cDNA topics
-
cDNA topics
cDNAs
We've had considerable discusssion about this problem within the
DOE genome program. Procedures for making/rendering cDNA libraries
equimolar have been devised in labs of Sherman Weissman at Yale
(recent PNAS paper) and Agiris Efstratiadis at Yale. Computer
search for references.
However you may be better off working with a vactor providing
for directional cloning of the cDNA, but not normalizing. The
formalization process can tend to eliminate members of multigene
family which are individually interesting. Thus:
- Make your library in a vector supporting directional cloning.
- In the reverse transcription, use saturating quantities of
polydT as this will tend to shorten the length of cloned polydA
3'insert tails.
- Prepare blots of your library.
- Probe first with ribosomal DNA probe to avoid picking these
dominant clones.
- Initially sequence 3' ends as these will tend to be most divergence
within members of a single multigene family, that is, their 3'
untranslated region.
- When you find that you've re-sequenced a 3'end, use it as a
probe on your library blot to eliminate that specie of abundant
clones.
- AND so forth.
In this way you'll make your way through from more abundant to
less abundant clones, with only about a 2 X resequencing of 3'
ends.
A second library produced through random priming on mRNA will
probably be usefull It can be used as a target (its blot that
is) for probing with the 5' ends of probes derived from the first
library. Thus 5' segments not reverse transcribed in the first
library may be found extended.
Subject: Poly(A)+ selction
I am trying to do Poly(A)+ selection of RNA using either Oligo(dT)
cellulose or Amersham Messenger Affinity Paper. I have been
having a lot of trouble with the procedure, and I need some advice.
I have dry oligo(dT) cellulose, which I prepare for selection
by first hydrating it in binding buffer (Tris/EDTA/SDS/0.5 M
NaCl), and then washing with several bed volumes of 0.1 N NaOH,
which is recommended in several published protocols. I am not
entirely sure what this step is supposed to do, however. If
anyone can answer that, I'd be grateful.
After the NaOH, I wash the cellulose with water, and then with
Binding Buffer, restoring the pH to 7.5. The cellulose returns
to a white color. I then heat my RNA sample to 65 C for 10 minutes,
and chill on ice. I add 2 x Binding Buffer, and apply to the
column.
The column is washed, after 10 minutes of binding, with 0.5
M NaCl and 0.1 M NaCl, and then the RNA is eluted with TE buffer
heated to 65 C. The eluate is reheated and reapplied to the
column to squeeze more stuff out of it.
When I run the "Poly(A)+" RNA on a gel, however, there
is no attenuation of the rRNA bands, compared with an equivalent
mass of total RNA. Nor is there an enhancement of these bands
in the Poly(A) fraction. It is as if there is no specific binding
at all. My yields, if that is the correct term, are around 1%
to 3% of the total RNA applied.
I have tried doing this using a batch method, as well. I have
also tried using messenger affinity paper, with the same results.
The discussion in Current Protocols and Maniatis suggests that
the rRNA bands should be almost undetectable in a poly(A)+ fraction.
What kind of mistakes could I be making to prevent specific
binding?
Is the NaOH wash step really necessary?
Why do some procedures call for LiCl and some for NaCl?
Why do many procedures call for SDS or Sarkosyl?
I wish to make a cDNA library with this RNA. As long as I am
using a poly(dT) primer, do I really need to fractionate the
RNA, or could I simply use an amount of total RNA in the first
strand reaction that would contain an appropriate amount of poly(A)+?
Am I stupid? clumsy? unlucky? or all three?
mRNA complexes with rRNA and these aggregates may need to be broken
before oligo dT chromatography will do any good in enriching
for the polyA+ fraction. See
Banile et al. (1976) Analytical Biochem. 72:413427.
>In his method (deleted above) Daniel states that he heats
the RNA sample to 65
>degrees C. This is sufficient for breaking such complexes
and taking care of
>any other secondary structures. At least to the extent needed
to obtain the
>quality mRNA he requires.
>
The fact that he originally posted a message complaining about
rRNA co purifying with his mRNA indicates that he is NOT getting
the quality mRNA required. In the Banile et al. paper, the authors
show that, at least in the case of mouse brain RNA in their preparations,
heating at 65C for 2min. in the presence of 0.5M NaCl was not
sufficient for complete removal of rRNA.
They hypothesized that either this treatment was not sufficient
to break some mRNA:rRNA aggregates, or that aggregates could
reform on the column. In any case, after an initial oligodT run
to remove most of the rRNA, the polyA+ fraction was eluted and
a more rigorous denaturation was performed by heating to 55C
in the presence of 80% DMSO and 1M LiCl. Following heating, the
sample was diluted tenfold and rerun on the column. Elution profiles
indicated that of the initial polyA+ fraction from the first
step, 36% did not bind to the column, suggesting that roughly
a third of the RNA from the initial step was polyA. After elution
of the remaining 64% that bound to the column, this polyA+ fraction
was run a third time, in which case 99% bound as a polyA+ fraction.
Subject: cDNA library
I am trying to construct a cDNA library in lambda phage (lambdaZAP)
using Stratagene's cDNA cloning kit. I am having trouble with
low recombinant titers. Although the literature that comes with
the kit seems to suggest that I can get up to 10^7 pfu from a
single cDNA construction, I am only getting about 200,000 pfu.
This is distressing.
When I make my poly(A)+ RNA and run it on an agarose gel, I seem
to get an adequate size range (from 200 nt to over 8 kb), but
my first strand cDNA seems to downshift in size to give a maximum
of about 2 3 kb. What kind of parameters influence the ability
to synthesize flulllength cDNA? For instance, could I tweak
the system to get larger reverse transcripts by using less RNA
for a given amount of enzyme?
I also wonder about my vector ligation reaction. If I run a sample
of the ligation on an agarose gel, I get a large "band"
about 20 + kb, with a very, very faint smear going up to the
well. I don't think an agarose gel can conventionally give the
kind of resolution needed to distinguish from a poor ligation
reaction to a good one. What other kinds of assays can be used,
which do not use up expensive packaging extracts, to test the
quality of ligation?
When I read discussions in Maniatis or the RED BOOK, the advice
mostly centers around the importance of RNA quality. What can
I do to test RNA quality for cDNA library construction (besides
running a gel to check for size, or doing a Northern to test
for integrity)?
Subject: Poly(A)+ selection
I seem to have a few questions regarding poly(A)+ selections,
so I thought I would try to pull them together here.
So far, it seems that the "type"numbers for oligo(dT)
cellulose refer to the length of the (dT) tails. This may have
some bearing on the binding capacity of the cellulose, or it
may not.
In my hands, I cannot get good recoveries off of columns run
using SDS in the buffer, nor can I make batch methods work.
Since this is a simple binding/elution reaction, I would think
that the column geometry is not as critical to the outcome as
it is in gelfiltration columns in which molecules of different
size must be separated. If this is so, then would an Eppendorf
tube with a small hole in the bottom work as well as a small
column as a 1 ml syringe?
I have been trying to figure out how much total RNA can be loaded
onto a given amount of cellulose. I looked in Maniatis, which
says that a 1ml packed volume (0.2 to 0.5 grams, I think) should
be sufficient for 10 mg of total RNA. In Methods in Enzymology,
they calculate the capacity from the published capacity of 100
A(260) units per gram. With some assumptions factored in, a
figure of 10 mg of _messenger_RNA is given! Over two orders of
magnitude greater than the estimate given in Maniatis. In a
batchwise protocol given by Promega, the loading capacity of
oligo(dT) cellulose is given as 500 micrograms of total RNA for
0.3 g of cellulose. Stratagene, however, takes 1 gram of cellulose,
suspends it in 20 ml of DEPCwater, and uses 920 microliters in
a column made from a 1 ml syringe. This is loaded with up to
0.5 mg total RNA.
The capacities given by Promega and Stratagene seem to be the
closest match, more or less (0.3 g per 0.5 mg RNA vs 0.1 g per
0.5 mg RNA). These figures still make up a large ballpark.
Is experimentation the only way to figure how much RNA to load
on a given column?
Washing steps are sometimes done by gravity and somethimes by
centrifugation. Volumes of wash range from 1 x column volume
to 10 x column volumes. Is a small wash with centrifugation
as good as a larger wash by gravity? I'd rather do a small,
fast wash for the sake of time.
PCR
In reply to Doug Prasher's quest for efficient isolation methods
and automated systems, I can offer the following: In a recent
Nuc. Acids Research an article appeared in which plant material
is squashed onto a hybridisation membrane (in the presence of
NaOH etc.). The membrane can then be stored or used immediately
for use in PCR reactions. This method was designed for rapid screening
for horticultural / agricultural applications. I have not tried
it yet, but will do so in the near future. Doug Prasher's moths,
being a squishy sort of organism, might be suited to this type
of procedure. The reference is: Nuc. Acids Res. Vol19(24):6954.
Langridge et al. Squashes of plant tissue as substrate for PCR.
>I have read that it is exceedingly difficult to blunt-end
ligate products from
> PCR (without cleaving first in internal restriction sites).
I am attempting
>to clone the product of a PCR reaction into a blunt end cutter
site (eg SMA 1)
>in pBluescript. Does anyone have suggestions?
The following worked for me:
After the PCR reaction, I added 10 units of T4 DNA polymerase
to the sample (I didn't remove the oil, one has to careful to
ensure the sample + T4 pol is well mixed). I incubated it at
37 deg for 15 min, and then did a regular clean up and agarose
gel/glass milk excision of the fragment. I find that if I heat
inactivate the T4 pol, it gives poorer yields... maybe going up
to 70 deg sends the exo activity nuts after the pol activity has
quit... who knows.
There is also a methods paper (in Nucl Acids Res , I believe)
that recommends a proteinase K treatment of the PCR'd DNA. The
authors say that this treatment improves cloning ability of PCR
products... I haven't tried it.
By the way, when I cloned my 650 bp PCR product, the positive
clones were all light blue.... I was cursing because the transformation
plates had few white colonies.
I would like some help/advice with problems encoountered using
PCR cloning. I've been following the PCR threads hopefully none
of this is rehashing.
We reverse transcribe total RNA to obtain specific cDNA for immunoglobulin
V region genes. We then PCR amplify with a 5' leader primer
and the same 3' primer used in RT'n or one upstream. This is
followed by phenol extraction, double restriction digests (Sac
I and Xba I for example), then DNA gel electrophoresis and band
isolation by glassmilk. Cloning is into pGEM11 (also double
digested) and transformation into DH5(alpha), plated on IPTG/Xgal.
Problems:
1. Often the number of recombinants is *very* low. Expect 100200
cfu in plating and get maybe 220 cfu's.
2. White colonies almost always DO NOT have inserts of the expected
size (we are checking light blue colonies, blue centered colonies,
etc).
3. Some colonies do have the correct size but the sequence of
these plasmids is NOT a V region immunoglobulin gene. A southern
blot of the PCR amplified product with a V region probe shows
great hybridization.
Subject: Amplifying *large* DNA...??
Does anyone out there have a TRIED and true recipe for amplifying
large [>4kb] peices of DNA, using PCR, that they would care
to share?
I just failed to amplifiy a 4.9kb fragment using the "vector
PCR" protocol that was published in last months BioTechniques
[p446452]:
100uL rxn vol.
200uM dNTP
1.5mM MgCl2
50mM KCl
10mM Tris [pH 8.3]
0.01% gelatin
25 pMoles each primer
2.5 U AmpliTaq
110 ng template
2 min @ 94 C; 2 min @ 52 C; 5 min @ 74 C for 30 rounds.
I actually used 50 pMoles of a single primer that should prime
at either end of my insert and an unknown amount of template
[~ 200ng].
Subject: PCR of larger DNA Fragments: summary
I recently posted here, looking for help with my PCR optimizations.
I am trying to amplify a 5kb transposon out of maize. The protocol
I stared with was based on a "vector PCR" protocol
published in October's BioTechniques [p446452]:
> 100uL rxn vol.
> 200uM dNTP
> 1.5mM MgCl2
> 50mM KCl
> 10mM Tris [pH 8.3]
> 0.01% gelatin
> 25 pMoles each primer
> 2.5 U AmpliTaq
> 110 ng template
>
> 2 min @ 94 C; 2 min @ 52 C; 5 min @ 74 C for 30
rounds.
The following are some of the suggestions that I recieved:
JUST_W@rz.uniulm.dbp.de <Walter JUST> writes:
>Please refer to Nucl.Acids Res. _18/4_, p.1079 (1990). A colleague
at our >university was using gp32 for enhancing PCR of products
up to 6.5 kb. >gp32 is available from Pharmacia Biosystems.
I haven't tried it yet...it's on order (and I will let you know
if it helps).
JBISHOP@POMONA.CLAREMONT.EDU <J. Bishop> writes:
> .....most important is that you get
a working
>primary amplification of your template. I have found that
it often
>works to set your first cycle of denaturing, annealing, and
amplification
>for twice as long as the others, i.e. 4 minutes at 94, 4 minutes
at 52, and
>10 minutes at 72. Subsequent cycles can then be performed
at the original
>times. I might suggest that you lengthen time period allowed
for amplification
>Five minutes is not necessarily enough time to lengthen a
5 kb fragment. Third,
>you did not state how long your primers were, but you might
want to try using
>a lower annealing temperature. For 10mers I use 42 degrees.
>These are just basic places to start, but they have worked
for me in the past.
>
> oh, yeah, you might want to include a ten minute run off
at the end, though
>I don't suspect that will help if you aren't getting any amplification
at all.
I haven't tried this..and at the moment I don't intend to, the
times just seem WAY too long. The run off at the end sounds
good, particularly if I was intending to clone the product.
snella@biotek.ifas.ufl.edu <Liz Snella> writes:
>Do your primers work when you try to amplify smaller regions?
