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Hello,
First at all thank you for these great protocols!
I would like to try your BOMB protocols for our nucleic acid isolation and maybe use it as our main methods in the near future.
However I would like to ask a few questions before I start.
We don’t really have the means to synthesize the beads, since you require kinda anaerobic conditions. However I already bought the Sera-Mag SpeedBeads cat. 45152105050250 which you recommend in the paper, but I am not sure how I should use these.
My first question is if we need to re-suspend the beads in other buffer or we can use the beads in the buffer in which they arrive?
Second, can we treat this bead solution as the “stock” solution in your protocols. Or we should adjust the amount of beads in each protocol? If we need to adjust the amount of beads, by how much we should?
Lastly, regarding this sentence from the protocol #4.3 “Silica-coated BOMB beads in TE(1:10 diluted from stock) Or, 1:50TE diluted carboxylate-coated magnetic beads can be used; for example, purchased from GE Healthcare” . My question is, should we dilute the beads 1:50? or you refer that the stock which we get from GE healthcare is already 1:50 diluted compared to your stock?
Thank you for your help,
Sebastian
Hi Sebastian,
Apologies for the slow reply here! The buffer the Speed Beads arrive in is not suitable for DNA handling – it contains Sodium Azide and is much too concentrated. What you need to do is take 1 mL of the resuspended beads as they arrive, wash 2 x in TE buffer (i.e. magnet, remove Aizde, resuspended in 1 mL TE, magnet, remove TE, add 1 mL TE, magnet and remove TE). Finally resuspend the beads in 50 mL of TE buffer (for DNA/RNA/TNA extraction) or 50 mL of PEG binding buffer (for cleanup/size exclusion).
Cheers,
TimH
Hi Tim,
I did exactly what you said, I diluted them 1:50, since many other protocols suggest the same thing. Also the washing was written on other protocols which I found on the internet.
Now I have some other questions specifically related to carboxyl beads. While the clean-up works perfectly and it is a good protocol, the other two protocols which I wanted to implement in our lab don’t work well with carboxyl coated beads.
The first one, the plasmid purification #5.1. For me it recovers kinda 1/3 of the amount compared to a commercial kit. I understand, your protocols are not optimized for carboxyl coated beads, so I am not surprised. However, I think there are several reason why this doesn’t work well.
1. The beads tend to clump in buffers which contain guanidine, and they don’t re-suspend in the wash buffer, they stay as small clumps, but never a uniform brown color as it should be when the beads are re-suspended. However, the beads can be re-suspended in the EB buffer, in the last step.
2. I think for the carboxyl beads the binding time should be larger. Also the volumes should be slightly lower to accommodate things in one microcentrifuge tubes. My experience (with ampure beads not specifically these ones, and for nanopore libraries) is that lower volumes and higher binding and elution times drastically increase the yield.
With these two things in mind I did a protocol for plasmid which uses a modified recipe for beads from the cleanup/size exclusion (so NaCl and PEG) and removed the guanidine. And I recover consistently around 80% of the plasmid compared to a commercial geneaid kit for plasmid. Eg. for a 5 mL LB culture I got 390 ng/ul (nanodrop) in 50 ul of EB compared to around 460 ng/ul with the geneaid kit. However, this is good enough for us. If you want I can share with you the protocols and test it with your synthesized beads and maybe publish it on the webpage for the people.
The other protocol which I tried to use is the gel purification. However, here I never managed to make the protocol work. First, using PEG does not help, if I actually use PEG and NaCl buffer, the beads form a sort of slime with something from the tube and they are really hard to work with.
With your original protocol my recovery is very low (speaking about large fragments above 600 bp) and I have a lot of salt contamination. No matter how much I wash (I tried 4 washes). Here I also notice that the beads form a clump, but this time around, I am literally unable to re-suspend the clump at all. They stick together no matter what. Again, my suspicion is the GITC in the buffer, but also somehow I think the agarose it is an issue, I think somehow the agarose can bind carboxylated beads (I know that in some nanopore protocols during the sequencing the pores die quickly and people think is a polysaccharide carryover, even if during the library prep you have 2-3 cleanups with ampure xp beads). Because of this experience with nanopore, I think that also agarose might bind to the beads and this is why I see this solid compact beads pellet. Interestingly, in some reactions during testing this did not occur, and I have no idea what was the case there.
