Miniprep without EDTA

[Mega]

Hello @all,

I was wondering if I could do a dirty miniprep without EDTA.

What I have: weak centrifuge, SLS (sodium lauryl sulfate), NaOH, vinegar, ethanol (I have access to both wodka and pure ethanol), water.


Can you do a mp with this limited resources?
What I want: Some ~60 to 70 % plasmids, some remaining proteins, RNA, mabe some fragments of chromosomal DNA.


I think with this percentage you can still do a transformation with E.Coli? Because Proteins will either be used or digested, RNA my do it's job inside the cell, chromosomal DNA has no origin of replication.

Only the plasmids will be able to replicate and thus you will get some transformants. These you select with ampicillin anyway. (Clearly, you won't get so much transformants, but who cares? )

Will DNAses inside the solution possibly destroy the plasmids??

[Cory Tobin]

> I was wondering if I could do a dirty miniprep without EDTA.

It will work without EDTA.

> What I have: weak centrifuge, SLS (sodium lauryl sulfate), NaOH, vinegar,
> ethanol (I have access to both wodka and pure ethanol), water.

Make sure the vinegar is distilled white vinegar. Also you'll need
potassium chloride, sodium chloride and diatomaceous earth.

> Can you do a mp with this limited resources?
> What I want: Some ~60 to 70 % plasmids, some remaining proteins, RNA, mabe
> some fragments of chromosomal DNA.

I'm not sure what you mean by 60%. But I've been able to get 200ng/uL
using only these reagents.



-cory

[Andreas Sturm]

I mean it doesn't have to be pure.
It must not be degraded by DNases. But it should work for a transformation.


distilled white vinegar? Have to look about that.

potassium chloride, sodium chloride and diatomaceous eart ?
What do I need these for? Do you have a protocol for the whole miniprep? That would be really great!!

2012/5/27 Cory Tobin <cory....@gmail.com>

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[Cory Tobin]

> potassium chloride, sodium chloride and diatomaceous eart ?
> What do I need these for? Do you have a protocol for the whole miniprep?
> That would be really great!!

http://wiki.biohackers.la/Miniprep


-cory

[Andreas Sturm]

Amazing, thanks!!!!!

Is the KCl relly necessary? Can't be replaced by NaCl?


What about Ethanol instead of Isopropanol?
Feasible?


(lye is NaOH, right? )

2012/5/27 Cory Tobin <cory....@gmail.com>

-cory

[Cory Tobin]

> Is the KCl relly necessary? Can't be replaced by NaCl?

You need a potassium ion. In the commercial miniprep kits (ie.
Qiagen) they use potassium acetate, but you can only get that from
chemical suppliers. You can get KCl at the hardware store. It is
used in water softener tanks. I actually found it at a grocery store.

> What about Ethanol instead of Isopropanol?
> Feasible?

Yes, I used isopropanol because I thought people would have easy
access to 100% isopropanol whereas 100% ethanol is more difficult to
find.

> (lye is NaOH, right? )

Yes.


-cory

[Cathal Garvey]

It's really easy, but somewhat hazardous, to make your own potassium acetate. You can get potassium hydroxide cheaply online, and titrate a solution oud it with acetic acid (vinegar).

Two things about vinegar:
One, 'distilled' vinegar isn't itself distilled: it means, rather, that it was fermented using a very rapid fermentation process from distilled ethanol. So, there are fair odds that it's not a pure or sterile product. However, in a miniprep that's probably not a big issue.

Two, vinegar is about 5%, pretty weak for titrating etc.. But, I gather you can easily get stronger acid by boiling off some water, because the boiling point of acetic acid is slightly higher and it doesn't form an azeotrope with water. Once the concentration is high enough, you can also crystallise pure acetic acid from solution by putting it in the fridge. However, BE WARNED: high-purity acetic acid is a pretty strong acid, and can burn skin, eyes or lungs, or react vigorously and perhaps dangerously with other chemicals. Please don't 'titrate' by dropping crystals of KOH in it, for example!

Cory: thanks for the protocol! Can't wait to follow that link on my laptop. License?