Yes! Good question though, I was really worried about that for
a while [o/n].
>If you have any old 32P hanging around, try filling in the
ends of the products
>with label. The 8 KB fragment that I mentioned is not visible
on Ethidium gels
>but is visible if you endlabel. Then run the labled fragments
on a agarose gel
>dry it and expose it. Even with month old label it only takes
an overnight
>exposure.
I blotted and did a southern instead...I had hot probe already.
[nothing!!]
>Try doing a "hot start", before you place the tubes
in the machine have it up
>to 94 already and pause for several minutes before you start
the first cycle.
Good idea: but you need to leave either the Taq or the template
out untill the reactions are at 94oC for it to be effective (I'm
sure this IS what Liz meant.)
The weird thing is that noone suggested mucking with the [MgCl2]
(too obvious?) I dropped it 3 orders of magnitude (1.5uM) and
started to see amplification.
NBR@AC.DAL.CA <Bruce Ramsey> (in an unrelated posting) wrote:
>...... I've on occasion obtained PCR products
>which consist of lambda sequences entirely due
>to priming from the 3' seven or ten bases of a 17mer
>and 20mer respectively. While the front end of these
>primers were dissimilar to any lambda sequence, the
>downstream ends were perfect matches.
>
>In other words, we should definitely check the 3'
>half in particular against the vector sequences.
I agree 500%! I have 3 or four smaller bands ( more intense than
the product I'm actually after) that ARE due to a 9bp and two
7bp stretches of homology at the 3' end of my primer! Very poor
design on my part.
I still have a ways to go before I get this particular amplification
optimized and I'm more than willing to share my experiences with
those who are interested (I'll post another summary if I get
more that a couple of requests).
Subject: Re: Thermostable polymerases without templateindept activity
In a previous article, bchtantw@NUSCC.NUS.SG (Dr Tan Tin Wee)
says:
>Could anyone recommend me thermostable DNA polymerases for
PCR that
>do not have DNAtemplateindependent polymerase activity?
>I understand that Taq polymerase puts in A's preferentially
at the
>3'ends of the PCR product (which is the basis of the TA cloning
technique).
Are there any DNA polymerases that do not have this activity?
Every DNA dependent DNA polymerase and reverse transcriptase
that I have tested has such an activity (although to varying
extents). These include:
AMV RT
MuLV RT
DNA Pol I
Klenow fragment of Pol I
T4 DNA Pol
T7 DNA Pol
Taq DNA Polymerase
Tth DNA Polymerase
Tfl DNA Polymerase
I'm not sure that I understand why this nontemplated addition
of a nucleotide occurs, but it is reasonably easy to reproducibly
observe. And the nucleotide does not have to be a dA. The standard
system that I used to look at this activity was to incubate a
EcoRV digest of a plasmid (pRCASBneo) with alpha labelled dTTP
(3.3 uCi/ul, 20 uM final conc.) and the enzyme (in the recommended
buffer) for 30 minutes at the recommended incubation temperature.
You can clearly detect labelled fragments on an 30 minute autorad
of a 1% agarose gel.
Regards,
Ashok
P.S I have not tried NEBs thermostable enzyme or the one sold
by Stratagene but I suspect that they will have this activity
too.
Subject: Re: PCR:Degenerate Primers
]In article <1991Nov30.124052.328@ucbeh.san.uc.edu>, penas@ucbeh.san.uc.edu
writes:
>Hellow!
>Another question on PCR. I am trying to amplify fragments
of about 160 to 200
>bp using degenerate primers targetted specifically to the
DNA binding domain of
>a group of transcription factors. The two primers that I
am using are
>degenerate (not highly because the target region is very conserved)
and are a
>20mer and a 22mer with 30 and 50% GC content, respectively.
I have been
>running my PCRs both at 45 and at 55 C. So far I get amplification
but it
>seems that it is not being specific because I don't get bands
but smears. I
>really don't know if this is good enough ( I intend to clone
the amplified
>fragments). I would appreciate comments and advise in terms
of how to improve
>my PCRs. I have been working with 2mM Mg++ reactions. Thanks
in advance for
>your comments. I will summarize your recommendations and comments.
Aaaahhh!! Degenerate PCRs a subject that has caused me great
pain for 3 years! The following experiences are based on using
degenerate primers (a 17mer and a 22mer each with 192 (!) degeneracies)
used to amplify a 700 bp segment from an archeobacterial strain
using a COY Tempcycler.
In my experience, I have found that with degenerate primers, they
really HAVE to be clean... that reduces the smear somewhat.
After playing around lots with anneal temperatures and time,
I found that using as high an anneal temp as possible with a
longish anneal time worked well. (I know it's not a precise condition,
but each case will be different). I ended up cycling at 94/1.5
min; 55/2.5 min; 72/2 min X 30 cycles. I also used 100 pmol
of each primer in the 50 ul reaction and .5 ug of genomic DNA,
and used a Mg conc of 2.5 mM. The primer concs may seem high
but the "correct" primers were in 1/192th of that
I suppose you need a balance so that the correct primers are
available to prime, yet you don't want too much of the "junk"
primers to prime spuriously.
Lowering the anneal temp to 45 deg gave more bands, lowering the
anneal time gave less intense ones which kinda made sense to
me the right primer out of 192 at each end isn't in that high
a conc, and you could be encouraging spurious priming at lower
temps.
You probably still may get multiple bands. I would definitely
find some way of sequencing the product to see if you've got
the right one I chased a red herring by assuming the most intense
band was my product which it wasn't. I eventually cloned some
bands to check that the correct product's ends matched the amino
acid sequence from which the primers came.
Subject: re:PCR sequencing
We are doing alot of PCR sequencing. In my own experience, sequencing
symmetric PCR fragments can be difficult. The things that have
really made a difference are 1) lowering PCR primer amount to
5 pm each (or removing PCR primers with centicon 100) and 2)
using Taq polymerase for sequencing and to cycle the sequencing
reaction using a labeled sequencing primer( in our case a fluorescent
primer). By cycling the sequencing reaction, you will get a
linear amplification of extension products which should give
plenty of sequence data. Also, with this approach you can directly
sequence symmetric PCR fragments without any purification! If
you want more info just let me know....Good-luck.
Sincerely, Sandy Koepf
Applied Biosystems Inc.
From: mats@bio.embnet.se (Mats Sundvall)
Subject: Re: inverse PCR
> Does anybody have any particular tips for doing inverse-PCR?
> We've tried alterring magnesium conc., dilute ligations to
encourage
> intramolecular ligation, long extension times - but no product
(dispite
> direct PCR working on template)
> Any ideas welcome.
A couple of new methods that is useful if you only know one end
has been published. It all depends on your specific application
if they are useful to you. Our experiance with inverse PCR are
not the best. The methods we have used depend on ligation of a
linker to the ends and amplifying only the ones with one internal
primer. Variations one this is scheme is published as bubbel-PCR
by Riley et al NAR Vol 18, No 10, page 2887. Another very similar
method was published in a more recent NAR but I do not have this
reference.
From: Chris_Jones@vme.ccc.nottingham.ac.uk
Subject: PCR cloning
I've just seen an ad for Invitrogens PCR cloning kit where they
claim that PCR products always have a 3' A added on. Does anyone
know anything about this (reference?). I have always done a klenow
step before cloning but didn't realise that the ends were ragged
because extra bases were added on? Quite a few of my applications
would be wrecked by an extra A so any tips about eliminating them
(if they exist) would be great.
Subject: Re: Oligonucleotide probing
It might be that asymmetric PCR is an answer:
i.e.
Primer A = 50 - 100 pmol
Primer B = 0.5 - 2 pmol
Subject: re:PCR CLONING
A reference that may be of interest to you is Perkin Elmer's newsletter
(Amplifications, May 1990, Issue 4). There is an article called
"DNA generated by PCR (...) has non-template nucleotide additions:
Implications for cloning PCR products. They suggest using Klenow
to remove the extra nucleotide (which probably is an A). Hope
this is of some help to you.
Subject: RE: PCR sequencing
We at @FINNPHI.BITNET do a lot of PCR-sequencing with reasonable
success.
Here's our protocol in which you may find the answears you need:
1. denature 0.6-1.8 pmol of pcr product (purified) by heating
2 min at 100 C in water.
2. snap cool on ice for 5 min.
3. add 6 pmol primer.
4. add buffer.
5. anneal primer for 10 min at +37 C
6. add enzyme, label and do the labelling for 5 min at +37 C
7. divide to extension/termination reactions and keep at +37 C
8. stop reaction (loading buffer)
There are a few more tricks:
- if your DNA is dirty (as agarose purified) use double amount
of enzyme, this countereffects the agarose inhibitory effect on
enzyme activity.
- if you want to read near the primer add manganese buffer from
the newest Sequenase kit (one-tenth of total reaction volume)
(Tabor et Richardson:PNAS 1989;86, 4076-4080).
We do our sequencing using the Amersham Multiwell system in a
30 mikrol reactionvolume, but i know about people using Sequenase
with similar success. Those of you using Klenow outhere....
Forget about ancient inferior systems, T7 rules OK!
Subject: PCR Sequencing
If you are directly sequencing PCR products, I need your advice.
We are trying to sequence PCR products of 400bp and 750bp in length
and having almost no success. We have tried changing a number
of variables none of which improve the situation. Our M13 control
reactions done in parallel ALWAYS work.
OUR STANDARD CONDITIONS are: 0.6 pmol ds template, 6 pmol primer
(also used in PCR), boiled 10 minutes, snap frozen in liquid nitrogen.
Thawed on ice and then sequenced using Sequenase.
THE GENERAL RESULT is: No sequence ladder (or extremely faint)
but there is a fairly intense band in all four lanes at a position
in the sequencing gel approximating the full-length fragment.
On some gels, a short sequence ladder is observed just below
this intense band, which we intrepret as termination at the end
of the fragment.
We have assumed the problem results from a low concentration of
the template-primer complex in the labelling reaction. We believe
there is enough nucleotide present in the labelling reaction
such that the Sequenase makes the complexes completely double-stranded
before the dideoxys are added in the termination reaction.
We have tried the following variables, individually of course,
with no improvement:
1) 4 pmol ss template (via assymetric PCR) instead of 0.6 pmol
ds template.
2) Increasing sequencing primer to 40 pmol.
3) Changing the thawing conditions:
i) leave on ice 90 min
ii) place in a -20C block, let warm to 7C or 15C.
4) Dilute labelling mix: 1:5 is normal, 1:20, 1:75, 1:300.
5) Shorten labelling reaction time at RT and at 0 C.
Subject: PCR cloning
I thought I might say a word or two in response to recent inquiries
about cloning PCR products.
As far as the concept of Taq DNA pol leaving an overhanging A
residue at the 3' end of PCR products, I'd say that the A (or
some other base) is not there in some percentage of the DNA.
You certainly can clone PCR products in a blunt end site with
very good efficiency with no other post- (or pre-) PCR treatment
than kinasing.
If we add T4 DNA pol to the PCR mixture immediately following
PCR, and incubate at 37 degrees for 15 minutes, cloning efficiency
is doubled. Thus, your overhanging base. A 1-hour treatment at
15 degrees with a few units of Klenow seems to work fine also.
You guys not wanting to spend the bucks on those TA cloning kits
might want to consider this method.
Subject: Re: PCR frag. cloning into M13 vectors
RP>Can anyone help me with m13 cloning I have some problems
RP>with ligation a PCR product into m13 vector (m13mp18)
RP>The size of fragments is varying from about 100bp to
RP>300bp size in lenght. I have opened the m13 vector
RP>by pstI /HindIII double digestion and also I have
RP?di>tested these PCR products same way because we have
RP>planned primers so that PCR is (probably) producing
RP>pstI and HindIII sites to the ends or near of the ends
RP>of these fragments.... If anyone has tried to introduce
RP>DNA fragments into m13 vector please let me know of
RP>the problems and how did you solve those problems...
RP>Especially talking about PCR fragment cloning into
RP>m13 and also into expression vectors too...
I haven't tried M13 but have tried the following protocol using
BRL's phagemid (pUC derivatives). I cut the vector with HincII,
and do a standard pcr reaction to get insert DNA. After the pcr
reaction, separate the reaction from the mineral oil, add approx.
10 units of T4 DNA Pol, incubate it at 37deg for 30 min, and gel
purify the fragment.
I ligate this DNA (1 or 2 ul) with the vector (25 ng), electroporate
into DH5alpha cells and plate for colonies on ampicillin/xgal
media. I usually get 100 to 200 colonies per plate with about
5% to 15% white colonies. I ignore this, because in the case
of two 600 bp pcr products, the correct clone (identified by subsequent
sequencing) was light blue on xgal plates! I do colony lifts using
the gel-purified pcr product as a probe, and usually about 5 to
15 colonies (per plate) give signals.
It has worked for me at least a half-dozen times. Another thing
of interest... yesterday, I ran across an NAR paper that was titled
"Improved cloning efficiency of polymerase chain reaction
(PCR) products after proteinase K digestion." by Crowe et
al., 1991. Nucl. Acids. Res. vol:19 No:1 page 184. The authors
claim that Taq pol may stick to the ends of pcr'd fragments thus
reducing "clonability". They claim that proK treating
the fragments increases cloning efficiency umpteen fold... sounds
like I may need to try this next time I have to clone a pcr product!
Subject: Re: PCR fragment cloning into m13 vectors
> .... If anyone has tried to introduce (PCR)
> DNA fragments into m13 vector please let me know of
> the problems and how did you solve those problems...
> Especially talking about PCR fragment cloning into
> m13 and also into expression vectors too.
