Methods in Biotechnology

Purification of Plasmid DNA

noreply@blogger.com (Unknown) — Thu, 11 Feb 2010 04:29:00 +0000

After the initial characterization, it is possible to purify further some or all of the plasmid DNAs by RNase digestion and extraction with organic solvents. This further purified DNA is suitable for techniques such as DNA sequencing, subcloning or the production of gene probes. In order to purify plasmid DNA after the isolation process, any residual RNA and contaminating protein are removed. This purification step involves two main steps, which are, first, removing residual RNA by using RNase in order to digest RNA and, second, extract contaminating protein using organic solvents, phenol-chloroform.

Materials:

RNase A: Make up as a solution in water at 10 mg/mL, Heat for 10 min in a boiling water bath or heating block to eliminate any DNase activity. Aliquot and store at -20^oC.
0.4 M Ammonium acetate.
Chloroform= A 24: 1 mix of chloroform and isoamyl alcohol. Store at 4^oC.
Phenol/chloroform= 25:24: 1 mix of TE-equilibrated phenol, chloroform, and isoamyl alcohol. Store at 4^oC.
100% Ethanol.
Sterile wooden toothpicks.

Methods:

Add 50 microliters of 4 M ammonium acetate containing 200 micrograms/mL RNase A to each miniprep and incubate it at room temperature for 20 min.
Add 100 microliters of phenol/chloroform to each DNA preparation.
Vortex briefly and centrifuge at high speed for 2 min in a microfuge. Remove the top layer containing the DNA and place it in a new sterile tube.
Add 100 microliters of chloroform to each tube.
Vortex briefly and centrifuge at high speed in a microfuge for 2 min. Remove the DNA in the top layer and place it in a second sterile tube.

For phenol/chloroform extractions avoid removing material from the interface.
Add 200 microliters of 100% ethanol to each tube.
Shake briefly to precipitate the DNA and centrifuge at high speed for 5 min at room temperature.

Done. Now you can extract, isolate, and purify Plasmid DNA using methods which I had described in this blog. Hopefully those can be useful for you.

Reference: Number 18 on References

Estimation of Disulfide Bonds Using Ellman’s Reagent

noreply@blogger.com (Unknown) — Thu, 08 Oct 2009 07:17:00 +0000

Ellman’s reagent has been widely used for the quantitation of thiols in peptides and proteins. It has also been used to assay disulfides present after blocking any free thiols (e.g., by carboxymethylation) and reducing the disulfides prior to reaction with the reagent. It is also commonly used to check the efficiency of conjugation of sulfhydryl- containing peptides to carrier proteins in the production of antibodies.

Ellman’s reagent or DNTB (5,5'-dithiobis(2-nitrobenzoic acid)) was first introduced in 1959 for the estimation of free thiol groups. The procedure is based on the reaction of the thiol with DTNB to give the mixed disulfide and 2-nitro-5-thiobenzoic acid (TNB) which is quantified by the absorbance of the anion (TNB 2-) at 412 nm.

Materials :

Reaction buffer: 0.1M phosphate buffer, pH 8.0.
Denaturing buffer: 6M guanidinium chloride, 0.1M Na2HPO4, pH 8.0.

It is not recommended to use urea in place of guanidinium HC1, since this can readily degrade to form cyanates, which will react with thiol groups
Ellman's solution: 10 mM (4 mg/mL) DTNB in 0.1M phosphate buffer, pH 8.0.

Unless newly purchased, it is usually recommended to recrystallize DTNB from aqueous ethanol
Dithiothreitol (DTT) solution: 200 mM in distilled water.