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[Cory Tobin]

> Two things about vinegar:
> One, 'distilled' vinegar isn't itself distilled: it means,
> rather, that it was fermented using a very rapid fermentation
> process from distilled ethanol. So, there are fair odds that
> it's not a pure or sterile product. However, in a miniprep
> that's probably not a big issue.

Yeah, the only reason I said to use distilled white vinegar is because
I tried a bunch of different vinegars and this type worked the best.
In terms of maximizing 260:280, they all worked relatively well, but
the distilled white also maximized the 260:230 which means there was
less carbohydrate contamination.

Pure acetic acid was the best (slightly higher DNA concentration and
less carbohydrate) but I wouldn't recommend anyone use that without
proper protective gear and ventilation. - preferably a fume hood.


> License?

Just added a CC BY 3.0 license on the page.


-cory

[Cathal Garvey]

Awesome detail, and thanks for the license Cory!

[Cory Tobin]

One more note about the KCl: the KCl is only used to precipitate the
SDS which subsequently traps the cell debris including the genomic
DNA, allowing it to be separated by centrifuging (sodium dodecyl
sulfate is soluble in water whereas potassium dodecyl sulfate is
insoluble). The NaCl is used in the next step along with the vinegar
to bind the DNA. DNA will bind to silica (diamonaceous earth) when
NaCl > 2M and 4 < pH < 6*. If you could find a way to lyse cells and
remove the debris without SDS then you could eliminate the KCl.

Regarding the vinegar: in this protocol the only purpose of the
vinegar is to get the pH down to 4.5. In commercial kits, the acetic
acid forms a buffer with the potassium acetate (weak acid + conjugate
base), but I've found the pH range in which DNA binds silica is quite
large so getting a precise pH is not very critical. I tried
substituting HCl since you can get pure HCl (37% in water) from
swimming pool supply stores under the name "muriatic acid" but since
HCl is a strong acid, getting the pH within the acceptable range
without a buffer was complicated. Other weak acids might be
acceptable substitutes for vinegar.

Finally, an anecdote regarding the NaCl:

All of the commercial kits I am familiar with use guanidine
hydrochloride to make the DNA bind to silica. Most of the literature
on DNA purification revolves around the use of chaotropic salts like
guanidine hcl, but those chemicals are expensive, often toxic and are
only available through chemical distributors.

One time the lab that I work in had received a shipment of defective
Qiagen kits, so instead of waiting for Qiagen to fix the problem I
mixed my own solutions using guanidine hcl. I noticed the solutions
with guanidine had properties similar to water saturated with NaCl,
like low surface tension and low adhesion to plastic (while pipetting
the solution tends to drip out of the tip instead of staying in the
tip during transfer). So I Googled/Pubmeded for any reports of NaCl
being substituted for guanidine and found only a single paper (Lacksmi
et al, 1999, Analytical Biochemistry) that used NaCl for minipreps.
Apparently the use of inexpensive NaCl has been overlooked by the
miniprep kit manufacturers. Sucks for the thousands of labs who have
been overpaying for guanidine for the last decade, but good for
DIYers.


*The pH 4-6 range is somewhat nebulous. I haven't explored the pH
dependence thoroughly so the boundaries might actually be somewhat
larger or smaller than that.


-cory

[Cathal Garvey]

It's also a powerful denaturant, so having guanidine in the lysis/neutralisation buffer is handy for that reason, too. Helps kill tough proteins like nucleases, and reduces waiting time. Another common chaotropic salt is urea, which should have very similar properties to guanidine hcl.

However, for convenience nothing beats salt! That's an awesome hack/discovery I wasn't aware of, thanks!

[Andreas Sturm]

So tomorrow I'll go get some diatomaeous earth and some KCl....


When I do it without a liquid media overnight culture (from agar directly), how many bacteria do I have to take with the inoculation loop?

[Cory Tobin]

> When I do it without a liquid media overnight culture (from agar directly),
> how many bacteria do I have to take with the inoculation loop?

I've never heard of anyone doing a miniprep from bacteria growing on
solid media. Why not do a liquid culture?