I clone PCR fragments into M13 vectors all the time, by pretty
much the same method you are trying to use. I design my primers
so that a restriction site is included near each end. I can think
of some problems that may be causing your cloning difficulties:
1. The primers should be designed so that the restriction
cleavage site is near the center of @20mer. If you are hoping
that the enzyme will cleave off the very last nucleotide or two
of the PCR product, you may have trouble. A method was
described in Nucleic Acids Research 18:6156 (1990) for "Efficient
cloning of PCR generated DNA containing terminal restriction endonuclease
recognition sites" In this article, Jung et al. showed that
if the PCR product is generated with 5'-Phosphorylated primers
and then concatemerized with T4 DNA ligase, it is more readily
digested at the terminal restriction sites. I have not had a
reason to try this, but it sounds reasonable.
2. The M13 DNA may not be completely digested with both PstI
and HindIII.
There was a BRL Focus article about double digestions of the multiple
cloning site (Focus 8:3 p. 9, 1986) in which they investigated
the importance of using the restriction enzymes in particular
order. They found that HindIII should be used before PstI and
not the reverse. Also, it would be difficult to use them simultaneously
because their sites are so close together.
3. You can verify that your M13 DNA is cut by both restriction
enzymes by a simple test described in FMC Resolutions newsletter
vol.2 no. 3. It basically involves taking aliquots of your DNA
after both the first and second restriction digestion and digesting
them together with an enzyme which cuts outside of the polylinker
- like MstI. This produces two pieces of DNA which differ in
length by the distance between the first two sites (like PstI
and HindIII). Separation on 4% NuSieve agarose makes it possible
to distinguish the two bands, which may be as few as 10 bp apart.
I have used this protocol many times and would be glad to fax
you a copy, if you need it.
If none of these ideas solves your problem, you could directly
sequence your PCR product, but that involves a whole new set
of problems!!!
Arabidopsis chloroplast isolation procedure.
Ludwig, R.A. (1991) "Plants shuttle CO2 equivalents as carbamoyl
phosphate from mitochondria to chloroplasts" in Molecular
Approaches to Compartmentation and Metabolic Regulation, (A.C.
Huang, ed.), Am. Soc. Plant Physiology, 1991.
Intact Arabidopsis chloroplasts can be highly purified from illuminated
leaves as follows:
1. Ultrasonicate 20 g. leaves in bath sonicator with sterile,
distilled water. Spin in salad dryer.
2. Mix leaves with 50 ml ice cold grinding buffer (10 mM sodium
pyrophosphate, 5mM MgCl2, 2 mM sodium ascorbate, and 0.33 M sorbitol,
pH 6.5), and whirl 10 sec in Cuisinart. Note: grinding buffer
should be "partially frozen" in dry ice - ethanol bath
and then vigorously shaken to suspend pyrophosphate.
3. Homogenize 10 ml aliquots 3-5 seconds with Polytron. Note:
This is the most critical step in the isolation procedure. While
only a small fraction of cells are lysed, virtually all chloroplasts
remain intact.
4. Filter homogenate through 2 layers of cheesecloth and Miracloth.
Quickly centrifuge filtrate 1 min at 8,000 ~ g in a swinging bucket
rotor at 2xC. Wash pellet with grinding buffer.
5. Resuspend pellet in 0.25 ml resuspension buffer (50 mM HEPES
pH 7.6, 2 mM EDTA, 1 mM MgCl2, 1 mM MnCl2, and 0.33 M sorbitol).
6. Layer resuspension on 17 ml 50% (w/v) Percoll gradient in gradient
buffer (50 mM sodium MOPS pH 7.8, 2 mM EDTA, 0.15% BSA, and 0.33
M sorbitol); use 30 ml Corex tube. Spin 10 min at 5,000 ~ g in
a swinging bucket rotor at 2xC. Note: Preform the gradient by
spinning the Percoll solution in gradient buffer 100 min at 10,000
~ g in a swinging bucket rotor at 2xC.
7. Remove the lower (intact) chloroplast band with a plastic,
disposable pasteur pipette. Pellet chloroplasts in swinging bucket
rotor. Resuspend intact chloroplasts in 0.5 ml resuspension buffer,
examine them by light microscopy, and freeze in 0.05 ml aliquots
at ~70xC.
PURIFICATION OF DNA BY BINDING TO GLASS POWDER
Binding and Wash Solutions
NaI solution: 90.8 g NaI
1.5 g Na2SO4
in 100 ml H2O. Filter through Whatman No.1. Put dialysis bag
containing 0.5 g Na2SO4 in bottle to keep solution saturated.
Store foil-wrapped at 4 oC.
NEET Wash: 100 mM NaCl
1 mM EDTA
50 % EtOH
10 mM Tris pH 7.5
Store at -20 oC.
DNA Purification:
To purify DNA from agarose gel, weigh gel slice.
Add 2 - 3 ml NaI solution per gram of gel.
Incubate at 37-50 oC, mixing frequently until agarose is totally
dissolved.
Add 1 5l of glass slurry per 5g of DNA.
Incubate on ice 5-10 mins, mixing occasionally.
Spin 5-10 secs in microfuge, remove and discard supernatant.
Wash glass pellet with 250 5l NaI (or 10 x volume of glass if
larger).
Spin and wash pellet 2-3 times with EtOH wash (same volume).
Dry pellet well, removing all residual liquid (air dry or use
Kimwipe carefully).
Resuspend pellet in H2O or TE (> 10 5l) and elute DNA at
50 oC for 5-10 mins.
Spin 1 min in microfuge and remove eluted DNA in supernatant.
DNA is ready for ligation, restriction, radiolabelling etc.
DNA bind to glass at high salt and low temp, elutes at low salt
and high temp.!
(To purify DNA from solution, add 3 volumes of NaI solution, immediately
add glass and put on ice).
PREPARATION OF GLASS POWDER
Use silica 325 mesh (a powdered flint glass available from ceramic
shops)
Resuspend 400 g of glass powder in 800 ml ddH2O in a 2 litre
flask.
Stir for 60 mins.
Allow to settle for 90 mins.
Take the SUPERNATANT (which contains the "fines" of
interest) and pellet in Sorvall (GSA rotor, 10 mins at 6000 rpm).
Resuspend pellet in 200-300 ml ddH2O.
Add nitric acid to 50 %.
Bring close to boil in fume hood.
Allow to cool.
Pellet glass as before, wash pellet 4-6 times with JddH2O (check
pH returns to neutral).
Store final pellet as 50 % slurry in ddH2O.
Store at -80 oC, working aliquot at 4 oC.
Cost for 2.5 kg of glass powder is approx $3
Subject: Re: PrepaGene
I purchased 1015 lbs of glass powder for ~$20 several years ago.
Needless to say we don't bother reusing glass milk.
My source was:
Cutter Ceramics
11908 Old Baltimore Pike
Beltsville, MA.
20705
Ask for 325 mesh powdered flint glass fines.
Sigma Chemicals sells powdered silica (also known as flint) for
about $15 for
500 g. This is the stuff we use for this... Don't get the _fumed_
silica
that's too small.
You have to do a size (1g sedimentation) cut taking the powder
that
settles out between 2 mins and 60 mins, then boil it all in 10
volumes of
50% nitric acid for an hour, and wash extensively.
You'll make enough for yourself and all your friends for a millenium
100 UL HMGR ACTIVITY ASSAY
In a 1.5 ml microfuge tube add:
20 ul of BSA/DTT solutionA
10 ul of NADPH solutionB
20 ul of 14C-HMG-CoA solutionC
50 ul of enzyme preparationD
Start the HMGR reaction by adding the enzyme and incubate for
20 minutes in a 30 C water bath.
Phenolics Extraction from Plant Tissue
Extraction of Free Phenolics
1. Grind 1 g of fresh tissue in 4 ml water with a Polytron
at room temp. for 30-60 s.
2. Add 4 ml acetonitrile, sonicate at 100 W for 30 s.
3. Spin in a table top centrifuge at full speed for 10
min, collect supernatant.
4. re-extract the residue with 4 ml acetonitrile for 4
h at room temperature, spin, collect and combine supernatant.
5. The residue is extracted with 4 ml acetonitrile again
overnight at 4oC. If necessary, extract the residue one more
time until the debris is completely decolorated. Save residue
for wall bound phenolics extraction.
6. Dry supernatant with nitrogen at 40-50oC up to 4 ml
(H2O), adjust its pH to 7 with 0.1 N NaOH (pH paper).
7. Extract the aqueous with 2 ml hexane in a capped tube
with shaking for 1 min, sit for 5 min. Remove organic phase (upper).
Repeat this step.
8. Residual hexane in the aqueous phase is removed with
nitrogen at 40-50oC, adjust its pH to 1.5-2 with 1 N HCL.
9. Load onto a C18-Sap-Pak cartridge (waters) pre-washed
with 4 ml acetonitrile and equilibrated with 4 ml 0.01 N HCl,
0.1% B-mecaptoethanol (BME), wash with 3.5 ml 0.01 N HCl, 0.1
0.1% BME, elute with 3.5 ml acetonitrile:H2O 4:6 v/v) 0.1%BME.
Skip first two drops, collect rest from the column.
10. Reduce vol. to 2 ml with nitrogen. Before HPLC or
spectrophotometer analysis, the sample should be centrifuged or
filtered to remove residual debris.
Extraction of Wall Bound Phenolics
1. The residue (wall material) from step 5 above is hydrolyzed
with 4 ml 2 N HCL at 90oC for 1 h, spin, remove supernatant to
a fresh tube. Save debris for lignin analysis.
2. Add 2 ml hexane to supernant, votex, centrifuge to separate
two phases, remove organic phase (upper) to a fresh tube, extract
aqueous phase with 2 ml hexane again, combine organic phase.
3. Dry hexane with nitrogen at 45oC, suspend pellets with
1 ml 0.01 N HCl. Before HPLC analysis, centrifuge or filter the
sample to remove residual debris.
HPLC Analysis of Phenolics
* See attached HPLC method and reference 1
Ligin Analysis -- Thioglycolic Method
1. Wall material after phenol extraction is incubated
in 5 ml 2 N HCl plus 0.5 ml thioglycolic acid at 90oC for 1 h.
2.
ISOLATION OF AN ENDOGENOUS ELICITOR
Media from elicited cells was fractionated using Sep Paks and
an increasing percentage of Acetonitrile, namely: 20, 40, 60,
80, and 100 percent. These fractions were concentrated and used
to elicit cells. PAL assays reveiled an active fraction, the
20 - 40 % cut.
This fraction was run out on a TLC plate and the 18 sections
(each 1 cm wide) were excised, eluted, concentrated, and used
to elicit cells. These PAL assays showed activity, but the sectioning
of the plate was not appropriate to isolate the active fraction.
Therefore, the TLC procedure was repeated and 10 spots were excised
individually. The 7th spot had the highest activity.
The same procedure of isolation was automated using the HPLC.
Fractions came off every minute during a 22 minute gradient increasing
from 20 to 40 percent. These fractions were concentrated and
are currently in the -70 freezer waiting for good cell lines for
elicitation.
Cells that were elicited by the 10 TLC spots were prepared for
the HPLC using the Isoflavone procedure. We have the Chromatographs
of each. We did the same with cells elicited by: old and new
C.lind., Pmm, Yeast, and a water control.
We also ether extracted media that had been treated with elicitor.
We extracted at pH 6 and pH 2 and have the HPLC results from
both samples.
PREPARING POTATO TUBER DISKS FOR USE IN AN EXPERIMENT
1. Set the tubers out at room temperature in a dark place the
day before using them. Wash the tubers with mild soap and water.
Use only certified seed tubers that have been stored for at least
one month at 4 C.
2. Prepare sterile water by autoclaving several flasks containing
glass distilled water. The flasks must be cooled to room temperature
before the water can be used!
3. Also sterilize: filter paper circles to place in petri plates
(12.5 cm circles for 15 x 1.5 cm plates,
or
9 cm circles for 10 x 1.5 cm plates)
small vials for sonication of AA (if needed)
forceps
hockey sticks
4. The next day surface sterilize the tubers by immersing them
in 70 % ethanol for 3 min. Set the tubers on paper towels to
air dry.
5. Cut disks from the tubers using a flame-sterilized knife and
#15 cork borer. Do this on a plastic cutting board that has been
wiped down with 70 % ethanol. Put the disks immediately into
sterile water (this prevents browning).
6. Rinse the disks three times with sterile water and place them
into petri plates containing filter paper. Add enough sterile
water to saturate the paper; don't get water on the tuber disks.
Store the disks in a cool, dark area until treatment.
TREATMENT OF TUBER DISKS WITH ARACHIDONIC ACID
1. Transfer an aliquot of arachidonic acid (AA) stock to a sterile
vial or test tube.
2. Drive off the solvent (hexane/isopropanol, 3:2) under a stream
of nitrogen gas. A clear oily residue will remain.
3. Add sterile water to make the desired concentration of AA
and sonicate the vial with a probe sonicator until a milky white
emulsion is created (usually ca. 5 seconds). Avoid foaming and
introducting air into the emulsion; don't oversonicate.
4. Typically, AA is applied to tuber disks (#15 cork borer size)
in 100 ul aliquots at a concentration of 0.5 mg/ml. The emulsion
is spread uniformly across the disk surface with a sterile "hockey
stick".