Methods :

A. Analysis of Free Thiols

It may be necessary to expose thiol groups, which may be buried in the interior of the protein. The sample may therefore be dissolved in reaction buffer or denaturing buffer. A solution of known concentration should be prepared with a reference mixture without protein. Sufficient protein should be used to ensure at least one thiol per protein molecule can be detected; in practice, at least 2 nmol of protein (in 100 micro-liter) are usually required.
Sample and reference cuvets containing 3 mL of the reaction buffer or denaturing buffer should be prepared and should be read at 412 nm. The absorbance should be adjusted to zero (A_buffer).
Add 100 micro-liter of buffer to the reference cuvet.
Add 100 micro-liter of Ellman's solution to the sample cuvet. Record the absorbance (A_DTNB).
Add 100 micro-liter of protein solution to the reference cuvet.
Finally, add 100 micro-liter protein solution to the sample cuvet, and after thorough mixing, record the absorbance until there is no further increase. This may take a few minutes. Record the final reading (A_final).
The concentration of thiols present may be calculated from the molar absorbance of the TNB anion.
U-A₄₁₂ = E₄₁₂TNB² - [RSH] (1)
Where U-A₄₁₂ = A_final- (3.1/3.2) (A_DTNB – A_buffer)
and E₄₁₂TNB²- = 1.415 x 10⁴ cm^-1 M^-1.
If using denaturing buffer, use the value E₄₁₂TNB²- = 1.37 x 10⁴ cm^-1 M^-1.

Standard protocols for use of Ellman's reagent often give E₄₁₂TNB²- = 1.37 x 10⁴ cm^-1 M^-1.

B. Analysis of Disulfide Thiols

Sample should be carboxymethylated or pyridethylated without prior reduction. This will derivatize any free thiols in the sample, but will leave intact any disulfide bonds.
The sample (at least 2 nmol of protein in 100 micro-liter is usually required) should be dissolved in 6M guanidinium HC1, 0.1M Tris-HC1 pH 8.0 or denaturing buffer, under a nitrogen atmosphere.
Add freshly prepared DTT solution to give a final concentration of 10-100 mM. Carry out reduction for 1-2 h at room temperature.
Remove sample from excess DTT by dialysis for a few hours each time, with two changes of a few hundred mL of the reaction buffer or denaturing buffer. Alternatively, gel filtration into the same buffer may be carried out.

Regards.

Reference: Number 21 at References

Separation of DNA Fragments Using PAGE Method

noreply@blogger.com (Unknown) — Thu, 24 Sep 2009 07:06:00 +0000

This method is able to separate DNA fragments with the size of as small as 10 bp and up to 1 kb with the resolution of as little as 1 bp. While agarose gel electrophoresis is only able to separate DNA fragments with the bigger size that PAGE does or in the size range of 100 nucleotides to around 10 – 15 kb.

Materials:

Gel apparatus: Many designs of apparatus are commercially available. The gel is poured between two vertical plates held apart by spacers

The plates should be cleaned thoroughly and then wiped with ethanol. To help ensure that the gel only sticks to one plate when the apparatus is disassembled, apply silicon to one of the gel plates. This is easily done by wiping the plate with a tissue soaked in dimethyl dichlorosilane solution and then washing the plate in distilled water followed by ethanol. If the plates are baked at 100^oC for 30 min, the siliconization will last four to five gel runs.
Deionized H₂O: Autoclaved water is not necessary for the gel mix or running buffer, but it should be used for diluting samples and purification from gel slices.
l0x TBE: 108 g of Trizma base (Tris), 55 g of boric acid, and 9.3 g of ethylenediaminetetraacetic acid (EDTA) (disodium salt). Make up to I L solution with deionized H₂O, which should be discarded when a precipitate forms.
Acrylamide stock: 30% acrylamide, 1% N,N'-methylene bisacrylamide. Store at 4^oC. This is available commercially, or it can be made by dissolving acrylamide and bisacrylamide in water, which should be filtered. Acrylamide is a neurotoxin and therefore must be handled carefully. Gloves and a mask must be worn when weighing out.
APS: 10% Ammonium persulphate (w/v). This can be stored at 4^oC for 1-2 months.
TEMED: N,N,N',N'-tetramethyl-l,2-diaminoethane. Store at 4^oC.
5X sample buffer: 15% Ficoll solution, 2.5X TBE, 0.25% (w/v) xylene cyanol and 0.025% (w/v) bromophenol blue.
Ethidium bromide: A l0-mg/mL solution. Ethidium bromide is a potent mutagen and should be handled with care. Store at 4^oC in the dark.

