a low abundance transcript, so it takes a bit to get enough products). The last step is 20 cycles. Would over amplification cause this kind of trace? Should I maybe reduce my template concentration?
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Old 12-01-2015, 06:28 PM   #14
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Quote:
Originally Posted by SunPenguin View Post
I think that's definitely possible, but I don't have enough experience to know what that looks like.

The product goes through several round of PCR, with a SPRI/gel extraction between each step. The total number of cycles right now sit at 70 (it's a low abundance transcript, so it takes a bit to get enough products). The last step is 20 cycles. Would over amplification cause this kind of trace? Should I maybe reduce my template concentration?
Overamplification could cause this kind of trace through formation of bubble product and daisy chains.
When primers are depleted during PCR, non-complementary strands can anneal to each other and show up as high mw products on the BA. If this is what's happening, reducing cycles or reducing template will help.

You can determine if this is what is occurring by running a portion of your library on a denaturing gel to see if the upper end stuff goes away.
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Old 12-01-2015, 06:46 PM   #15
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Quote:
Originally Posted by SunPenguin View Post
I think that's definitely possible, but I don't have enough experience to know what that looks like.

The product goes through several round of PCR, with a SPRI/gel extraction between each step. The total number of cycles right now sit at 70 (it's a low abundance transcript, so it takes a bit to get enough products). The last step is 20 cycles. Would over amplification cause this kind of trace? Should I maybe reduce my template concentration?
Can you describe your protocol with more detail, including what your DNA (or cDNA? you mentioned a transcript) sample consists of and what you're amplifying? I've never had to resort to repeated rounds of purification and PCR -- there might be a simpler way to do this by e.g. touchdown PCR.

jdk787's suggestion of a denaturing gel will definitely clear things up. You also might want to run your gDNA/cDNA sample out to see if you're getting high-molecular weight products that are beyond the range of the Bioanalyzer chip you're running. Alternatively, if you have access to a Tapestation the gDNA Screentapes are quite decent at resolving larger fragment sizes.
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Old 12-01-2015, 09:13 PM   #16
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Can you describe your protocol with more detail, including what your DNA (or cDNA? you mentioned a transcript) sample consists of and what you're amplifying? I've never had to resort to repeated rounds of purification and PCR -- there might be a simpler way to do this by e.g. touchdown PCR.

jdk787's suggestion of a denaturing gel will definitely clear things up. You also might want to run your gDNA/cDNA sample out to see if you're getting high-molecular weight products that are beyond the range of the Bioanalyzer chip you're running. Alternatively, if you have access to a Tapestation the gDNA Screentapes are quite decent at resolving larger fragment sizes.
It goes through multiple rounds because each round uses a different set of primers that are nested to each other. I start with a 5'Race RT (no repeat, just 1 cycle), and then two rounds of nested PCR, then 1 final round of PCR to attach Illumina seqeuencing handles (as opposed to ligation, which in my hands hadn't been efficient).

Right now I've definitely not optimized all the parameters. Library prep is definitely still a bit difficult for me to grasp.

Hmm I think it definitely could be overamplification, since it does seem on the samples that I start with lower concentration of template (just by checking gel band intensity) I tend to have less of this kind of issue. The qPCR results for these sample wasn't super high though, so it's a little hard for me to imagine the reaction being depleted of primers (it's added to the reaction at 0.5uM). Maybe I'm somehow losing amplified product to annealing due to over cycling?

I'm also using Phusion hot start for this reaction. I'm wondering maybe that has something to do with it? perhaps not enough Mg?

Last edited by SunPenguin; 12-01-2015 at 09:16 PM.
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Old 12-02-2015, 06:02 AM   #17
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Originally Posted by SunPenguin View Post



The qPCR results for these sample wasn't super high though, so it's a little hard for me to imagine the reaction being depleted of primers (it's added to the reaction at 0.5uM). Maybe I'm somehow losing amplified product to annealing due to over cycling?
What is the molarity of your final product? Your BA traces look like your libraries are pretty concentrated?

Also, is the final concentration of primers in your reaction 0.5uM or are they 0.5uM before you add them? If they are 0.5uM in the reaction (2uL of 12.5um in 50 uL) I think you should be ok depending on how much material you have going in to the PCR reaction.
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Old 12-02-2015, 06:14 AM   #18
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The graph is consistent with my experience; 0.4X isn't going to work and even 0.5X is dodgy. For the record, you really should recalibrate your SPRI conditions for your specific reaction mix, because certain components (like magnesium, or especially PEG from a "quick ligation") change the binding chemistry.

But then your Bioanalyzer trace looks perfect except for being overamplified.

Instead of a denaturing gel, you can also denature a sample yourself (a couple of minutes at 95 C) and then run it on a Bioanalyzer RNA chip to profile the ssDNA. This won't get artifacts from daisy chains, bubbles, or whatever overamplification causes.

Also FYI Phusion has been obsolete for some time now; NEB uses the new Q5 polymerase in its library kits, and all the cool kids are using Kapa HiFi, even for amplicons in the 10+ kb range.

Titrate that PCR. Just make a bunch of duplicate tubes and take one out after each cycle, then purify per usual and run on the Bioanalyzer (DNA chip). I bet you'll see that a few cycles lower gives you just as big a peak in the desired range and none of the long tail. FYI overamplified libraries sequence just fine; the only problem is that you can't believe the Bioanalyzer numbers to quantify them, but qPCR doesn't care.

Last edited by jwfoley; 12-02-2015 at 06:19 AM. Reason: more information
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Old 12-02-2015, 06:32 AM   #19
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The total number of cycles right now sit at 70 (it's a low abundance transcript, so it takes a bit to get enough products).
Is this a typo? Even starting from a single molecule, 70 PCR cycles would give you about 21 g of DNA, according to a quick back-of-the-envelope calculation, though of course you'll exhaust the reagents first (and primers are usually the first thing to go, by design). I'll grant you some loss in the intermediate steps, but still. Overamplification definitely sounds like a possibility.
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Old 12-02-2015, 06:58 AM   #20
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What is the molarity of your final product? Your BA traces look like your libraries are pretty concentrated?

Also, is the final concentration of primers in your reaction 0.5uM or are they 0.5uM before you add them? If they are 0.5uM in the reaction (2uL of 12.5um in 50 uL) I think you should be ok depending on how much material you have going in to the PCR reaction.
The final concentration of the primers are 0.5uM. BioA estimates about 5-10nM on these peaks, and qPCR estimates about 10-20nM. I don't think that's super concentrated?

Edit: actually I looked at the traces again. The ones that show more of a shoulder have 5-10nM estimated by bioA. The ones that have less of a shoulder shows upward of 50-60 nM by bioA.

Last edited by SunPenguin; 12-02-2015 at 07:00 AM.
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