Did you guys observe these issues in gel purification during testing of the gel extraction?
Thank you,
Sebastian
Hi Sebastian,
Many thanks for the detailed feedback. I have tried to split the two main challenges you raise so it is easier for others to follow. However, due to the common issue of clumping, I suspect the answer for both is the same (you need to more beads for capture, or less starting material):
PLASMID EXTRACTION
You are right, most of the protocols are optimised for the lab-made ‘BOMB’ silica beads, so how well the carboxylate “Speed Beads” work in various situations has not always been tested or optimised. Plasmid extraction is one of those.
Nevertheless, I have done a lot of DNA/RNA/TNA extraction from cultured cells and tissue using the carboxylated speed beads and guanidine buffers. The only time I see clumping is when there is a lot of DNA/RNA/TNA due to excess input material. What I assume happens is that instead of the nucleic acid just precipitating to the beads in solution, the beads and nucleic acid form one giant complex (almost as if the nucleic acid ‘captures’ the beads instead of the other way around). This bead complex is now effectively saturated, and unable to collect all of the nucleic acid in the solution, hence one reason why you are not getting the yields you expect to see. There are two things I think you could do to improve this: (1) use less starting material (I suspect you will get less clumping as beads won’t saturate) and (2) scale up your extraction if deep well plate will allow – more beads/volume will mean greater capacity to capture nucleic acid.
GEL EXTRACTION
I am not sure if we ever tried doing PEG/NaCl cleanup from a solution that contains agarose, but I imagine it would make for quite a viscous/slimy solution as you describe…between the PEG crowding agent and melted agarose this is perhaps to be expected.
When using the original protocol (#4.3), you mention more clumping. Again, we did not see this in our experiments, and I suspect this is part of the reason why you have a lot of salt contamination. The dense clumps tend to trap GITC/salt, and are not easily washed.
Have you tried adding more (TE-diluted) Speed Beads to the solution? The protocol suggests to add 5-10 uL of STOCK SOLUTION ‘BOMB’ beads (i.e. undiluted), however, if you added 5-10 uL of DILUTED Speed Beads, you may not have enough.
Would be great if you can post back the result so we can help other users trying to use speed beads for plasmid/gel extraction.
Cheers,
TimHi Tim,
Plasmid extraction.
At least for the plasmid purification I don’t think the beads amount is an issue. I used the same amount of beads for both your #5.1 protocol and my current protocol. The only difference is that in my protocol I don’t have GuHcl in the N3 buffer, but instead is the N3 buffer from #5.2 (2.3 M KoAc). And for the binding (in my protocol) I make sure I have a 1:1 ratio between the supernatant recovered and the PEG buffer . I did both of these protocols in parallel and in the #5.1 I always see clumping (but the clumps can be broken is smaller pieces, however they can never be re-suspended in the PE wash buffer) as opposed to the one with PEG where I just vortex them and the beads form nicely a brownish solution with the PE wash buffer. Since I use the same amount of starting material for both protocols, the same culture, same plasmid, etc. I don’t think the issue is the amount of material. This is why my suspicion is towards the guanidine.
My experience with clumping is very vague, and everything relies on AmpureXP beads and preparation of libraries for nanopore sequencing. The only time I saw clumping was when I tried to use high molecular weight DNA to prepare a nanopore library as it was mentioned also here. http://seqanswers.com/forums/showthread.php?t=47522
Based on nanopore support, clumping is because the long strands of DNA bind tightly to the beads. Also because of this reason, eluting HMW DNA from carboxylated beads is hard, requires larger elution times and temperatures at around 65C (https://www.researchgate.net/post/How_can_I_fully_elute_whole_genomic_DNA_off_of_Sera-Mag_paramagnetic_beads ).
But this is not the case here, as the plasmid is 8kb or less.
Another reason why I don’t think that the bead amount is the issue is the following. I used the modified protocol by me for a Midiprep. In this case I used 70 mL of culture (exactly the same culture as above, this is why it is such an odd number) so roughly 14 times more than for a miniprep. But I increased the beads amount just 5 times. Yet the midiprep worked perfectly (785 ng/ul in 300 ul of elution buffer).