-cory

[Nathan McCorkle]

Need DIY broth medias for mammalian and bacterial cells.

Also KCl might be found as salt-alternative at food stores.

[Jeswin]

Minipreps use bacteria grown in liquid media. I guess there is a
higher density of cells in liquid so that you get better DNA
purification efficiency.

[Andreas Sturm]

Tried to get diat. earth at a store. They didn't have it.

Another store, the Lagerhaus (don't know if there's an equivalent in english, mainly farmers buy there, but also private people)  had already closed when I came from university.



do you think, in university we will have some of it? Anyway, tomorrow I'll ask there (although I don't know what university would do with it...)

2012/5/29 Jeswin <phill...@gmail.com>

[Simon Quellen Field]

Swimming pool supply stores have it.

Most swimming pools use diatomaceous earth filters.

You can also get food-grade on-line, even using Amazon.

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[Mega]

Was at a swimming pool store. In former times it was used, but now they sell nano powder.

I put the glowing bacteria into a marmelade jar containing some 200-300 mL of LB-Medium. It was glowing for some days but now it's not glowing anymore... I assume most of the bacteria are dead. Can I still use them for miniprep, or are the plasmids destroyed when doing apoptosis???

[John Griessen]

On 05/28/2012 01:14 AM, Cathal Garvey wrote:
> I tried
>>substituting HCl since you can get pure HCl (37% in water) from
>>swimming pool supply stores under the name "muriatic acid" but since
>>HCl is a strong acid, getting the pH within the acceptable range
>>without a buffer was complicated. Other weak acids might be
>>acceptable substitutes for vinegar.

Are you meaning complicated because no buffer, or because of needing to titrate
to get the right concentration?

[Cathal Garvey]

Answer: probably, but leave to lose for longer, and expect smaller yields.

Also, technical note: Apoptosis is a deliberate form of tidy cell suicide found in multicellular or at least eukaryotic organisms. Bacteria and archaea sometimes show evidence of orderly suicide, but it's not apoptosis. When they die by other causes and burst open messily, it's called 'lysing'.

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[Mega]

What do you mean by 'leave to loose'? not shaking it to keep the bacteria out of solution?

would it be much better to do a new liquid culture, 3mL or so, in a 50 or 100 mL tube? How can I make sure that they get enough oxygen without contamination? alu foil?

Good News: I asked @university, if I could get some KCl and d.earth. Now I have several gramms of both.

[Cory Tobin]

>>> substituting HCl since you can get pure HCl (37% in water) from
>>> swimming pool supply stores under the name "muriatic acid" but since
>>> HCl is a strong acid, getting the pH within the acceptable range
>>> without a buffer was complicated.  Other weak acids might be
>>> acceptable substitutes for vinegar.
>
>
> Are you meaning complicated because no buffer, or because of needing to
> titrate
> to get the right concentration?

It is complicated because there is no buffer and we are essentially
titrating a strong base (the lysis solution contains NaOH) with a
strong acid, so once the NaOH is neutralized the pH quickly drops
below 2 which is too low for the DNA to bind the silica. We need the
pH to get in the 4-6 range. If you look at this titration curve
http://www.files.chem.vt.edu/chem-ed/titration/graphics/titration-strong-weak.gif
showing the titration of HCl and acetic acid against NaOH, there is a
lot more room for error in the acid concentration when using acetic
acid. It's possible to do it with a strong acid but your measurements
would have to be very precise or you would need to use a buffer.

-cory

[Nathan McCorkle]

On Wed, May 30, 2012 at 5:52 AM, Mega <masters...@gmail.com> wrote:
> Was at a swimming pool store. In former times it was used, but now they sell nano powder.
>
>
> I put the glowing bacteria into a marmelade jar containing some 200-300 mL of LB-Medium. It was glowing for some days but now it's not glowing anymore... I assume most of the bacteria are dead. Can I still use them for miniprep, or are the plasmids destroyed when doing apoptosis???
>

First do you mean glowing or fluorescing, we need to be consistent
with terminology here.