Methods:

For 50 mL, enough for a 18 x 14 x 0.15 cm gel, mix l0x TBE, acrylamide, H₂O, and APS as described in Table below.

Just prior to pouring, add 50 microliters of TEMED and mix by swirling.
Immediately pour the gel mix between the gel plates and insert the gel comb. Leave to set; this takes about 30 min.
Fill the gel apparatus with 0.5X TBE and remove the comb. Use a syringe to wash out the wells, this may take multiple washes. It is important to remove as much unpolymerized acrylamide as possible because this impairs the running in of the samples

If you are separating very small fragments, e.g., less than 50 bp, the gel should be prerun for 30 min, as this elevates the resolution problem experienced with fragments running close to the electrophoresis front..
Add 0.2 volume of 5x sample buffer to each sample, usually in 10-20 microliters of TE, water, or enzyme buffer. Mix and spin the contents to the bottom of the tube

High-salt buffers (above 50 mM NaCl) will affect sample mobility and tend to make bands collapse. In this case, salt should be removed by ethanol precipitation..
Load the samples on the gel and run at 200-300 V (approximately 10 V/cm) until the bromophenol blue band is two-thirds of the way down the gel; this takes about 2.5 h

Do not run the gel faster than 10 V/cm, as this will cause the gel to overheat, affecting the resolution. The gel can be run more slowly, e.g., 75 V will run overnight.
Disassemble the gel apparatus and place the gel to stain in I mg/mL of ethidium bromide for approximately 30 min. View the stained gel on a transilluminator.

Happy separating DNA fragments.

Reference: Number 20 on References

Polyacrylamide Gel Electrophoresis: Advantages and Disadvantages

noreply@blogger.com (Unknown) — Mon, 21 Sep 2009 03:17:00 +0000

You must have known that agarose gel electrophoresis is generally adequate for resolving nucleic acid fragments in the size range of 100 nucleotides to around 10-15 kb. But, for nucleic acid which its fragments below those range, it will be difficult to separate and hard to visualize because of diffusion within the gel matrix. These problems are solved by native polyacrylamide gel electrophoresis (PAGE). Using native PAGE, fragments as small as 10 bp and up to 1 kb can be separated with a resolution of as little as 1 bp.

Polyacrylamide Gel Electrophoresis has a number of advantages, which are:

PAGE has a high loading capacity, up to 10 micrograms of DNA can be loaded into a single well (1 cm x 1 mm) without significant loss of resolution.
Polyacrylamide contains few inhibitors of enzymatic reactions.
PAGE is an ideal gel system from which to isolate DNA fragments for subcloning and other molecular biological techniques.

As any other methods, PAGE also has disadvantages:

The mobility of the fragments can be affected by base composition making accurate sizing of bands a problem.
Polyacrylamide quenches fluorescence, making bands containing less than 25 ng difficult to visualize with ethidium bromide staining.

As an alternative method for visualizing DNA fragments instead of using Ethidium Bromide staining is carried out by wrapping the gel in a UV-transparent plastic film, such as Saran Wrap, and then placing it onto a thin-layer chromatography plate that contains a UV fluorescent indicator. Long-wave UV light is shone through the gel onto the chromatography plate, causing it to glow. Regions of high DNA concentration leave a "shadow" on the plate as the transmitted UV is absorbed by the DNA. The position of the DNA can be marked on the Saran Wrap with a fiber-tip pen and then cut from the gel.

The methods to separate and to purify DNA fragments will be described later in the next post.

Reference: Number 20 on References

Rapid Boiling Method for Plasmid DNA Isolation

noreply@blogger.com (Unknown) — Sat, 29 Aug 2009 17:16:00 +0000

In the previous post, it had already explained about how to extract plasmid DNA by using alkaline lysis method. In this particular post, it will be explained the other method to extract plasmid DNA instead of using alkaline lysis method. It is Rapid Boiling method, an alternative to alkaline lysis method that was developed by Holmes and Quigley. Here, the cells are lysed partially allowing plasmids to escape, whereas the bacterial chromosomal DNA remains trapped in the cell debris. High temperature is then used to denature the chromosomal DNA, after which reannealing allows the plasmids to reassociate. Centrifugation removes the chromosomal DNA along with the cell debris, leaving the plasmid in suspension, from where it is recovered by isopropanol precipitation.