5. The treated tuber disks are incubated at 20 C in darkness.
FLUOROGENIC ASSAY FOR BETA-GLUCURONIDASE ACTIVITY
GUS Extraction Buffer (300 ml)
50 mM NaPO4, pH 7.0 150 ml of 0.1 M stock buffer
10 mM Na2EDTA 12 ml of 250 mM stock
0.1 % sodium lauryl sarcosine 3 ml of 10 % stock
0.1 % Trition X-100 3 ml of 10 % stock
add 132 ml distilled water
10 mM beta-mercaptoethanol 7 ul/ml of extraction buffer
(add beta-MCE fresh before
use of the buffer)
Extraction of GUS activity
Grind a leaf disk in a 1.5 ml microfuge vial with a Kontes pellet
pestle. Typically, a tobacco leaf disk cut with a #9 or #15 cork
borer is homogenized in 700 ul of extraction buffer.
The extracts are spun briefly to sediment the particulate matter
and the supernatant is assayed.
Assay of GUS activity
Reaction buffer is 1 mM 4-methyl umbelliferyl beta-D-glucuronide
(Sigma M-9130) in extraction buffer (22 mg MUG in 50 ml). This
is good for at least one month in the refrigerator.
End point assay.
To a microfuge tube containing 100 ul of reaction buffer add
20 ul of the extract supernatant and mix well. Incubate the reaction
at 37 C for 60 min, then stop the reaction by adding 900 ul of
0.2 M Na2CO3.
Time course assay.
To a microfuge tube containing 500 ul of reaction buffer add
50 ul of the extract supernatant and mix well. Incubate the reaction
at 37 C. At 5, 15, 25 and 35 minutes after the start of the reaction
remove 100 ul aliquots of the reaction mixture and transfer to
tubes containing 900 ul of stop buffer (see above).
Determine the MU concentrations with the spectrophotofluorimeter,
excitation at 365 nm, emission at 455 nm.
EFFECT OF UV LIGHT ON GUS EXPRESSION IN TRANSGENIC TOBACCO
PLANTS
Methods
1. One cm disks are cut from leaves with a no. 9 cork borer and
placed abaxial (lower) side up on moistened filter paper in petri
dishes.
2. With the cover off the petri dish the leaf disks are exposed
to short wave UV light in a Chromato-View cabinet, model CC-60.
The optimal exposure peroid was determined with a simple dose
response experiment. For this setup irradiation for 1 to 2 min
gave the best induction.
3. The cover is replaced on the petri dish after treatment and
the leaf disks are incubated at room temperature in darkness.
4. GUS activity is determined with the fluorgenic assay as described
by R. A. Jefferson, 1987 (Plant Mol. Biol. Rep. 5:387-405). Each
leaf disk is homogenized in a 1.5 ml microfuge vial containing
700 ul of GUS extraction buffer. After a brief spin 20 ul of
the supernatant is added to 100 ul of the GUS assay buffer and
incubated at 37 C for 1 hour. The reaction is stopped by adding
900 ul of 0.2 M sodium carbonate.
IEF PAGE
* Gel preparation (12 ml)
2 ml 30% acrylamide (29:1)
2.4 ml 50% glycerol
0.6 ml Ampholyte (or 0.5 ml desired ampholyte plus 0.1 ml ampholyte
pH3-10, check instructions on specific ampholyte).
7 ml H2O
Mix well, degas for 5 ml under house vacuum.
25 ul 10% APS
20 ul TEMED
Mix carefully, do not introduce air boubles, pour a gel.
* Sample preparation
2X sample buffer: 60% glycerol, 5% ampholyte (pH3-10 is ok)
Mix equal vol. of sample and sample buffer.
* Electrophoresis
Assemble gel apparatus, add tank buffer
upper tank buffer: 25 mM NaOH
Lower tank buffer: 25 mM acetic acid
Load sample the same way you do SDS gel.
Electrophoresis at 200 V for 1.5-2 h and 400 V for further 2
h.
If you desire to determine pI, electrophoresis at 200 for 2 h
and then at 400 V for 3.5-4 h.
If Staining is desired, fix the gel with 10% TCA for 10 min with
gentle shaking, and then soak the gel in 1% TCA for at least 2
hr (prefer overnight) to remove ampholite.
Stain the gel with Coomassie Blue the same way you stain a SDS
gel.
To run second dimension gel, soak the gel slice (removed from
the IEF gel) in 2X SDS sample buffer for at least 30 min at room
temp. carefully place it into a slot of prepared SDS gel. Run
the gel the same way you do a regular SDS gel.
The reaction is stopped by adding 10 ul of 6 N HCl. Mix well
and let the reaction mix stand at room temperature for approximately
30 minutes to alow the mevalonic acid to lactonize.
Spin the reaction vials at full speed in a microfuge for 3 minutes
to pellet the protein. Apply 15 ul of the supernatant to a channeled
thin layer silica gel plate and when the spot is dry repeat with
another 15 ul. Use of a blow dryer will greatly speed the process.
Chromatograph with chloroform:acetone (2:1) in a TLC tank. A 20
cm plate takes about 1 hour.
The TLC system separates the labelled product of the HMGR reaction
(mevalonolactone) from the 14C-HMG-CoA and 14C-HMG which remain
at the origin. The migration of the mevalonolactone is determined
by adding 500 ug of unlabelled mevalonolactone and 30 ul of reaction
mix to the TLC plate before separation. The mevalonolactone is
detected after chromatography by iodine vapor. Alternatively,
labelled mevalonolactone can be used with reaction mix in an unneeded
lane, and detected by scrapping off 1 inch sections of the lane
and counting radioactivity.
Subject: Lambda DNA Prep's
In response to Charlie Hussey's request for information on preparing
lambda DNA from Dr. Meyerowitz's RFLP clones, we've had good success
in obtaining nice yields of digestable DNA with the procedure
described below:
Lambda DNA Preparation (adapted from D. Chisolm, 1989 BioTechn
7:21-23)
1. Grow suitable host (e.g. LE392) on plate. Inoc colony to NZCYM,
make overnight culture. Transfer 1/1000 to 100 mls NZCYM, and
grow to OD600=0.2. Pellet cells, resus 20mM MgSO4 to OD600=1.0.
Cells usable for two weeks @ 5 degrees C.
2. Combine 0.5 mls cells with lambda stock. Use 5x10^4 to 10^5
pfu of concentrated phage, or if from plates, elute plugged plaque
into 1 ml SM for at least 2 hrs and use 10-100 ul of that as inoculum.
Incubate @38 degrees C for 30'. Then combine this with 37
mls NZCYM in a 250 ml Erlenmyer or magenta jar. Shake overnight
@ 38 degrees C. Complete lysis should be evident, with clots
of cellular debris.
3. Transfer culture to oak ridge tube containing 100 ul CHCl3
and mix well. Add 370 ul nuclease soln and incubate @ 37 degrees
C for 30'. Add 2.1g NaCl and dissolve. Spin 20U @ 7K slow decel',
angle head rotor.
4. Transfer super' to fresh oak ridge tube containing 3.7 g PEG
(6-8K) and dissolve gently. Set on ice for 60'. Preheat 65 degrees
C bath.
5. Pellet phage @7K 20' (5 degrees C). Gently resus' pellet
in 0.5 ml SM containing 50 ug/ml RNAase A. Transfer to 1.5 ml
eppy'. Incubate at 37 degrees C for 10'. Add 0.5 ml CHCl3, and
mix gently but thoroughly. Spin 5' in microfuge.
6. Transfer super' to fresh tube containing :
20ul 0.5M EDTA
5ul 20% SDS
10ul proteinase K (2.5 mg/ml)
Incubate 65 degrees C for 30'.
7. Extract with phenol, then CIA. Reextract with CIA. Transfer
final super' to tube containg 170 ul 6M AmOAc. Add 0.7 ml isopropanol.
Cool 10' (-70 degrees C) to 30' (-20 degrees C), and pellet 8'.
Wash with 70% ETOH, vacuum dry, resus' in 200 ul TE.
8. Yields are sufficient that a labelling rxn using 4ul of phage
DNA makes good probes.
NZCYM per liter:
10 g NZ amine (casein hydrolyzate, type A)
5 g NaCl
5 g yeast extract
1 g casamino acids
2 g MgSO4 -7H2O
pH = 7.5
SM per liter:
5.8 g NaCl
2.0 g MgSO4
50 mls 1 M Tris (Ph 7.5)
2% gelatin
Nuclease Solution per 10 mls
5.0 mls glycerol
0.1 mls 3M NaOAc
H2O to 10.0 mls
50 mg DNAse I
50 mg RNAse A
Problems are usually due to too high or too low inoculation levels.
If the cultures are turbid after ON growth, then increase the
phage (cells reached stationary phase before phage caught up
to them); if the cultures are cleared, but not especially viscous,
and with little cellular debris, then lower the phage inoculation
(lysis prior to good cell growth). Hope this works.
Terry Delaney (DELANEY@SALK)
(postdoc with Dr. J. Chory)
Phage DNA isolation
Prepare DEAE:
Equilibrate DEAE-cellulose (DE52, Whatman) resin in 10mM Tris-HCl
(pH 7), 1mM EDTA, 0.01% Na azide. Keep resin as a 40% slurry.
Before use, wash slurry 2 times in L-broth and resuspend as a
80% L-broth slurry containing 0.01% Na azide. This can be stored
at 4 C for 1 week.
CTAB Method:
Prepare plate or liquid lysate from desired phage. The following
applies to 1 ml of lysate. It can be scaled up proportionally
for larger volumes.
To cell lysate, add DNaseI (Sigma) to a final concentration of
20 ug/ml and incubate at room temperature for 20 min. During last
5 min spin at 8000g for 5 min. Transfer supernatant to a new tube
and add gelatin at 50 ug/ml final concentration (gelatin stock:
0.3% with 0.01% Na Azide). Add equal volume DE52 cellulose (80%
slurry) and mix on a rotating wheel for 10 min. Centrifuge down
the pellet, collect supernatant, and add reagents to make supernatant
50 ug/ml proteinase K and 20mM EDTA. Incubate at 45 C for 15 min.
Then add CTAB (Sigma) solution to a 0.1% concentration (CTAB stock:
5% w/v in 0.5M NaCl). Heat tube at 68 C for 3 min, then cool on
ice for 5 min. Spin lysate at 8000g for 10 min at RT. Discard
supernatant and resuspend pellet in 1/5 the original volume of
lysate (or 1/10 if a large scale prep) with 1.2 M NaCl. Add 2.5
vol ethanol to the Na-DNA precipitate. Centrifuge at 8000g for
10 min, wash pellet in 70% ethanol, and re-centrifuge. Dry pellet
and redissolve in water or TE.
For more information: see Manfioletti & Schneider, NAR 16:2873-2884
(1988)
mannopine synthesis
Hi. Here is the info on mannopine synthesis: Use the methods of
Petit et al (1983) Mol. Gen. Genet. 190:204-214, with the following
modifications.
1. Mannopinic acid is more stable and works well for selection.
2. Do the indicated synthesis but cut amounts by one fifth--better
to do several smaller reactions.
3. on p. 207, after Dowex 50, gas evolves--this must be done under
vac., otherwise it takes too long and degradation occurs. Follow
the progress of the reaction by paper electrophoresis and silver
staining plus ninhydirin staining of parallel samples to follow
production of mannopinic acid and disappearance of glutamine.
4. The above information was supplied by the person who did all
of the synthesis for Tempe. His name is Pierre Guyon and he says
that you can easily make 60 grams in 2 weeks using the above.
If you so desire he would be happy to pay you a visit and make
some mannopine for you. He is a very fine person, a couple of
years from retirement. He says he is sorry that he didn't take
any mannopine with him when he left Tempe's lab.
Polysome Isolation from Alfalfa Suspension Cells
1. Grind fresh or frozen tissues in liquid nitrogen with a mortar
and pestle to fine power.
2. Add enough polysome isolation buffer (PIB) to wet the power
( 2 ml/g tissue) and allow to thaw. Grind further for 2 min,
then add more PIB (4-5 ml/g tissue) and grind for another 1 or
2 min.
3. Centrifuge at 5000 rpm (rotor JA20) for 10 min at 4oC, transer
supernatant to fresh tubes.
4. Add Triton X-100 (50%) to a final concentration of 1%, mix
well and incubate on ice for 10 min.
5. centrifuge at 15,000 rpm (rotor JA20, 27000 g) for 15 min at
4oC. At the same time, add 3 ml sucrose cushion to each sealable
tube.
6. Layer the supernatant to sucrose cushion using a sterile pipette,
about 8.5 ml per tube, seal the tubes.
7. Centrifuge at 44000 rpm (T-1270, Serva) for 3.5 to 4 hr at
4oC.
8. After centrifugation, punch a hole on shoulder of the sealable
tube and use a syrenger to remove 2-3 ml of liquid through second
hole on shoulder, cut top off with a lazer blader, remove all
liquid with a pipetter connected to a vacumn system. Pellets are
rather firm and you do not have to worry about disturbing the
pellets.
9. Depend on what you want to do with the polysome, resuspend
the pellets with appropriate buffer. For polysomal isolation,
use following buffer, I usually use 100 ul for each tube.
For in vitro translation, following buffer should be used, I
usually use 50 ul for each tube.
Nuclei Isolation from Alfalfa Suspension Cells
Weiting Ni
I. Buffers and Solutions:
Honda Buffer
25 mM Tris.HCl pH 7.8
0.44 M Sucrose (add before use)
5 mM MgCl2
2.5% Ficoll Type 400 (Sigma)
5% Dextran T40 (Sigma)
10 mM -mercaptoethanol (add before use)
2 mM spermine (add before use).
To 650 ml DEPC treated H2O, add 3.025g Tris, 5 ml 1 M MgCl2 and
pH to 7.8 with 1 M HCl, then add 25 g Ficoll and 50 g Dextran.
Heat the solution to about 45 C with stirring. The solution may
be turbid, do not filter. QS to 770 ml with DEPC treated water.
Autoclave for 30 min.