I've purified GFP protein from cell lysate, and it still fluoresced,
so it doesn't depend on cell's being alive, but certainly a lysed cell
would have proteases all over the place and the GFP could then degrade
rapidly


--
Nathan McCorkle
Rochester Institute of Technology
College of Science, Biotechnology/Bioinformatics

[Andreas Sturm]

Glowing. Bioluminescence.

They produce no more luciferin, so they are dead. Or dying.

2012/5/30 Nathan McCorkle <nmz...@gmail.com>

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[John Griessen]

On 05/30/2012 12:21 PM, Cory Tobin wrote:
> If you look at this titration curve
> http://www.files.chem.vt.edu/chem-ed/titration/graphics/titration-strong-weak.gif
> showing the titration of HCl and acetic acid against NaOH,

I see what you mean about the steep change, then a plateau.

They sell 9% vinegar at supermarkets for pickling and clothes wash additive.

[Nathan McCorkle]

and also tartaric acid "cream of tartar"

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[Andreas Sturm]

My NaOH 'powder' is in a bad container, and now it has sucked water.... Now it's a very pure NaOH solution...
Have to get some new.




BUT: I succeded in getting a professional miniprep kit. I'd like to use it now.

I recently got a Dremelfuge from a friend who built his own 3D printer. As an inauguration present ;) 
1) Will it be spinning fast enough to do a miniprep (Get most of the bacteria out of soultion)??  I once tried it, but it gave very little yield. But maybe, the lb marmelade jar was not full of of bacteria (althought they didn't glow anymore, so lysis?)

2) In which container shall I breed the 3-5 mL LB solution for the bacteria??
They will need oxygen, so above the tube shall be space. I have e.g. those https://encrypted-tbn0.google.com/images?q=tbn:ANd9GcQ8Q34n7YkKUL1gx99j4OL3efA0_NN9AgO2BHv28QgmxIfdT0w8 (the right one, 15 mL)

Is that sufficient?

3) Do they need 37°C and shaking? I don't have a device that heats or shakes them. But I have a hacked play-station controller which shakes when I apply a voltage. I could tape it to the controller and let it shake (never tested it for more than 1/2 an hour, but I'm confiden't it won't blow up)
How will I know they are ready for being harvested?


4) Shall I get some ampicillin for that? When they grow at 37°C, they will grow faster thus losing plasmids more poften. When I keep them at room temperature, they will lose much less plasmids I read. The yield may decrease, but will it be significantly?

[Cathal Garvey]

> I recently got a Dremelfuge from a friend who built his own 3D printer. As
> an inauguration present ;)

> 1) *Will it be spinning fast enough to do a miniprep* (Get most of the

> bacteria out of soultion)?? I once tried it, but it gave very little
> yield. But maybe, the lb marmelade jar was not full of of bacteria
> (althought they didn't glow anymore, so lysis?)

That depends. As always, and especially because you printed your own, I
can't offer any suggestions or guarantees of safety. But, I find that
5-10 minutes on the second setting of a Dremel 300 is enough to pellet
bacteria, although if you're in a hurry you can do it faster at setting
3-5. Remember that the longer/harder you spin them, the harder the
pellet will be to resuspend, and the higher the chances are that you'll
kill/damage whatever you're trying to pellet/resuspend.

But yes; Dremelfuges are easily able to pellet bacteria.

> 2) *In which container shall I breed the 3-5 mL LB solution for the
> bacteria??
> *They will need oxygen, so above the tube shall be space. I have e.g. those

> https://encrypted-tbn0.google.com/images?q=tbn:ANd9GcQ8Q34n7YkKUL1gx99j4OL3efA0_NN9AgO2BHv28QgmxIfdT0w8(the
> right one, 15 mL)
>
> Is that sufficient?

A wide & shallow-bottomed container is perfect if you have a jar or
erlenmeyer flask, but if you can shake them then a test tube or similar
is fine too. If you're using a deep container like a test tube, and you
don't have a shaking incubator, try to keep the volumes low and keep the
tubes slanted so that there's maximum air-liquid exposure.