Materials:

LB broth bacteria culture medium. The content are 1% Tryptone, 0.5% yeast extract, 200 mM NaCl, then it sterilize by autoclaving in suitable aliquots.
STET mix which is contain 5% (v/v) Triton X-I 00, 50 mM Tris-HCl, pH 8.0, 50 mM EDTA, pH 8.0, 8% (w/v) sucrose. Store the mix solution at room temperature.
Lysozyme: Dry powder. Store at -20^oC.
70% Ethanol.
Isopropanol.
TE solution: 10 mM Tris-HCl pH 8.0,1 mM EDTA.
A boiling water bath: An opened bottom tube rack is required because the tubes must be placed directly in the water to achieve rapid heating.
Sterile wooden toothpicks.

Methods:

Set up a culture for each miniprep by inoculating 2-3 mL of L-broth, containing an appropriate antibiotic (e.g., 100 micrograms/mL ampicillin) with a bacterial colony. Grow overnight at 37^oC with vigorous shaking.

Where plasmids have a high copy number, the growth time may be reduced to approx 6 h.
Before starting the miniprep, begin boiling the water and make up a fresh solution of 1 mg/mL lysozyme in STET mix.
Fill a 1.5-mL labeled microfuge tube with an aliquot from each culture. Pellet the bacteria by centrifugation for 1 min at 12,000 g. Carefully aspirate off the supernatant using a drawn-out Pasteur pipet.

The short centrifugation time leaves a loose pellet that is easier to resuspend. If the pellet does not readily resuspend, pipet the solution up and down to dislodge it. Do not suck the pellet directly into the pipet tip.
Vortex each pellet for a few seconds to break up the pellet. Add 20 micrliters STET mix to each tube. The pellet should now easily resuspend by vortexing.
Immediately place the tubes in the open-bottom rack, and place in the boiling water for exactly 45 s. Ensure that each tube is at least half submerged.
Centrifuge the tubes at 12,000 g for 10 min. A large, sticky, loose pellet should form.
Remove the pellet from each tube by "fishing" it out with a sterile wooden toothpick. Because the pellet is quite slippery, it is useful to have a paper tissue at the top of the tube to catch the pellet and prevent it from slipping back down into the tube.
Add 200 microliters isopropanol to each tube, and centrifuge at 12,000 g for 5 min.
Aspirate the supernatant, and wash the pellet in 500 microliters 70% ethanol. Centrifuge the tube for 1 min to compact the pellet, and then aspirate the 70% ethanol.
Air dry the pellets for 10 min, and resuspend each one in 100 microliters TE buffer. Vortex and shake for 10 min before use to ensure complete dissolution.
Use 10 microliters (equivalent to 100 ng of plasmid for most vectors) and analyze by gel electrophoresis.

It is possible to scale up procedures for the isolation of plasmid.
Now, you have two options method in extracting plasmid DNA, which are alkaline lysis method and Rapid Boiling method.

Reference: Number 18 on References

Plasmid DNA Extraction Using Alkaline Lysis Method

noreply@blogger.com (Unknown) — Wed, 26 Aug 2009 05:24:00 +0000

Plasmids can be isolated by a variety of methods many of which rely on the differential denaturation and reannealing of plasmid DNA compared to chromosomal DNA. One commonly used technique developed by Birnboim and Doly involves alkaline lysis. This method essentially relies on bacterial lysis by sodium hydroxide and sodium dodecyl sulfate (SDS), followed by neutralization with a high concentration of low-pH potassium acetate. This gives selective precipitation of the bacterial chromosomal DNA and other highmolecular-weight cellular components. The plasmid DNA remains in suspension and is precipitated with isopropanol.