2 M Sucrose
342 g Sucrose in 250 ml H2O, QS to 500 ml with water, add 0.25
ml DEPC. Incubate O/N and autoclave for 30 min.
Nuclei Washing Buffer (NWB)
50 mM Tris.HCl pH 8.5
5 mM MgCl2
10 mM -mercaptoethanol (add before use)
20% Glycerol
To 380 ml DEPC treated water, add 3.025 g Tris, 2.5 ml 1 M MgCl2,
pH to 8.5. Add 100 ml glycerol, QS to 500 ml with DEPC treated
water. Autoclave for 30 min.
Nuclei Resuspension Buffer (NRB)
50 mM Tris.HCl pH 8.5
5 mM MgCl2
10 mM -mercaptoethanol (add before use)
50% Glycerol
To 230 ml DEPC treated water, add 3.025 g Tris, 2.5 ml 1 M MgCl2,
pH to 8.5 with 5 M HCl. Add 250 ml glycerol and QS to 500 ml with
DEPC water. Autoclave for 30 min.
II. Nuclei Isolation
1. Prepare Honda buffer
77 ml Honda buffer
22 ml 2 M sucrose
78 ul -mercaptoethanol
1 ul 200 mM spermine
1 ml triton X-100 (optional)
Mix well and place on ice
2. Grind 5-10 g cells with a mortar and pestle in liquid N2
to fine powder, transfer the powder to a glass homogenizer, add
5-10 ml Honda buffer to homogenize for 1 min.
3. Filter the homogenate through 4 layers of Myracloth, the
Myracloth is pre-wetted with Honda buffer.
4. Centrifuge at 1000 g for 5 min, resuspend the pellets with
20 ml Honda buffer. Soft brush may employed to help resuspending
the pellets. It is always a good idea to check the intactness
of the nuclei and contamination at each step from now on. Large
starch grain, if exist, can be removed by centrifuge at 100 g
for 5 min.
5. Centrifuge as above and resuspend the pellets in 20 ml nuclei
washing buffer (NWB).
6. Centrifuge again as above and the nuclei pellets should be
white in color (slightly grayish). If necessary, wash the nuclei
again with 20 ml NWB (check with a microscope).
7. Resuspend nuclei pellets with 20 ml nuclei resuspension buffer
(NRB) and pellet the nuclei at 1000 g at 4 oC for 5 min. Remove
NRB and allow residual NRB at the bottom of the tube to resuspend
the nuclei. Final vol. is about 0.5-1 ml.
8. Aliquot 0.1 ml to Eppendorf tube and freeze with liquid nitrogen.
The nuclei so prepared remain active after five times freeze
and thaw.
"RUN ON" TRANSCRIPTION WITH COTTON COTYLEDON NUCLEI
All steps are done on ice except indicated.
Take 50 ul nuclei, spin for 10 sec. and remove 25 ul of supernatant,
then add:
10 ul 1 M (NH4)2SO4 100 mM
4 ul 10 mM MgCl2 4 mM
1 ul 100 uM Phosphocreatine 1 uM
4 ul 0.25 mg/ml Phosphocreatine Kinase 10 ug/ml
12.5 ul RNAsin 500 U
31.5 ul H2O
2 ul NTP mix 500 uM each
1 ul UTP 30 uM for UTP
10 ul 32P-UTP 100 uCi
---------------------------------------------------------------
Total 100 ul
Incubate at 30oC
Stop reaction with 10 fold stop buffer (1% SDS, 10 Mm EDTA)
To test RNA stability: add 1 ul 100 mM UTP after 10 min
incubation and chase for 50 min.
To determine linear range of 32P-UTP incorporation:
Take 10 ul reaction mix at following time points: 5 10 20 30
45 60 min and directly add to 100 ul stop buffer. Vortex for
20 sec. and spot 100 ul to DE81 filter, let air dry or use a lamp.
Wash the filter with 0.5 M Phosphate buffer (Ph 7.2) 3 times,
4 min each; water 3 times, 4 min each and finally with 100% ethanol.
Air dry the filter and count in non-aqueous fluid.
Or alternatively use spin columns.
Usually the incorporation is linear at first 10 to 20 min incubation.
RNA extraction
After stop the reaction with 1% SDS, 10 mM EDTA,
1. Add 0.1 vol of 2 M sodium acetate,
2. Extract with equal vol. of chloroform:phenol (1:1),
3. Incubate at 55oC for 5 min. and then cool on ice for 5 min.
4. Centrifuge at 10,000 g for 15 min. at 4oC.
5. Extract aqueous phase with chloroform:phenol and centrifuge
as step 4.
6. Add 2 vol of ethanol to aqueous phase and mix well and incubate
at -70oC for 1 h.
7. Pellet RNA at 10,000 g for 15 min. at 4oC.
8. Wash pellets (resuspend) with 70% ethanol and recover RNA
as step 7.
9. Repeat step 8.
10. Dry RNA pellets and resuspend in 100 ul DEPCed water and
pass a spun column.
Note: stop at step 7 if the RNA is used for RNA gel analysis.
IN VITRO TRANSLATION OF POTATO TUBER mRNA
The reaction mixtures are prepared in 0.5 ml vials as follows
(volumes in ul)-
vial # 1 2 3 4 5 6
water 4 3 2 1 0 BMV
mRNA (0.5ug/ul) 0 1 2 3 4 RNA
translation cocktail 9.5 9.5 9.5 9.5 9.5 9.5
Add the water to the vials then the RNA and then make up the translation
cocktail. The stocks for making up the cocktail are kept on ice
as is the vial in which the cocktail will be mixed.
The translation cocktail is made up as follows (volumes in ul)-
1X 8X
amino acids (minus methionine) 1 mM stock 0.2 1.6
35-S metionine (10 to 15 uCi/ul) 1.5 12.0
RNasin (40 units/ul) 0.4 3.2
Promega reticulocyte lysate 7.0 56.0
Keep the cocktail on ice; add the lysate to the cocktail just
before adding the cocktail to the reaction mixture. Mix the cocktail
gently with the pipet.
After adding the cocktail to the reaction mixture briefly vortex
the vials GENTLY and spin the tubes just enough to bring the droplets
to the bottom of the vials. Incubate the reaction vials at 30
C for 60 min. Terminate the reaction by freezing at -20.
formamide for hybridization and blotting membranes.
As the poster surmised, formamide reduces the Tm of DNADNA,
DNARNA, and RNARNA hybrids. For DNADNA, the effect is 0.7oC/%
formamide. So why not do hybridizations at room temp in 60%
formamide? Because, in addition to reducing the Tm, formamide
reduces the nucleationrateconstant for reassociation, so that
the reaction would take 5 10X as long to hybridize to an equal
extent. How do people get away with using formamide? Either
they (1) have such a high concentration of probe (or DNA on the
filter in the case o plasmids) that it does not matter, or (2)
they use a accelerating agent, such as dextran sulfate, which
provides a 10X speed up, or (3) their blots don't work very well.
For genomic blots, dextran sulfate will let you use formamide
and still get good results. The problem is, especially for nitrocellulose,
dextran sulfate + formamide can cause horrendous background problems.
We routinely do genomic southern blots and northern (RNA) blots
at 65oC with no formamide, no dextran sulfate, hybridizing for
40+ hours with excellent results. I do not believe there is
any good reason to use formamide, especially now that nylon membranes
are available that are not affected by high temperature. Most
people who use formamide do not realize that it slows down the
hybridization reaction, so that usually there is a net decrease
in hybridization.
Re: nylon vs nitrocellulose. Nitrocellulose has a marginally
higher capacity for DNA or RNA, this can be important for dotblots
and slotblots. Otherwise, it is much more fragile (it tears
very easily), it ages very poorly, it is more difficult to handle
(wetting can be difficult), and it is much less reproducible
from lot to lot. I blotted for 5+ years on nitrocellulose before
changing to nylon (AMF Cuno's Zetabind or S+S Nytran) and I would
never go back. No baking, no tearing, no background, its great.
Subject: Re: formamide/ membranes for hybridizations.
Just a quick reply to why someone might use formamide in hybridizations.
I was doing some Northerns with an Algal gene probe, where we
needed the highest possible stringency, we used 1X SSC, 60 %
formamide, and a temperature of 65 oC. Without the formamide
the hybridization solution would have to have been >100 oC!
We also found that our genomic blots were better with the formamide/dextran
sulfate procedure, but only if the probe concentration was 5
10 ng/ml. With higher probe concentrations the background was
truly horrendous.
In article
<1991Nov5.164459.13185@murdoch.acc.Virginia.EDU>,
wrp@cyclops.micr.Virginia.EDU (Bill Pearson) writes:
> Regarding formamide for hybridization and blotting membranes.
>
> As the poster surmised, formamide reduces the Tm of DNADNA,
>DNARNA, and RNARNA hybrids. For DNADNA, the effect is 0.7oC/%
>formamide. So why not do hybridizations at room temp in 60%
>formamide? Because, in addition to reducing the Tm, formamide
reduces
>the nucleationrateconstant for reassociation, so that the
reaction
>would take 5 10X as long to hybridize to an equal extent.
How do
>people get away with using formamide? Either they (1) have
such a
>high concentration of probe (or DNA on the filter in the case
o
>plasmids) that it does not matter, or (2) they use a accelerating
>agent, such as dextran sulfate, which provides a 10X speed
up, or (3)
>their blots don't work very well. For genomic blots, dextran
sulfate
>will let you use formamide and still get good results. The
problem
>is, especially for nitrocellulose, dextran sulfate + formamide
can
>cause horrendous background problems.
>
> We routinely do genomic southern blots and northern (RNA)
>blots at 65oC with no formamide, no dextran sulfate, hybridizing
for
>40+ hours with excellent results. I do not believe there
is any good
>reason to use formamide, especially now that nylon membranes
are
>available that are not affected by high temperature. Most
people who
>use formamide do not realize that it slows down the hybridization
>reaction, so that usually there is a net decrease in hybridization.
>
> Re: nylon vs nitrocellulose. Nitrocellulose has a marginally
>higher capacity for DNA or RNA, this can be important for
dotblots and
>slotblots. Otherwise, it is much more fragile (it tears very
>easily), it ages very poorly, it is more difficult to handle
(wetting
>can be difficult), and it is much less reproducible from lot
to lot.
>I blotted for 5+ years on nitrocellulose before changing to
nylon (AMF
>Cuno's Zetabind or S+S Nytran) and I would never go back.
No baking,
>no tearing, no background, its great.
>
>Bill Pearson
freeze & squeeze, for isolating DNA from lowmelt agarose
"oligo GEB": from Meth. Enz. 65, 506.
500mM NH4OAc 1 ml of 5M stock
10mM MgOAc 0.1 ml of 1M stock
1mM EDTA 0.02 ml of 0.5M stock
ddH2O to 10 ml
Some folks also add a small amount of SDS to this buffer and say
it helps. I have tried the protocol you mention and there are
commercially available spin columns for this but if you use the
pinhole, you may wish to add some siliconized glass wool to prevent
the gel from being extruded through the hole.
We use the following:
Elution of DNA from agarose gels
1. The DNA sample is loaded onto a lowmelting preparative agarose
gel (0.7 to 1.0%), and electrophoresed as required. Ethidium
bromide (20 5l of 10 mg./ml) may be included in the gel.
2. Visualize DNA bands on a long wave UV glow box and photograph.
Excise the desired DNA bands with a clean, ethanolwashed scalpel.
Transfer the gel slice to a clean microfuge tube.
3. Melt the agarose at 65 deg C for 5 minutes. Add one volume
of TEsaturated phenol and vortex for a few seconds. (Optional:
centrifuge for 3 minutes and then skip to step 6.)
4. Freeze the agarosephenol mixture at 70 deg C for at least
15 minutes.
5. Thaw the sample for a few minutes at room temperature. Centrifuge
for 3 minutes.
6. Remove the aqueous phase to a clean microfuge tube. Add onehalf
volume of phenol, vortex and centrifuge for 2 minutes. Repeat
phenol extraction one more time.
7. If sample volume is 0.5 ml or less, skip to step 8. If sample
volume is greater than 0.5 ml, add 1.5 volumes of nbutanol, vortex
and centrifuge for 2 minutes. Remove butanol (upper) phase and
discard. Butanol extraction may be repeated until the sample
volume is 0.5 ml or less.
8. Extract once with one volume of watersaturated diethyl ether.
Centrifuge for 1 minute to separate phases, discard ether (upper)
phase.
9. Add onetenth volume of 3M NaOAc, pH 5.2 and 2 volumes of cold
95% ethanol. Precipitate DNA by storing at 70 deg C for at least
one hour.
10. Pellet the DNA by centrifugation for 15 minutes at 4 deg
C. Wash once with 0.5 ml of 70% ethanol and dry for a few minutes
under vacuum.
Subject: BETTER METHOD THAN: Freeze & Squeeze: DNA from Lowmelt
In article <1991Oct23.233111.18225@agate.berkeley.edu> nigel@codon1.berkeley.edu
(Nigel Walker) writes:
>He has described to me a technique which he calls freeze &
squeeze, for
>isolating DNA from lowmelt agarose. The excized band is diluted
in buffer 1
>[this is where the cookbook is important!], frozen with liq
N2, and centrifuged
>through a tiny hole in the bottom of a 0.5ml eppendorf tube
(into a 1.5ml tube)
Many of the responses to this post have included protocols using
extractions and glass wool. We have had much success recently
with a much simpler protocol adapted from a blurb in the April
1991 TIG (p. 119?). Instead of glass wool, you use a piece
of Whatman filter paper. If done correctly, the paper prevents
the agarose from entering the 'eluate'. There is no need for
freezing or extraction. We use a very high grade agarose to
minimize impurities, and get excellent yields from TBE gels.