> 3) *Do they need 37�C and shaking?* I don't have a device that heats or

> shakes them. But I have a hacked play-station controller which shakes when
> I apply a voltage. I could tape it to the controller and let it shake
> (never tested it for more than 1/2 an hour, but I'm confiden't it won't
> blow up)

> *How will I know they are ready for being harvested?

If you have access to 37C and shaking, that's perfect. But, I grow my
E.coli at 30C without shaking, and I still get good yields on a
miniprep. You just need to make sure they have plenty of air-broth
contact by inclining them or by only incubating in erlenmeyer flasks
with ~3mm of liquid at the bottom.

Vibration, as from a controller (awesome that you're using a PS
controller! :P), will certainly help to aerate the cultures by agitating
the water (preventing stratification of high-medium-low oxygen fluid)
and creating more surface area due to standing waves. So yea, vibration
is probably good. But do test the motor for overheating; vibrator motors
in phones and games consoles aren't usually designed for extended use.
There is a commonly available sort of vibrator motor that *is* designed
for extended use though.....that's right, massage wands! :D Glad we're
all on the same page!

> 4) Shall I get some ampicillin for that? When they grow at 37�C, they will
> grow faster thus losing plasmids more poften. When I keep them at room
> temperature, they will lose much less plasmids I read. The yield may
> decrease, but will it be significantly?

If you're using a conventional cloning plasmid, you'll need to apply
selection constantly, as they are usually missing DNA regions critical
for stable reproduction in wild plasmids.

For good yields, I recommend "rinsing" the cells you will be inoculating
with in plain broth first. I've seen this recommended by others as a way
to remove any beta-lactamase enzyme that they've secreted into their old
medium; the enzyme genes may be dormant by the time you harvest them for
inoculation of another culture, but the enzymes that remain can quickly
destroy any ampicillin in your new culture, reducing stability and
plasmid yield.

So, suspend the cells in new LB buffer, gently vortex or mix, and
centrifuge back out of that buffer before putting them into your new
Ampicillin-LB for maximum yields.

You don't need to do this for routine stuff, like subculturing a strain
just to keep it stable. It's just a good idea if you want high plasmid
yield, which I think you want.

If you *really* want good yields, you could consider using a different
broth: LB is actually very poor for plasmid yield. TB ("Terrific Broth")
is a common substitute using similar ingredients, which gives better
yields (I am told).

Most basic thing to bear in mind: don't use old cells. Use them while
they're growing happily in exponential-phase. Because you're using pVIB,
this is easy: harvest them for a miniprep when they are glowing at their
brightest, or just after their peak brightness. That indicates lots of
cells in exponential phase, which is just what you want. The moment you
let them fall into stationary phase (old) growth, your yields will drop
significantly.

Good rule of thumb: Harvest 6-8 hours after starting the culture. If
you're doing it overnight, start incubating them late and miniprep the
very next morning.

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PGP Public Key: http://bit.ly/CathalGKey

[Andreas Sturm]

>If you're using a conventional cloning plasmid, you'll need to apply
>selection constantly, as they are usually missing DNA regions critical
>for stable reproduction in wild plasmids.

Interesting. So natural plasmids are far more stable? A professor of mine always tells us that bacteria don't have luxury and so lose their plasmids kind-of-always.
I don't know if pVIB has a pUC origin or something similar. The Carolina homepage doesn't say.

>Good rule of thumb: Harvest 6-8 hours after starting the culture. If
>you're doing it overnight, start incubating them late and miniprep the
>very next morning.

6-8 hours also when 20°C room temperature??
Won't they need more time as grown-at-37°C-ones?

2012/6/7 Cathal Garvey <cathal...@gmail.com>

[Cathal Garvey]

> 6-8 hours also when 20�C room temperature??
> Won't they need more time as grown-at-37�C-ones?

No, I culture at 30C, which is still nice and warm. I do leave them
longer though, maybe 10 hours. When you get used to them, you can often
tell by the turbidity (cloudiness) and dispersion of a culture roughly
how "happy" it is. Exponential cells seem, to me, to "clear" less when
allowed to settle after a shake, probably because they're still actively
swimming around. On the other hand, stationary cells seem to me to be
more easily stratified by resting on a bench.