Materials:

Luria Bertani (LB) broth bacteria culture medium: 1% Tryptone, 0.5% yeast extract, 200 mM NaCl. Sterilize by autoclaving in suitable aliquots. In order to ensure retention of the plasmid, media should be supplemented with the appropriate antibiotic(s).
1.5 mL Microfuge tubes.
Sterile tubes: Must have a volume of at least 10 mL to ensure good aeration.
Lysis solution: 200 mM NaOH, 1% SDS. Store at room temperature.
Resuspension solution: 50 mM glucose, 50 mM Tris-HCl, pH 8.0, 10 mM ethylene diamine tetraacetic acid (EDTA). Keep at 4^oC to prevent growth of contaminants.
Potassium acetate (neutralizing solution): 3 M potassium/5 M acetate. For 100 mL, take 29.4 g of potassium acetate, add water to 88.5 mL, and 11.5 mL of glacial acetic acid. Store at room temperature.
TE: 10 mM Tris-HCl, pH 8.0, 1 mM EDTA.
Isopropanol.
70% Ethanol.

Methods:

Take a number of separate sterile tubes and place 2 mL of L-broth into them. Inoculate from individual bacterial 37°C overnight with shaking.
Transfer each culture to a labeled 1.5-mL Eppendorf tube, and centrifuge for 30 s at high speed in the microfuge.

Here, a tight creamy pellet may be seen.
Decant the supernatant and place tubes in a rack vertically for 5-10 s. Remove any of the remaining liquid by aspiration with a fine Pasteur pipet.
Add 100 microliters of resuspension solution into each tube, close the lids, and resuspend the bacteria in each tube by shaking or vortexing to dissociate the bacterial pellet.
To each tube add 200 microliters of lysis solution and mix by inverting the tube several times.

The solution should quickly turn transparent and become more viscous indicating bacterial
lysis has taken place.
Allow at least 2-3 min for lysis to take place and leave the tubes to stand for 60 s before opening. This will allow the liquid to return to the bottom of the tube.
To each tube add 150 microliters of neutralizing solution and invert the tubes several times. At this point bacterial chromosomal DNA is usually seen as a white precipitate.
Centrifuge the tubes for 2-5 min at full speed in a microfuge.
Place new sterile tubes into a rack, label them, and add 250 ~ of isopropanol to each tube.
Remove the tubes from the microfuge, being careful not to disturb the precipitate.
Remove the supernatant with a 1-mL pipet, avoiding the white precipitate as much as possible.

Some of the precipitate may float, so it is critical to use a pipet and disposable tips to
recover the supernatant rather than pouring it.
Transfer the liquid phase into the new set of labeled tubes containing the isopropanol.
Vortex the tubes for 5-10 s and centrifuge the tubes in the microfuge for 30 s at high speed. The plasmid DNA precipitates as a white pellet.
Decant the supernatant and wash the pellets by adding 750 mL of 70% ethanol, vortex briefly, and centrifuge at high speed for 30 s.
Decant the ethanol, and centrifuge again for 10 s to collect the remaining ethanol at the bottom of the tubes. Carefully aspirate the remaining ethanol and leave the tubes to air dry on the bench for 5 min.
Dispense 50 microliter of TE into each tube, and resuspend the pellet

It is not advisable to vortex, as this may lead to DNA shearing. The sample is best left for
3-5 min with occasional finger flicking of the tube.
Take 10 microliters of the resuspended pellet and analyze by agarose gel electrophoresis (see on my posting entitled Agarose Gel Electrophoresis of Nucleic Acids).

Actually, you can use another method to extract plasmid DNA instead of Alkaline Lysis. I will try to describe it on the next post. Hopefully, this method will be useful for your lab work.

Reference: Number 18 on References

96-Well DNA Isolation Method From Leaf

noreply@blogger.com (Unknown) — Mon, 29 Jun 2009 08:25:00 +0000

The following protocol provides a high throughput, low cost method of producing a superior DNA yield of high quality which is suitable for TILLING, map based cloning or any application which requires long term DNA storage. The protocol has been designed for DNA extraction from leaf material, preferably young leaf tissue should be used as this minimizes samples being contaminated with polysaccharides and phenolics.