The resulting eluate can be directly added to a ligation mixture.
This proceedure has saved me gobs of time and is practically
effortless. I highly recommend giving it a try.
Subject: Elution of DNA from agarose gels
K.R. Raj writes:
>I am quite tired of electroeluting DNA out of agarose. Could
>anyone suggest a better way other than using the geneclean
kit ?
> Thankyou.
> kraj@uk.ac.crc
I've had a lot of success with simple phenol extraction of melted
lowgelling temperature agarose. Cut out the bands, trim away
excess
agarose, place the gel slice in a 1.5 ml microfuge tube. Place
at
65 degrees for 5 minutes. Bring volume to 0.5 ml with TE buffer.
Add
an equal volume of Trissaturated (pH 8) phenol, vortex vigorously,
place at 70 degrees for at least 15 minutes (or on dry ice for
at
least 5 minutes). Remove tube from freezer (or dry ice) and place
directly in a microfuge. Spin at top speed for 5 minutes. Remove
the
aqueous (top) phase to another microfuge tube. Extract twice
with
onehalf volume of phenol, and once with one volume of watersaturated
ether. Add onetenth volume of 3M NaOAc, pH 5.2 and two volumes
of
ethanol. Place at 70 degrees for at least 15 minutes. Centrifuge
at
room temperature for 15 minutes to pellet DNA. Wash once with
1 ml of
70 percent ethanol and dry briefly under vacuum.
In my hands, recovery of 8595 percent is typical. In most cases,
I
use the eluted DNA for cloning or direct sequencing with no further
purification.
I just want to second this, and to add that for many procedures,
we
find that we do not need to remove the agarose at all. All of
our
ligations, sequencing, and random oligo primed labelling are now
done
in low melting point agarose directly. Melt the agarose at 65
degrees
C for five minutes, cool to 37 degrees, mix components, and run
the
reaction at the usual temperature. (It doesn't matter if the
agarose
hardens again durring the reaction). We have also done restriction
digests this way, but for technical reasons it is usually easier
to
purify the reprecipitate the DNA first in this case.
BTW, people in my lab are quite superstitious about sources and
lots
of agarose. The only LMP agarose they will use is Genetic Technology
Grade SeaPlaque GTG agarose from FMC BioProducts. I have no
connection with FMC except as a satisfied customer, and I certainly
don't mean to suggest that other brands don't work. I just want
to
suggest that if you have difficulties, you might try a different
source of agarose.
Subject: RE: Elution of DNA from agarose gels
I use a method presented in Trends in Genetics, June 1990, 6(6).
The
basic procedure is to make a small hole in the bottom of a 500
ul tube (I use a
22 guage needle), place a small amount of siliconized glass wool
in the bottom,
then place the gel slice (from a 1% gel) inside the tube. Remove
the cap, place
the tube in a 1.5 ml eppendorf tube, and spin in a microfuge at
6000 rpm for 10
minutes. The collected DNA is then purified by phenol:chloroform
then
chloroform extractions and ethanol precipitation. You can expect
a recovery of
over 90% for fragments from 100 to 3000 base pairs, and a slightly
lower
recovery above this range. DNA collected by this method is quite
adequate for
subcloning and for preparation of radiolabeled probes.
Subject: Re: Elution of DNA from agarose gels
We have a good deal of experience cloning fragments extracted
from agarose gels
by electrophoresis into a strip of DEAE cellulose paper (DE81,
Whatman). The
method we use in the lab is as follows:
1) run a preparative agarose gel (0.6% to 1%, depending on the
size of the
fragment of interest) containing 0.5 ug/ml ethidium bromide.
2) Transilluminate with a long wave lenght UV source (do not use
short wave to
avoid nicking).
3) Make a cut parallel to the fragment just in front of it with
a razor blade.
Do not cut the entire gel because the pieces will float in the
tank later.
4) Insert a 34 mm width strip of DE 81 paper (previously autoclaved)
and
continue the electrophoresis at about 50 mAmps for 5 min.
5) Take the paper into a 250 ul eppendorf tube with a small hole
at the bottom,
and put this tube into a 1.5 ml eppendorf tube.
6) Wash 5 times with 100 ul of TE with brief spins between washes.
7) Transfer the small microfuge tubes to clean 1.5 ml tubes.
8) Elute the DNA with 5 times 100 ul of 1M NaCl.
9) Add 1 ml ethanol and keep at 20 for at least 30 min.
10) Spin down, wash with 70% ETOH, dry and go for it!
Some precautions:
a) If your DNA fragment is small and migrates with the dye front,
use xylene
cyanol dye instead of bromophenol blue. BFB will be eluted and
affect ligation.
b) Make a small hole with a heated needle. Big holes will let
pieces of paper
come through. This also affects ligation.
c) Do all the steps with gloves.
We have cloned fragments of various sizes, and also used DNA fragments
isolated
by this method (even a 14 Kb fragment) as templates for in vitro
transcriptiontranslation. By the same token, we use this method
for preparing
probes.
The only expense is the paper (about $300), but there comes so
much that you
buy it only once in a life time.
Miguel A. Valvano, Microbiology and Immunology, Univerisity of
Western Ontario,
London, Ontario, Canada, N6A 5C1.
> reaction at the usual temperature. (It doesn't matter if
the agarose
> hardens again durring the reaction). We have also done restriction
> digests this way, but for technical reasons it is usually
easier to
> purify the reprecipitate the DNA first in this case.
>
> BTW, people in my lab are quite superstitious about sources
and lots
> of agarose. The only LMP agarose they will use is Genetic
Technology
> Grade SeaPlaque GTG agarose from FMC BioProducts. I have
no
> connection with FMC except as a satisfied customer, and I
certainly
> don't mean to suggest that other brands don't work. I just
want to
> suggest that if you have difficulties, you might try a different
> source of agarose.
>
Subject: Re: Freeze & Squeeze: DNA from Lowmelt
Freeze squeeze protocol for extraction of DNA or RNA from agarose
gels.
Ref: Locker,J. (1979) Anal. Biochem. 98; 358367
Tautz, D. & Renz, M. (1983) Anal. Biochem. 132; 1419.
1) Excise band from gel and place into 0.5ml microcentrifuge
tube plugged
at the bottom with siliconized glass wool.
2) Add 400ul of 0.4M TrisHCl, pH 7.8, 0.3M MaCl, 2mM EDTA and
freeze
the tube in liquid Nitrogen (dry ice also works)
3) Pierce the bottom with a 25 gauge needle heated in a flame.
4) Place the 0.5ml centrifuge tube (still frozen) into a 1.5ml
tube
and centrifuge at full speed for 5 minutes. If some agarose
pieces
remain, repeat the centifugation.
5) Extract the pooled eluate with secbutanol to reduce volume
and remove
EtBr then EtOH ppt.
6) DNA ready for use.
RNA extraction
1) Run a methyl mercury gel and remove the lane of interest,
but don't stain
Place agarose in 10mM NaPO4, pH 7.6, 10mM BEtSH
2) Remove and stain a lane of markers and use to estimate position
of the
RNA band you want to extract.
3) Remove band of interest and extract RNA as described for DNA
4) Adjust salt and ethanol ppt RNA directly following the extraction
without
butanol extraction
5) The RNA can be used for cDNA synthesis or translation
In your posting, you requested information about other methods
to elute DNA
from agarose besides electroelution and Geneclean (glass milk?).
Do you really need to elute the DNA? For example, if you use
one of FMC's
high quality low melt agaroses such as SeaPlaque or NuSieve, you
can perform
ligation, restriction digestion, labelling, etc. with the gel
present.
However, if you really do need to get rid of the agarose, here
are two methods
that I use.
First of all, for both of these methods, you still need to use
SeaPlaque or
NuSieve (or some other highquality, low melting point agarose)
and you need
to run the gel in TrisacetateEDTA buffer (not TrisborateEDTA).
1. Warm phenol method After slicing out your band from the gel,
add 100 ul
of sterile distilled water per lane, and melt at 65C for 10 min.
At the same
time, heat some Trissaturated phenol in a separate tube in
the 65C bath. Add an equal volume of warm phenol to the DNAagarose
and
vortex vigorously. Microfuge to separate phases and remove the
aqueous phase
to a fresh tube. Add another equal volume of TE to the organic
phase and heat
at 65C again for 10 more minutes. Vortex as before and separate
phases.
Combine the two aqueous phases and ether extract. Then, precipitate
the DNA
with 2 volumes of cold (20C) EtOH. You don't have to add any
salt. Leave on
ice for 10 min and microfuge 15 min. Wash the pellet with 70%
EtOH and dry.
Resuspend in TE or dH2O. This DNA can be used for ligation, digestion,
labelling, etc.
2. Agarase digestion Use one of the commercially available agarases
to
digest the agarose present in the slice and then just EtOH precipitate
the DNA
without any extractions. Both New England Biolabs and Epicentre
Technologies
have agarases available. Epicentre calls theirs "GELase".
I have used GELase
many times with good results. It is especially nice if you have
to do a second
restriction digest and then run a second gel. This is hard to
do with the
agarose present because it has to remain hot while you load it
on the second
gel!
Subject: Doublestranded DNA sequencing
Hi! Does anyone have experience with doublestranded DNA sequencing
using
Sequenase version 2.o of USB? I would appreciate ANY information
for
optimization of results. Thanks in advance.
Use 510 ug DNA, add 0.2M NaOH (final) for 30' at 37 deg.
EtOH precipitate and follow the USB protocol. Load 4ul
on the sequencing gel. DNA miniprepped by the boiling
method (Maniatis) gives good results.
Subject: summary Double Stranded DNA Sequencing
Hi! I want to summarize what I have been doing for my double stranded
sequencing reactions. Sorry it took me so long but I was out
of town
for a conference during the last two weeks. As I mentioned in
a
previous minisummary the most critical aspect seems to be the
cleanliness of the template DNA to be sequenced. The following
is the
miniprep protocol that I am using which is from INVITROGEN:
MINI Plasmid Preparation Protocol
(Reference: Zhou et. al., Biotechniques Vol 8., No. 2:172 (1990))
1. Grow 2 mls of bacterial culture at 37 C overnight.
2. Spin 1.5 mls of the culture in a microcentrifuge tube for 10
sec, decant supernatant leaving 50100 ul of medium in the tube.
Vortex
to resuspend cells completely.
3. Add 300 ul of TENS (10mM TrisHCl, pH 7.5, 1mM EDTA, 0.1N NaOH,
0.5% SDS), vortex for 25 sec.
4. Add 150 ul of 3.0M Na (or K) Acetate, pH 5.2, vortex for 25
sec.
5. Spin for 2 min to pellet cell debris and chromosomal DNA.
Transfer supernatant (just 300 ul) to a fresh tube, add 900 ul
100%
ETOH and vortex. Freeze at 70 C for 30 min.
6. Spin 5 min to pellet plasmid and RNA. The pellet should have
a
white appearance. Discard supernatant and wash pellet twice
with 1 ml
of 70% ETOH. Remove any residual ETOH.
7. Resuspend pellet in 2050 ul of sterile H2O.
Note that this protocol does not require phenol:chloroform extractions
or RNAse treatment.
Following this procedure I clean my plasmid DNA with prepAGene
(BIORAD). PrepAGene gets rid of RNA and any contaminant on your
DNA
sample.
The next step is to denature the plasmid DNA. I have been using
a
modification of a protocol sent by William J. Buikema of the Univ.
of
Chicago.
Alkaline Denaturation:
1. ETOH precipitate the DNA obtained from PREP_A_GENE (I use large
amounts of elution buffer to increase my yield.
2. Dissolve pellet in 20 ul of Sterile H2O and add 2 ul of freshly
prepared (I don't know how critical is it to use freshly prepared
solution since I did all my sequencing reactions (20) at once,
and I
haven't repeated any yet) 2 N NaOH, 2 mM EDTA. Mix well and
incubate
for 30 min at 37 C.
3. Add 3XETOH (100%) mix and incubate for 20 min at 70 C.
The next step is to do the sequencing reactions. I followed the
protocol of Sequenase version 2.0 (using their Manganese buffer).
My results have always (consistently) been optimum. I am able
to read
200250 bases.
Subject: quick n' easy plasmid sequencing
I would like to share a quick n'easy plasmid sequencing protocol
we have
recently lifted, which works really well.
Kuchuan Hsiao : A fast ands simple procedure for sequencing double
stranded
DNA with Sequenase . NAR 19:2787
Basically you simply dissolve clean miniprep plasmid DNA in 2550ul
TE;
to 5ul add 1ul appropriately diluted sequencing primer (10ng/ul);
add 1ul 1M NaOH & mix;
37 deg C 10'
add 1ul 1M HCL & mix;
add 2ul 5X Sequenase buffer
(optional: 37 deg C 5 min)
and thereafter go for it according to the Sequenase manual.
Cuts plasmid sequencing time down drastically, and will really
be a help when
sequencing more than one template.
Subject: Re: dsDNA cleanup with Sephadex
>Does anyone out there have a suitable method for the removal
of nucleotides
>and, if possible, 20 mer primers from a reaction mixture using
Sephadex
>in a home made spin column. The dsDNA in the mixture is at
least 120 bp and
>requires some concentration as well as clean up.
>The sample should also be desalted in the clean up step.
>
>Any help as to column preparation, spin times and gforce would
be greatly
>appreciated.
>PS Any suggestions for alternative techniques are welcome.
(But we haven't
>got a centrifuge that will take Centricon devices!!)
>
Amicon (I think) or maybe Schleicher and Schule makes a centriconlike
device
that is the size and shape of a microfuge tube....
This might be an option.
We use spin columns with p6DG (from BioRad) frequently but I'm
not sure
you can separate 120 nt from primers or worse primerdimers (~40
nt?)