> Interesting. So natural plasmids are far more stable? A professor of mine
> always tells us that bacteria don't have luxury and so lose their plasmids
> kind-of-always.
> I don't know if pVIB has a pUC origin or something similar. The Carolina
> homepage doesn't say.

Not so! Plasmids wouldn't even exist in the wild if they were that unstable.

Basically, most of our plasmids use a mechanism called "Rolling circle
replication" to make copies of themselves. This mechanism involves a
step where the plasmids are single-stranded DNA, a very unstable state
for DNA to be in.

In wild plasmids, there's a non-coding region (often called a "Single
stranded origin" or SSO, but names differ) that folds into a special
shape that encourages transcription into mRNA by host ribosomes, but
only for 5-20 nucleotides or so. This mRNA acts as a "primer" for DNA
polymerases to replicate the other strand of the plasmid, stabilising it.

However, when we first started "hacking" wild plasmids to make early
cloning vectors, we just hammered in antibiotic resistance sites to make
manipulation easier, and then started cutting away whatever we could
remove while still having a functional plasmid. With
antibiotic-resistance genes and antibiotics forcing the cells to retain
even highly unstable plasmids, many or most cloning vectors of the time
ended up having their SSO's cut out, and because very unstable. Because
most modern vectors are simply re-hashes of older vectors with fancy new
features, they are similarly unstable.

This instability is one of the things I aimed to target with my own
plasmid, by deliberately including a highly stable SSO close to the
"normal" origin and Rep genes. The available literature suggests that
unless a gene is particularly bad for a bacterium (toxic gene products,
overly strong promoters, etc..), the plasmid shouldn't get lost just due
to growth and replication. You get less than 2% of cells without
plasmids after ~10-20 cycles of replication without antibiotic
selection, versus 80-99% for conventional plasmids.

I don't yet know if my plasmid matches this stability. That's one of my
remaining questions to answer! ;)

[Andreas Sturm]

Amazing!!!

Yeah, it sounds totally logical.

Plasmids have to be more stable if they wanted to survive Mio of years!!

I think we would never have learnt this at university. At least before graduating.  We recently did some theory of plasmids , but it  wasn't much at all...

2012/6/7 Cathal Garvey <cathal...@gmail.com>

[Mega]

Guys, I have 10% acetic acid here. Can I use it instead of the distilled white vinegar??

(For my diy - dirty - miniprep)

 

Am Sonntag, 27. Mai 2012 17:54:23 UTC+2 schrieb Mega:

Hello @all,

I was wondering if I could do a dirty miniprep without EDTA.

What I have: weak centrifuge, SLS (sodium lauryl sulfate), NaOH, vinegar, ethanol (I have access to both wodka and pure ethanol), water.


Can you do a mp with this limited resources?
What I want: Some ~60 to 70 % plasmids, some remaining proteins, RNA, mabe some fragments of chromosomal DNA.


I think with this percentage you can still do a transformation with E.Coli? Because Proteins will either be used or digested, RNA my do it's job inside the cell, chromosomal DNA has no origin of replication.

Only the plasmids will be able to replicate and thus you will get some transformants. These you select with ampicillin anyway. (Clearly, you won't get so much transformants, but who cares? )

Will DNAses inside the solution possibly destroy the plasmids??

[Nathan McCorkle]

More likely than not, is there a particular reason you ask?
http://en.wikipedia.org/wiki/Vinegar#Distilled_vinegar

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[Cathal Garvey]

I made my buffers according to Cory's protocol using 50% acetic acid,
diluting to 5% to match most commercial white vinegars in Ireland.

I'll share my results when I test the buffers soon. :)

Also, I'm rewriting the protocol in Markdown and I'll be sharing a few
alternative routes for people without access to some of the ingredients,
but who can source reagents to make them. i.e., if you can't buy KCl,
but you can get Potassium Hydroxide and Hydrochloric Acid (AKA Muriatic
Acid AKA "Spirits of Salt"), what amounts of each to react to get the
desired amount of KCl, etc.