Materials:

Extraction Buffer

200 mM Tris-HCL pH 7.5
250 mM NaCl
25 mM EDTA
0.5% SDS

10 mM Tris-HCl pH 8.0
0.1 mM EDTA pH 8.0

TE and RNAse A

For 2 plates: 20 ml TE and 20μl DNAse-free RNase A (10 mg/ml)

The Methods are:

Pre-heat extraction buffer to 65°C.
Label collection tubes (Qiagen Cat. No. 19560) and add a single tungsten carbide bead (Qiagen Cat no. 69997) to each tube.
Prepare ice bucket and tube of ethanol to wash forceps after each harvest.
Harvest material (3 growing tips) into a single collection tube.
Add 400 microliters of extraction buffer to each tube (use a multi-pipettor). Put on lids (Qiagen Cat. No. 19566).
Homogenise material on the mixer mill (Retsch MM300) for 2 min/30s.
Incubate at 65^oC for 30 min to 1 hour.
Centrifuge for 10 minutes at full speed.
Label a new rack of collection tubes.
Remove 300 microliters of supernatant into a new collection tube (use a multi-pipettor set speed to slow) with extended length tips.
Carefully add 200 microliters of phenol:chloroform to each tube (THIS PROCEEDURE SHOULD BE CARRIED OUT IN THE FUME HOOD), use 200 microliters manual multi-pipettor with filter tips
Put lids onto the tubes and invert several times, so the samples are well mixed. Centrifuge for 10-15 minutes.
Label a set of storage plates.
Using a manual multi-pipettor with filter tips very carefully remove 200 microliters of the upper layer to a new storage plate (AB Gene Cat. No. AB 0765) (THIS PROCEEDURE SHOULD BE CARRIED OUT IN THE FUME HOOD).
Using a multi-pipettor add 1/10^th vol. of 3 M sodium acetate ( approximately 20 microliters) and an equal volume of isopropanol (approximately 220 microliters). Put on lids, mix well, and leave at -20^oC for a maximum of 1 hour.
Leave plates on desk until the have reached room temperature, as spinning when still frozen at high speeds can cause the plate to crack. Centrifuge at 5600 rpm for 45 minutes.
Remove supernatant and add 100 microliters of TE containing RNAse A at a final concentration of 10 microliters/ml and incubate 30 min at 37^oC.
Repeat the precipitation step (step 15). After centrifugation at 5600 rpm for 45 minutes remove the supernatant and add 200 microliters 70% ethanol. Put on lids and leave for 15 minutes or overnight.
Remove ethanol and leave to air dry. Add 100 microliters of TE and store in fridge/freezer.

Kindly Regards.

Reference: Number 17 in References

RNA Extraction From Fresh Blood

noreply@blogger.com (Unknown) — Mon, 08 Jun 2009 07:40:00 +0000

RNA can be extracted from blood since whole blood contains nucleated white cells that constitute an easily accessible source. RNA extraction from blood will be more successful if the nucleated white cells are first isolated from the red cells since the red cells are a rich source of ribonucleases that are able to degrade RNA. It is important to minimize degradation by following the appropriate recommendations for handling RNA. The methods of RNA extraction usually comprises of three steps which are cell lysis, partitioning of RNA into a solvent fraction, and recovery of RNA from the solvent by precipitation.

Here are the materials that you need:

X Phosphate-buffered saline (PBS).
Lymphoprep (Nycomed).
10-mL centrifuge tubes (polypropylene or glass).
RNAzol B (Note: RNAzol B contains guanidinium thiocyanate which is an irritant, and phenol which is a poison. It is therefore recommended that RNAzol B is handled in a fume cupboard.).
Chloroform.
Isopropanol.
Ethanol.
5 M sodium chloride.

And The Method is:

Dilute 10 mL of anti coagulated whole blood 1:2 with 1X PBS in a sterile plastic 20 mL universal.