Subject: Re: dsDNA cleanup with Sephadex
I make homemade spin columns in a 1 ml syrenge with a little glass
wool
stuffed in the bottom (can use silicon wool if you want to be
sure it is
not sticky) and fill syringe full of column matrix. Can use sephadex
G50
we prefer biorad P10. Put the column in a 15 ml falcon tube, spin
at 2000 RPB
oops, 2000 RPM for 2 minutes, the matrix should now look dry.
cut the lid off
a 1.5 ml microtube and put in bottom of falcon tube, put syringe
end inside
1.5ml tube, load sample, and spin 2000RPM for 2 minutes. I have
found this
really only works reliably in a swinging bucket rotor. Speed
and time are
variable, but do both spins the same. Discard syringe and your
sample is in
the micro tube. This matrix works well for desalting and removing
nucleotides. If you want to remove linkers can use other matrices,
some
commercial spin columns use CL2b, CL4B and CL6B to separate larger
oligos
from each other. I have no experience with those, but they should
work. Just
need to use a strong bead like an acrylamide bead so that it does
not crush
like agarose beads do.
Subject: Re: quick n' easy plasmid sequencing
In article <201153.292a4f30@uctvax.uct.ac.za>, rybicki@uctvax.uct.ac.za
writes:
> I would like to share a quick n'easy plasmid sequencing protocol
we have
> recently lifted, which works really well.
>
> Kuchuan Hsiao : A fast ands simple procedure for sequencing
double stranded
> DNA with Sequenase . NAR 19:2787
>
> Basically you simply dissolve clean miniprep plasmid DNA
in 2550ul TE;
>
> to 5ul add 1ul appropriately diluted sequencing primer (10ng/ul);
>
> add 1ul 1M NaOH & mix;
>
> 37 deg C 10'
>
> add 1ul 1M HCL & mix;
>
> add 2ul 5X Sequenase buffer
>
> (optional: 37 deg C 5 min)
>
> and thereafter go for it according to the Sequenase manual.
>
> Cuts plasmid sequencing time down drastically, and will really
be a help when
> sequencing more than one template.
>
> Ed Rybicki
Modification by Di James (Microbiology Dept, UCT)
FOR GCRICH TEMPLATES
after addition NaOH, INCUBATE 40 DEG C 30 MIN
KEEP AT 40 WHILE ADDING HCl AND 5X RXN BUFFER
COOL VIAL TO ROOM TEMP SLOWLY!! (+/ 30 MIN)
Suggestion: allow tupperware with water at 40degC to cool on bench
(with vial
in boat).
Subject: Re: quick n' easy plasmid sequencing
>> Basically you simply dissolve clean miniprep plasmid
DNA in 2550ul TE;
>>
>> to 5ul add 1ul appropriately diluted sequencing primer
(10ng/ul);
>>
>> add 1ul 1M NaOH & mix;
>>
>> 37 deg C 10'
>>
>> add 1ul 1M HCL & mix;
>>
>> add 2ul 5X Sequenase buffer
>>
>> (optional: 37 deg C 5 min)
>>
>> and thereafter go for it according to the Sequenase manual.
>>
>> Cuts plasmid sequencing time down drastically, and will
really be a help when
>> sequencing more than one template.
>>
>> Ed Rybicki
>
>
> Modification by Di James (Microbiology Dept, UCT)
>
> FOR GCRICH TEMPLATES
> I do not use miniprep DNA but 10ug of NUCLEOBOND AX preps
and get good
resolution of 600700 bases in most cases.
> after addition NaOH, INCUBATE 40 DEG C 30 MIN
>
> KEEP AT 40 WHILE ADDING HCl AND 5X RXN BUFFER
>
> COOL VIAL TO ROOM TEMP SLOWLY!! (+/ 30 MIN)
>
> Suggestion: allow tupperware with water at 40degC to cool
on bench (with vial
> in boat).
>
>I then continue using Sequenase and ROB SCHUURMAN AND WILCO
KEULEN'S
MODIFICATION utilising KLENOW POLYMERASE and HIGHER TERMINATING
TEMPERATURE.
Subject: Re: ss phagemid DNA
>I recently started trying to use Stratagene's "Altered
Sites"
>kit for mutagenesis. I have cloned about 5 kb into their pUC
>derived vector (~5 kb also). Attempts to isolate phagemid
>DNA have been dismal. I can barely see the
>ssDNA bands when loading 1/4 of a prep from 1.2 ml
>of phage supernatant. The E. Coli strain is JM109.
>I have tried growing the cells for 6 hrs or overnight
>in 2XYT + tet after adding either R408 or M13KO7
>helper phage. I have also tried adding 20 mM phosphate.
>My last experience with ss phagemid preps was using pEMBL,
with a much
>smaller insert. Subsequent to this I worked with M13, also
>with a small insert, and thought I was getting a low yield
>THEN, at least by comparison with pEMBL.
>
>Should I try growing the cells tet? Perhaps
>I should decrease the culture volume from 5 ml/50 ml tube
>to 2ml/50 ml tube, in order to improve aeration?
>I could scale this procedure up, but thought perhaps some
useful alternative
>suggestions might come of an inquiry to this group.
>
I have had quite good success with phagemids as long as your helper
phage is good and healthy. A problem with low yields may be due
a low
titre on your helper phage. A second potential problem is your
strain
of E. coli. JM109 is sick, and doesn't respond well to phage
infections especially if a plasmid is there. It tends to lyse
easily.
Try one of the similar but less cripled versions like MV1190 or
TG1.
To avoid satellite colony formation around ampresistant colonies,
we have
had good success with Timentin 3.1, a trademark of Smith/Kline/Beecham.
Timentin 3.1 is a 30:1 mixture of ticarcillin (a penicillin) and
clavulanic
acid. The latter is a substrate for betalactamase that, in a
small percentage
of reaction cycles, leaves a product covalently attached to the
enzyme, thus
inactivating it. In clinical use, timentin is effective against
betalactamase
producers that would otherwise be resistant to penicillins. We
have found that
under laboratory conditions, ampicilin resistant strains such
as those carrying
common plasmid vectors are resistant to timentin at a wide range
of
concentrations. Broth supernates of ampresistant bacteria grown
in the
presence of timentin seem to contain reduced levels of betalactamase
activity,
and formation of satellite colonies around ampresistant transductants
or
transformants is reduced on timentin plates. We have also found
that, in some
cases, overproduction of proteins from plasmid constructs is markedly
enhanced
when the cultures are grown with timentin instead of or in addition
to
ampicillin.
Timentin should be available through conventional pharmaceutical
wholesalers, not through companies that cater to researchers.
Be sure to
request timentin for injection, not the flavored oral formulations.
Timentin
3.1 comes as a vial containing approximately 4 g of powder (3
g of ticarcillin,
0.1 g of clavulanic acid, and the rest as salt mostly potassium,
I think).
It dissolves readily in water and the solution can be stored frozen
more or
less indefinitely (if it turns yellow, pitch it). For plates
or broth, we have
used concentrations of 4080 mcg/ml when replacing ampicillin.
It is cheap
enough not to matter (1020 cents per liter of broth or agar).
There is also Timentin 3.2 containing twice as much clavulanic
acid. We
have no experience with it.
I would be interested to hear of the experiences of anyone
else who has
used Timentin or gives it a shot (ram3@po.cwru.edu). I'll post
a summary if
indicated.
Subject: Re: FUSION PROTEINS FOR IMMUNIZATION?
I tried a number of fusions with trpE and T7 gene 10 before
getting successful expression. The gene I was trying to fuse was
a Dictyostelium protein with several hydrophobic regions my
initial constructs which included these regions were not
overexpressed in E. coil, so I started to delete down the size
of the fusion until it worked. The other esential factor for
success was to avoid growing without antibiotic selection, such
as above an OD of 1.0, and to streak out the glycerol stock to
get a FRESH colony before growing up for the induction. The
procedures are nicely explained in Studier et al, Meth. Enz.,
185: 61 (1990). The precautions that he suggests may seem
extreme, but I assure you that if your protein is hard to
express then they are essential.
I was finally successful using Promega's pGEMEX1 in Studier's
strain BL21(DE3) containing the pLysS plasmid. This plasmid
expresses T7 lysozyme, which is antagonistic to T7 polymerase,
at a low level. The result is a significant drop in the
noninduced expression which just may allow you to get your
fusion. (see Studier (1991) JMB 219, 37 for more info).
I should add that of the three fusions that I made, the two
with the extremely hydrophobic regions were expressed to less
than 0.5% of the total protein and stopped growth completely,
while the other was expressed to ~50% of total protein and
growth would continue for hours.
Subject: TrpEfusion proteins as immunoadsorbents
I am interested in using a TrpEfusion proteins (from a PATH
vector) for the affinity purification of antibodies. The polyclonal
sera was generated by immunization with the protein of interest
fused to the maltosebinding protein. Since high level expression
of
TrpE fusions usually result in the production of insoluble inclusion
bodies, my question is whether anyone has tried to use these
insoluble proteins as immunoabsorbents. The two main questions
are 1) are these proteins completely insoluble or would there
be
some leaching of protein into the antibody preparation and 2)
would
presumably hydrophilic epitopes be exposed in these inclusions
to
make them effective immunoabsorbents. If you have any thoughts
about these questions or any others that you can raise, please
respond. I know of many other possibilities for affinity purification
of antibodies (coupling to activated supports, nitrocellulose
blotted
proteins, glutaraldehyde immobilization inpolyacrylamide gels)
but
thought this might be an interesting method.
Subject: Melting temperature calculations
I need to calculate Tm for DNADNA hybrids and DNARNA hybrids.
I
am thoroughly confused by 3 sources of equations that I have found.
The main confusion relates to the change in Tm in relationship
to GC
content. Typing equations are hard, so I'll just type the relevant
parts of the equations and invite comments and corrections.
The 3 sources are: A. Maniatis, 2nd edition
B. S & S "Transfer and Immobilization..."
1987 ed.
C. S & S "Transfer and Immobilization..."
1990 ed.
For DNADNA hybrids, the relevant functions are
1. 0.41(fraction G+C) (A)
2. 0.41(% G+C) (B)
3. 41(fraction G+C) (C)
For DNARNA hybrids, the relevant functions are
4. 0.58(fraction G+C)+11.8(fraction G+C)^2 (A)
5. 58.4(% G+C) (B)
6. 0.58(fraction G+C)+11.8(fraction G+C)^2 (C)
Since expression (2)=expression (3) and they both came from the
same
place, I tend to believe that and assume that there was a mistake
in
Maniatis. Although expression (4) is identical to expression
(6), and
totally different from expression (5) (even without taking into
consideration
the second order term), I find it difficult to believe that it
is true
because it will wind up giving much lower Tm for DNARNA hybrids
than
a similar DNADNA hybrid under similar conditions based on the
rest of
the formula given in those references.
My *hunch* is the correct expression for DNADNA hybrid is 41(fraction
GC)
and that none of the published expressions for DNARNA hybrids
are correct
(i.e. none of 4, 5, or 6 above) for DNARNA hybrids. For DNARNA
hybrids,
I would guess the real expression ought to be
58(fraction G+C) +11.8(fraction G+C)^2.
I am not a molecular biologist and I have no access to basic nucleic
acid chemistry texts. Would someone please help me resolve this
dilemma?
ccy@po.cwru.edu
Subject: Re: nylon membrane
In article <E5A1E61A99FF40891A@twnas886.bitnet>, YHSUN@imb.as.tw
writes:
>I would like to hear comments on (1) comparison on charged
nylon vs. non
>charged nylon membrane, (2) What are the critical factors
of the various
>manufacturer's conditions for blotting, hybridization, and
washing? Are
>they really critical? Or the standard method works? (3) Has
anyone used 1M
>ammonium acetate for blotting with nylon membranes?
>Thanks.
>
>Y. Henry Sun
>Institute of Molecular Biology
>Academia Sinica
>Taiwan
I don't know the answer to (1) and (2), but I use Amersham's HybondN
membrane for Southerns dot blots and colony/phage lifts, and I've
mostly used ammonium acetate as my neutralize and transfer solution.
I've not noticed a difference compared to TrisHCl neutralize and
SSC
or SSPE transfer buffer.
Subject: Electrophorating with used cuvettes
G'day to JJW, John Nash, Paul Fisher and everyone else out there
who has been on about reusing electrophorator cuvettes (and why
is
it that so many ozzies have views on this...well here's another's).
We have been reusing electrophorator cuvettes for some time now
and havn't had any problems. Since >90% of the time we use
electrophoration to rescue plasmids passaged through
insect embryos, we have had to develop a rather thorough procedure
to guard against cross contamination. Immediately after use, cuvettes
are thoroughly rinsed in H20 and left to soak. At the end of the
day
we then incubate them overnight at 37oC in a solution of 50mM
Tris,
10mM MnCl2 and 2ug/ml of DNase. Cuvettes are then washed extensively
in
H20 and then left in 100% EtOH. Just before use we air dry them
to remove
the EtOH, but I guess that an 80oC dry sterilization would probably
be better.
Since starting to reuse cuvettes we always include a no DNA control,
and on only a few occasions have we seen a colony, but none
looked like coli.
For general transformation purposes though I'd recommend one of
the
procedures already posted (using DNAse is a double edged sword).
Subject: How to grow bacteria in microtiter wells?
C Liang writes Does anyone know how to grow bacteria directly
in microtiter plates, with
glycerol already in the growth medium? Do you shake the plate
while growing?
Is the growth media different than LB? We are making a library
and this would
really help us out.
We do this routinely: To 500 ml LB broth add K2HPO4 3.15g, KH2PO4
0.9g, Na citrate 0.225g, MgSO4.7H2O 0.045g, glycerol 22g, aliquot
and autoclave.