[Cory Tobin]

> Guys, I have 10% acetic acid here. Can I use it instead of the distilled
> white vinegar??

I would dilute it down to 5%. 10% might work but I've had good results with 5%.


-cory

[Mega]

Ok. this was my question. whether 10 is better than 5 or 9%. Gonna do that. The protocol from your link will have calculated certain pH values. If I took 10% it may be too acidic?

[Cory Tobin]

> If I took 10% it may be too acidic?

I don't know. You can try it and see if it works. But it's not too
difficult to just mix it with water. 1 part water to 1 part 10%
acetic acid.

-cory

[Andreas Sturm]

So you don't have a hint/idea what could be better? So just try it with 5% (maybe I'll make ~5.5 to 6% by taking a bit less water)

I have pH indication stripes. I assume, after the neutralization solution has been added, pH should be ~7 (?). So I just check the pH and add more 5% acid?  (For step 7 of  http://wiki.biohackers.la/Miniprep )

And the binding solution? Does it matter so much? Just try it out?

2012/7/10 Cory Tobin <cory....@gmail.com>

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[Cory Tobin]

> So you don't have a hint/idea what could be better? So just try it with 5%
> (maybe I'll make ~5.5 to 6% by taking a bit less water)

The mixture of the lysis solution, neutralization solution and binding
solution needs to be in the range of pH4-6. I haven't really explored
the exact boundaries, so 4 and 6 are rough estimates. 4.5 works. 3
does not work. So the boundary is in between there somewhere.

> I have pH indication stripes. I assume, after the neutralization solution
> has been added, pH should be ~7 (?). So I just check the pH and add more 5%
> acid? (For step 7 of http://wiki.biohackers.la/Miniprep )

Actually, "neutralization solution" is a misnomer because it's
actually acidifying the mixture. The lysis solution is very basic
(~pH 11 I think) and the "neutralization solution" brings the pH down
to around 5. So maybe it should be called the "acidification
solution".

> And the binding solution? Does it matter so much? Just try it out?

Yeah, try it out and let us know what works. 10% might work. I never tried.


-cory

[Andreas Sturm]

At university we have a machine/device that checks the quantity of DNA (ng/uL). Haven't used it yet, but maybe this or next week.  I may use it when I need / want.

When the diy miniprep is done, I'll tell you what concentration I reached.

2012/7/10 Cory Tobin <cory....@gmail.com>

-cory

[Eugen Leitl]

On Tue, Jul 10, 2012 at 07:16:45PM +0200, Andreas Sturm wrote:
> So you don't have a hint/idea what could be better? So just try it with 5%
> (maybe I'll make ~5.5 to 6% by taking a bit less water)
>
> I have pH indication stripes. I assume, after the neutralization solution
> has been added, pH should be ~7 (?). So I just check the pH and add more 5%
> acid? (For step 7 of http://wiki.biohackers.la/Miniprep )

Buy a commercial pH meter with a glass pH electrode and
calibration buffers. Everything else is for the birds.

[Andreas Sturm]

I got some 0.5 molar EDTA from the lab. Can I add some mL to the lysis solution to boost efficiency?
Or would it be better added to the wash solution?

(With the lysis solution a part will stick to the cell wall, right?)

(

> Everything else is for the birds.

In my language, we say "for the cat" :D
                )

[Cathal Garvey]

Make Tris-EDTA with it and use that for eluting/redissolving the DNA
after the miniprep.

If you do your Miniprep in short time, there's no need to worry about
EDTA. Using it for storage of DNA *after* the miniprep makes more sense.

[Cathal Garvey]

You can pick up cheap glass-bulb pH meters on ebay as "pool pH testers".
My pH meter is one of these; a little red handheld battery powered unit
I got for maybe �15: commercial ones can run into hundreds, as usual.

You may need to calibrate it though, and for that I got little pH
standard tablets from a supplier; dissolve a tablet in 100mls deionised
water to get pH x, then calibrate the meter with a little screw at the
back until it matches the calibration sample.