A suitable antrcoagulant is 3.8% trisodmm citrate diluted 1: 10 mto whole blood.
Carefully layer 10 mL of the diluted blood onto 3 mL of lymphoprep in each of 2 x 10 mL polypropylene or glass tubes able to withstand centrifugation at 800g. Ensure that a sharp interface is obtained with little or no mixing between the blood and separation fluid.
Centrifuge at 400g for 30-40 mm or 800 g for 15 min at room temperature.

g-Force can be converted into rpm using the formula g = 0.0000118 x r x N², where r = radius in cm and N = rpm.
Following centrifugation, a clear solution IS obtained with aggregated erythrocytes sedimented to the bottom of the tube. Mononuclear cells, including lymphocytes form a distinct, cloudy band within the clear solution at the interphase of the upper sample plasma layer and the lower Lymphoprep solution (see Fig. 1) Transfer the mononuclear cell layer to a separate tube using a pipet tip or Pasteur pipet. The upper layer may first be removed to just above the band, if desired.
Make the cell solution up to 5 mL with 1X PBS, invert to mix and centrifuge as in step 3.
Decant the supernatant. The lymphocyte pellet may be stored for several days in this condition at -20^oC before subsequent processing if desired.
Lyse the cells by the addition of 0.5 mL RNAzol B Solubilize the RNA by passing the lysate through the pipet a few times.
Transfer the lysate to a sterile Eppendorf, add 50 microliter of chloroform, shake samples vigorously for 15 s, and incubate on ice (or at 4^oC) for 5 min. (Samples can be stored in this state for 1-2 h).
Centrifuge the suspension at 12,000 g for 15 min in a microfuge.

Ideally, microfuge spins from this stage onwards should be carried out at 4^oC but may be performed at room temperature if a refrigerated micromge is not available.
Transfer the upper aqueous phase (carefully avoiding the interphase, which contains DNA and proteins) to a fresh Eppendorf, add an equal volume of isopropanol (approx 400 microliter), and store for 15 min at 4^oC (or at -20^oC overnight).
Microfuge samples at 12,000 g for 15 min. The RNA pellet should be visible at the bottom of the tube.
Remove the supernatant and wash the RNA pellet once by adding 800 microliter of 75% ethanol, vortex and centrifugation at 7500 g for 8 min.
Resolubilize the RNA in 0.2M sodium chloride (e.g., 192 microliter water + 8 microliter 5 M sodium chloride). Precipitate sample with 400 microliter of 100% ethanol at -20^oC for 1 h.
Centrifuge and ethanol wash as above (steps 12 and 13).
Allow the pellet to dry with tube open at room temperature for 15 min
Solubilize the pellet in 50 – 100 microliter of DEPC-treated water. The sample may be heated to 52-60^oC for 5-15 min to aid solubilization.
Quantitate and analyze the RNA.

Regards.

Reference: Number 16 on References

DNA Extraction From Fresh Bone

noreply@blogger.com (Unknown) — Mon, 01 Jun 2009 15:23:00 +0000

You can extract nucleic acids, such as DNA, from bone samples in order to analyze gene expressions, to look for somatic mutations of tumors or other pathological tissue, or for genotyping archive material when other sources of DNA are not available. You can use several kits that have already provided by biotech companies. But, if you are extracting DNA from large number samples, you can use a homemade method as described here to be effective in cost.

There are four procedures that ascertain the successful extraction of nucleic acids from tissue:

disrupting the tissue so that extraction reagents can reach the cells.
disrupting the cell membranes so that nucleic acids are liberated.
separation of the nucleic acid from other cellular components.
precipitation and solubilization of the nucleic acid.

Materials

DNA extraction buffer: Add 17.6 mL of 0.75 M sodium citrate, pH 7.0, 26.4 mL of 10% sodium lauryl sarkosyl, and 250 g of guanidinium isothiocyanate to 293 mL of distilled water and mix well. Add 7.2 microliter of beta-mercaptoethanol/mL of lysis buffer on the day of use

All chemicals should be of molecular biology grade. The solutions can be stored at 4^oC for up to 3 months.
0.5 M ETDA: Add 93.05 g of EDTA to 300 mL of distilled water and add 10 N NaOH to pH 8.0. Make up to 500 mL. Autoclave.