The bugs grow fine at 37 deg without shaking; I grow them for
12 hrs before freezing at 80. Hope this helps.
Subject: Re: Oligo length check
>I have had some trouble with my oligos, please help.
>
>Question:
>
>If i run a 20%, (95% akrylamide/5% bis) urea denaturing sequencing
gel loading
>5001000 ng oligo, will i detect one base length differences
after ethidium
>bromide staining, or will the base composition affect migration
too much? I run
>the gel in a BioRad miniprotean device (about 10 cm cel).
My oligos are mostly
>2022 mers with a couple of 16 and 25mers.
>
>Anyone with experience of such a check please respond. Help
is very much
>appreciated.
>
> Best regards, Mika Salminen
>
> msalminen@finnphi (Bitnet)
> msalminen@nphi.fi (Internet)
>
Mika
Bad idea.
Reason 1:
The oligo is liable to soak out in the ethidium stain, leading
to loss and
cross contamination
Reason 2: this gel is too little for good separation, though
we use 16 cm
gels that work ok
The best way to localize the oligo band is with uvshadowing.
The new
Maniatis (Sambrook) has a good description of this method, and
how to
purify the oligo after soaking it out.
2 notes: for uv shadowing they recommend a silica gel plate with
uv
indicator... If you don't have one handy, many (but not all) intensifying
screens will work. The best way to do this is have someone _show_
you;
I've seen folks struggle with no fluorescence. When it works
it's easy.
Note 2: Are you sure you need to purify the oligo? why? most
applications work with "crude" oligos. Try it out,
maybe?
good luck,
dennis
Subject: Re: Oligo length check
> Reason 1:
>
> The oligo is liable to soak out in the ethidium stain, leading
to loss and
> cross contamination
Not true; we routinely analyze all the oligos we make by running
them
in a 20 % gel and afterwards staining with ethidum bromide. Soaking
is not a problem, and staining is a good way of checking the amount
of
nonfulllength oligo. We have analysed over 600 oligos this way,
and
we have also noticed an interesting point, which is that using
non
denaturing gels the secondary structures in the oligos are conveniently
detected.
> Note 2: Are you sure you need to purify the oligo? why?
most
> applications work with "crude" oligos. Try it
out, maybe?
Subject: Acid-phenol extraction of circular DNA
Does anyone have any experience with the purification of closed
circular DNA by extraction from low-ionic-strength, low-pH buffer,
using phenol equilibrated to pH 4.0 with NaOAc? This method was
published by Zasloff et al. '78 (NAR 5:1139-1152), and it looks
both
easier and higher-yield than ethidium bromide-CsCl centrifugation,
but
it doesn't seem to be in common use. The claim is that, for reasons
"not as yet understood" (any ideas?), linear and nicked
circular DNA
partition into the organic phase under these conditions, while
covalently closed rings remain in the aq. phase. The method is
good
at least for rings 4 to 40kD in size (no data outside that side
range), gives quantitative (close to 100%) recovery, provides
>99%
purity with 3 iterations, and works on samples containing as little
as
..01% closed circular DNA. The acidic depurination rate was reported
to be 1/30,000 bases/hr, which seems acceptable. The only
disadvantage of the method is that it does not eliminate linear
DNA
<1500bp.
What I'm trying to do is to isolate and clone extrachromosomal
closed
circular DNA (eccDNA, aka cccDNA, aka spcDNA) from mammalian cells
growing in vitro. The above method, originally developed for
plasmid
preps, has in fact been used for this purpose [Truett et al. '81
Cell
24:753-763], but it doesn't seem to have caught on, so I'd appreciate
hearing from anybody who has actually tried it, either for plasmids
or
for organelle, viral, or eccDNA.
I need high purity, because I want to be able to say with confidence
that any resulting clone came from extrachromosomal DNA &
not from
contaminating chromosomal fragments. (There is an ATP-dependent
DNAse
method by Yamagishi et al '83 [Gene 26:317-321], but it only gives
96%
purity.) And the high yield of the acid-phenol method is very
attractive, because my starting material is available in limited
quantity (10^9 cells max). I have heard that the yield after
3
successive CsCl runs (which is what I would need for the requisite
purity, & seems to be the standard method in the literature)
is as low
as 10%. Any feedback on whether that's true? I also have concerns
about whether there's going to be enough DNA for me to see the
band in
order to collect it by side puncture, especially since I don't
want to
UV-irradiate the hell out of my poor DNA. Advice on either of
those
points (how much DNA do you need to see in a 5ml tube [with &
without
UV], & how much UV is OK) is welcome, too.
I plan to follow either the acid-phenol or the CsCl prep with
digestion by ATP-dependent DNAse, which looks like a great method
for
cleaning up any residual chromosomal DNA. Hopefully that should
take
care of the fragments <1500bp. One of my reservations, however,
is
that this enzyme is inhibited by ssDNA, so if a significant amount
of
that ends up in the aq phase, I may be out of luck. Any feelings
on
how likely that is?
Finally, if anybody has any other ideas on how to do this, I'd
love to
hear them. Has anyone ever tried commercial plasmid prep kits
(like
Magic Mini-preps) on eukaryotic samples? The ratio of small circular
DNA to chromosomal DNA is a lot lower than in bacteria -- ~.03%
eccDNA; maybe ~0.3% once you include mitochondrial DNA. And I
suspect that the amount of sample I could process this way would
be
too low. But who knows; it might work! (If it matters, I'm planning
to isolate the DNA from whole cells, and eliminate the mitochondrial
stuff later, during the cloning step.)
Subject: Re: freedom from RNAse
Brett, one common problem that I've seen with people doing
occasional RNA work is not with the actual isolation
procedure, but with the gel rig. Has your gel rig been used
for analyzing plasmid minipreps which have been digested
with RNase A? If so, the electrodes of the rig become
covered with RNase which will come off during
electrophoresis and gobble up your RNA on its way to the
(-)electrode. To clean up a gel rig I soak it in 1% SDS, 1%
NaOH & 0.1% EDTA overnight. I then reserve the rig for use
with RNA samples only.
I hope this is your problem since it is easily fixed.
However, if the problem lies elsewhere, remember that RNase
is stable stuff. Baking glassware at 150
degrees may not be harsh enough, better try 250 degrees.
Also, I don't think treating glassware with chloroform
serves any useful purpose. I try to use disposable
plasticware as much as possible.
I hope this helps--Roy
Subject: Re: freedom from RNAse
Our standard list includes;
-All glassware, pipettes, spatulas, weighing foils, etc baked
overnight
at 250 C to destroy RNAase. Cover openings and wrap other items
with
Al foil to protect from contamination until use.
-Soak non-heatable items with 1 M NaOH overnight and then rinse
with
DEPC treated water. (Stirring bars, electrophoresis gel boxes)
-All reagents are originally new and unopened and then stored
separately and used only for RNA work.
-Separate pH electrode purchased new and used only in RNAase-free
solutions.
-Gloves worn at all times and changed whenever they come in contact
with a non-rnase-free surface (just use common sense).
-Whenever possible, use disposable plasticware for solution preps
and
storage.
-Don't let people smoke in the vicinity as there is RNase in the
smoke particles (so I've heard).
Subject: Re: Paramagnetic particles
> We recently purchased some streptavidin paramagnetic particles
> -nucleic acid qualified (cat. no. Z524/1,2) from Promega
and
> would like to know some more information concerning these
beads.
>
> 1. What are the size of the particles?
Well they vary in size. I've imaged them in the TEM and with
my Atomic Force Microscope and they seem to average around
500nm but I've seen them as big as 1.5um and as small as
100nm. BTW these "MagneSphere" particles are in no way
round! They're more like "MagneBlobs". The Dynal M-280
magnetic particles are very round and have a mean dia. of
2.8um +/- 0.2um if that's what you need.
> 2. What is the concentration of particles in the supplied
buffer?
1 mg/ml
> 3. What is the buffer that the particles are shipped in?
PBS and it may have some BSA and/or Azide for the Streptavidin
> 4. What is the binding capacity of the beads for biotinylated
> oligonucleotides?
1.08 nanomoles biotinylated oligo d(T) per mg particles
1.1% Non-specific binding of radiolabeled oligo d(T) (no biotin)
is captured after incubation with 2ml particles for 10min at RT
> Paul N. Hengen
All of this info can be found on the Certificate of Analysis
provided with the MagneBlobs. The info given here is for
lot 08540. They get their particles from Advanced Magnetics
and I think they coat them with Streptavidin themselves.
Also, check out Edmund Scientific for a good source for
rare earth magnets. If you need more help drop me a line..
Subject: Re: paramagnetic particles
The paramagnetic particles are supplied by Advanced Magnetics,
Inc.
1-800-343-1346; however, the specs from Advanced Magnetics do
not apply
for some reason. I think the beads are further diluted after Promega
gets them. Also, these beads are a different size than the Dynal
Dynabeads M-280 Streptavidin so don't be fooled into thinking
they're
all the same, ie. different size = different binding capacity.
From Promega technical support, I got the following information...
1.The beads are ~ 0.5 um mean diameter. I imagine they vary depending
on the batch of beads etc. (maybe as big as 1.5 um; The Dynal
beads
are on the order of 3.0 um).
2.They are supplied in 1 x PBS (10 x = 11.5 g/l Na2HPO4; 2.0 g/l
KH2PO4; 80.0 g/l NaCl; 2.0 g/l KCl; pH 7.4), therefore, the
1 x
buffer is 4 mM Na2HPO4; 1.5 mM KH2PO4; 140 mM NaCl; 3 mM KCl;
pH 7.4.
3.They are supplied in aliquots of 600 ul at a concentration of
1 mg/ml.
4.The binding capacity has been determined by binding of a 25mer
biotinylated oligo where 1 mg of beads binds 1 nmol oligo. Since
the beads are at 1 ug/ul (1 mg/ml), 1 ul will bind 1 pmol oligo.
5.The measured surface area is 100-150 m^2/g beads. From my estimate,
the binding capacity of a single bead at 0.5 um diameter will
be
on the order of 5 - 10 fmol oligo (please check me).
BTW, I talked to Angela Ryan of Promega (biotec@pslc.psl.wisc.edu)
and
was told that Promega DOES NOT monitor messages addressed to
promega@biotechnet.com eventhough they specifically advertise
this in
BioTechniques. The address that is monitored is BIOTEC@WISCPSL.
Subject: Genomic DNA in small quants. - A summary.
Some time (quite long..) ago, I have posted a request for
protocols on isolation of genomic DNA. I have received some
answers W/ references. I'd like to repost the ref's W/ some
detailes. Beg pardon for the great delay in reposting, but
you know- problems, problems, problems...
1: (Received W/ thanks from Eric Atkinson).
Kim and Smithies, Nucleic Acids Research, vol. 16, no. 18,
pp 8887-8903 (1988). s.a. - Joyner et al. Nature,
vol. 338, pp 153-156, (1989)).
This method is based on a hypotonic lysis step followed by
a Proteinase K step (freez/thaw cycle here is optional). The
supernatant goes directly to PCR. The method suits a small number
of cells. I have tryed it with success. However,I got some non
specific bands (I probably used too many cells).
2: This is from Jeff Ross - Thanks Jeff.
Nucleic Acids Res., 19:6053 . This technique is for DNA
isolation from small tissue samples. While the protocol calls
for spooling the DNA onto a plastic pipette tip, centrifugation
would work for collecting amounts too small to spool. They have
used this technique with samples smaller than 200 mg, including
liver, lung, peripheral blood lymphocytes, C3H10T1/2.
I haven't tryed this method yet, but intend too.
3: Frank Chiafari sent me a very detailed protocol :-)
essentially for suspended cells.
The full protocol can be obtained by request. The outlines are:
a. Transfer cells into 15ml tube. Add Lysing Solution. Mix and
Centerfuge . Pour off solution, drain leaving pellet.
b. Resuspend pellet in ACE Shocking Solution, transfer to labelled
microfuge g
tube; Spin ; Pour off solution, saving pellet; Repeat ACE wash.
c. Add 300ul of Nuclei Lysis Buffer to each pellet.
d. Add SDS, Proteinase K and RNase A , vortex ; incubate for
1hr. at 60*C; vortex again and incubate (same).
e. Add Precipitating Solution. Shake . Let stand at room temp
for
3 min. Spin in microfuge.
f. Transfer supernatant containing DNA to new tube; add EtOH.
Mix until DNA comes out of solution. Spin tube ; Wash pellet with
EtOH and respin; rotovac untill barely dry; add TE & Incubate
at
60*C from two hrs. to o/n.
Frank writes that the DNA is easily restriction cut and ligated.
DNA isolated
from blood to be used with PCR will amplify, but will do better
if chelexed
first.
4. I myself, also found a convinient way of isolation W/ a kit
(yes friends, I know, I have read the writings on the walls
regarding your opinion on the use of kits). The kit is "Gnome"
from BIO-101. I downscaled the whole procedure 1:10, and I spin
the DNA down instead of spooling. It works O.K. in my hands.
Regarding the use of Chelex- Many people have suggested it's use
in purifying genomic DNA for PCR. Chelex 100 resin is claimed
to
enhance PCR's specificity and efficiency by at least 10 times.
This is produced by Bio-Rad, Cat. # 143-2832 Biotech. Grade.
Regretfully, I haven't tryed it out in my hands yet, :-(
since the stuff has to be imported to Israel which takes a month
(Not to mention about 50% added to the price).
Life's not that simple as you can see. We in Israel, have added
another law to Murfey's : "Whenever you'll need a material
or
a piece of equipment BADLY and URGENTLY, it will not be in stock
- and it will take it months to get there..."