Somewhere, some people mention (no scientific citation for this) that the binding capacity of 1:50 diluted sera-mag beads is around 7 ug/ul of diluted beads. Yet I don’t have any citation for this.
GEL extraction:
I have a more concentrated beads solution for the Gel extraction 1:12.5. So 4 times more concentrated compared to what dilution is standard. I did not try to increase the volume because I think the isopropanol ratios are critical for the protocol. Either way the same result. At this stage honestly I don’t have time to look into this protocol and figure out why it does not work in my hand. The beads are visibly starting to clump the moment when I add to the dissolved agarose in GITC. I was thinking to add first the isopropanol, then to try to add the beads. I will see if I will have time to test this.
I think that sera-mag beads are not behaving exactly as your synthesized beads, and maybe this is why we get this issue with them.
Best regards,
Sebastian
Hi, first of all thanks so much for all the amazing work you’ve done.
I have a question regarding the Speedbeads you suggest (45152105050250) vs what is suggested in the Openwetware protocol (65152105050250). Yours are hydrophilic, theirs are hydrophobic and both PEG/NaCl binding buffers are basically the same. I’ve tested hydrophilic vs hydrophobic Speedbeads together with Ampure XP beads in RNA purification and in enzymatic cleanup, and the hydrophilic beads clump in PEG buffer but the hydrophobic does not.
I know you protocols suggest binding RNA in guanidine buffers with isopropanol/ethanol but that seems like a bad idea for downstream NGS applications. Can you comment on this behavior, or suggest a way around of using the hydrophilic Speedbeads in RNA purification without guanidine buffers?
Hi Bernardo – sorry for the slow reply – we haven’t done a huge amount of testing using the hydrophilic beads. Did you get any further with this? We have done plenty of NGS (Illumina mostly) using GITC and the hydrophobic Speedbeads – it doesn’t appear to cause us issues….could you clarify if its an issue for you? As above, we usually only get clumping when a lot of input tissue/cells is used and high-molecular weight DNA is used – but others (e.g. Sebastian above) have had clumping issues when extracting plasmids.
Cheers, Tim H
Hi Tim and Sebastian,
Thank you for the amazing work. I’m attempting plasmid purification with Sera-Mag beads and Tim, I would love to have your modified protocol that doesn’t involve guanidine as a reference.
Best,
Shirley J
Ahoy,
With the forum having grown rather quiet over the last year I suspect Sebastian might not see your request. And while I am not willing to dox him I dare to say that you might be able to find his e-mail via google.
Anyhow I can also suggest the following as a binding/extraction buffer which has worked very well for me:The Urea+LiCl lysis and binding buffer recipe for 50 mL :
6M Urea (18g Urea)
3M LiCl (6.3 g LiCl)
500 mM TrisHCl (2.5 mL of 1M stock, pH 8)
30 mM EDTA (2 mL of 0.5M stock)
0.5% Tween20 (250 uL)I used this to replace the standard BOMB GITC buffer 1:1.
This buffer is a lot cheaper, less toxic and performed about as well as the GITC variant. There are two main downsides to this buffer compared to GITC, it does not keep as well, after about a week at room temperature I would always find it to start performing somewhat worse and at some point it would develop a shiny film on the surface and it has a tendency to precipitate when room temperatures are cool. I believe the short shelf live to be some form of Urea deterioration and not microbial contamination but I never tested it as the tiny extractions I perform require so little of the GITC buffer with its nearly infinite shelf life that it was just not worth switching to this alternative permanently.
In case you feel like this will be difficult to acquire, ask someone in a lab which has been doing RNA work for a while, there is a good chance the have high concentration LiCl solutions, as for Urea, it is even cheaper than LiCl.
Cheers
Tim M
Hi,
I am not sure what you mean by your request, as all plasmid isolation protocols that involve Sera-Mag beads, do not involve guanidine salts.
Guanidine salts don’t play nicely with the Sera-Mag beads, this is why in all protocols that I provided with Sera-Mag you will not find any guanidine salts as components of the buffers.
Best regards,
Sebastian
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