There might be a way to easily make a calibration standard at home using
common ingredients, but I'd expect to be off by a few tenths of a pH
point. This is somewhat significant considering that it's a logarithmic
scale, but for rough-and-ready minipreps where your desired ranges are
between pH 2-4 and 8-10, the inaccuracy isn't that bad.

[Mega]

Here is a plastic bottle that says TE-EP redissolving buffer. This should be Tris EDTA?

 

I'm going to ask if I can have a few milliliters of it.

[Mega]

I got 3 samples , two of them with some 209 ng/uL  and 176.5 ng/uL  and one experimental (didn't change the protocol for 1.5mL overnight culture to 3mL) with 54ng/uL.

So just double the ammount of chemicals when you make twice the volume ;D And get much better yields. The miniprep was done with just 3000 rpm for all and therefore longer peroids of rotation, I have to get a better device...

[Nathan McCorkle]

The first two concentrations are decent, did you records the 230, 260, and 280nm absorbance levels? The ratios between them tell you about organic solvent contamination (230nm, mainly phenol but others too) and DNA to protein level which is a measure of general purity (280nm is tryptophan's peak, which is used as the average/common protein peak)

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[Andreas Sturm]

Yeah, there was a 260:2801 ratio...

I didn't write it down, but it is still saved in the computer .... Gonna get that.

Did a transformation with one of those plamid (176 ng/uL), gotta look if it works and how many colonies... (Inkuating them at room temp so still no visible growth...)


 

2012/7/17 Nathan McCorkle <nmz...@gmail.com>

[Mega]

So, the values are:

For the 209.39 ng/uL plasmid  the 260:280 ratio is 1.46
176.51 ng/uL   -> 1.91
54.17 ng/uL  -> 1.74



I think 1.91 is the best? Should it be high or low? ;)

[Avery louie]

Iirc as close to 2 as possible.

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[Nathan McCorkle]

230nm is used for phenol and related organic detection, it doesn't factor into the 260:280 number

On Jul 31, 2012 7:31 AM, "Slangtry" <sarah....@gmail.com> wrote:

The 260:280 ratio should  actually be as close to 1.8 as possible.  Lower is indicative of too much protein and higher can indicate RNA.  (Could always tell when the RNase was going bad when all the preps were 2.1 or higher.)  Anything (approximately) outside the 1.7 to 2.0 range and it becomes difficult to trust the 230nm reading used to calculate the concentration. 

On Saturday, July 28, 2012 7:09:44 AM UTC-7, Avery wrote:

Iirc as close to 2 as possible.

On Jul 28, 2012 5:38 AM, "Mega" <masters...@gmail.com> wrote:

So, the values are:

For the 209.39 ng/uL plasmid  the 260:280 ratio is 1.46
176.51 ng/uL   -> 1.91
54.17 ng/uL  -> 1.74



I think 1.91 is the best? Should it be high or low? ;)





Am Sonntag, 27. Mai 2012 17:54:23 UTC+2 schrieb Mega:

Hello @all,

I was wondering if I could do a dirty miniprep without EDTA.

What I have: weak centrifuge, SLS (sodium lauryl sulfate), NaOH, vinegar, ethanol (I have access to both wodka and pure ethanol), water.


Can you do a mp with this limited resources?
What I want: Some ~60 to 70 % plasmids, some remaining proteins, RNA, mabe some fragments of chromosomal DNA.


I think with this percentage you can still do a transformation with E.Coli? Because Proteins will either be used or digested, RNA my do it's job inside the cell, chromosomal DNA has no origin of replication.

Only the plasmids will be able to replicate and thus you will get some transformants. These you select with ampicillin anyway. (Clearly, you won't get so much transformants, but who cares? )

Will DNAses inside the solution possibly destroy the plasmids??

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[Jeswin]

On Sat, Jul 28, 2012 at 5:38 AM, Mega <masters...@gmail.com> wrote:
>
> I think 1.91 is the best? Should it be high or low? ;)
>

What are you going to do with these clones? I guess the effect of
inhibition depends on what you plan to do with it.