Tris–EDTA: Add 1 mL of 1 M Tris to 200 microliters of 0.5 M EDTA. Make up to 100 mL with distilled water.
3 M Sodium acetate, pH 5.2: Add 401.8 g of sodium acetate to 800 mL of distilled water. Adjust pH to 5.1 with glacial acetic acid. Make up to 1 L with distilled water. Autoclave.
General Reagents: Tris-saturated phenol pH 7.8–8.0 (Sigma), Chloroform, Isopropanol 100%, Ethanol.

Methods

Collect the bone sample in a sterile container containing phosphate-buffered saline (PBS) and transport to the laboratory within 1–2 h.

If the DNA extraction is not initiated immediately, freeze the sample at –20^oC or below for later use.
Place the bone tissue in a clean glass Petri dish. Using bone cutters or a strong sharp pair of scissors, isolate a piece of bone measuring about 1 cm³ and transfer to a clean 5-mL bijoux container.
Add 1 mL of DNA extraction buffer and homogenize the tissue with the scissors until a slurry solution is obtained.
Transfer 500 microliters aliquots of slurry into screw-capped conical-bottomed 1.5-mL Eppendorf tubes.
Add one volume of Tris-saturated phenol, followed by one volume of chloroform per tube. Mix well by inverting the tubes a few times or by shaking. Do not vortex, because vortex-mixing causes long strands of DNA to shear.
Centrifuge the tubes at 10,000 g for 20 min to separate the phases.
Transfer the upper layer to a fresh centrifuge tube (taking note of the volume), being careful not to disturb the milky layer at the interface. Repeat steps 5–7 if the interface is disturbed.
Add one volume of ice-cold isopropanol and 0.1 volumes of 3 M sodium acetate to the supernatant. Mix well and allow to stand for 15 min on ice.
Centrifuge the tubes at 10,000g for 20 min to pellet the DNA.

Orientate the Eppendorf tube so that you can identify where the DNA pellet lies. A pellet should be visible the bottom of the tube.
Aspirate and discard the supernatant, taking care not to disturb the pellet. Wash the sample with 1.75 mL of ice-cold ethanol and centrifuge at 10,000g for 5 min. Aspirate and discard the supernatant and then repeat the wash.
Dissolve the DNA pellets in 10–50 microliters of water or Tris–EDTA buffer (you can pool DNA from the same sample at this stage) and quantitate by spectrophotometry or with Hoechst 33258.

Hoechst 33258 is a DNA-specific dye that can be used to quantitate DNA.
Store the sample frozen at –20^oC or below.

Kindly regards.

Reference: Number 15 on References

Cleave DNA Using Restriction Endonulease, A Bacterial Enzyme

noreply@blogger.com (Unknown) — Fri, 22 May 2009 08:17:00 +0000

In order to manipulate DNA you have to posses the ability to cleave DNA at specific sites by using bacterial enzyme, which is restriction endonulease. Restriction endonucleases are bacterial enzymes that cleave duplex DNA at specific target sequences with the production of defined fragments. The name of the enzyme (such as BamHl, EcoRl, AluI, and so on) tells us about the origin of the enzyme but does not give us any information about the specificity of cleavage. The recognition site for most of the commonly used enzymes is a short palindromic sequence, usually either 4, 5, or 6 bp in length, such as AGCT (for AZul),GAATTC (for EcoRl), and so on. Each enzyme cuts the palindrome at a particular site, and two different enzymes may have the same recognition sequence but cleave the DNA at different points within that sequence.

Materials

10X stock of the appropriate restriction enzyme buffer.
DNA to be digested in either water or TE (10 mM Tris-HCl pH 8.3, 1 mM ethylenediaminetetraacetic acid [EDTA]).
Bovine serum albumin (BSA) at a concentration of 1 mg/mL.

BSA is routinely included in restriction digests to stabilize low protein concentrations and to protect against factors that cause denaturation.
Sterile distilled water.

Good-quality sterile distilled water should be used in restriction digests. Water should be free of ions and organic compounds, and must be detergent free.
The correct enzyme for the digest.
5X Loading buffer: 50% (v/v) glycerol, 100 mM Na2EDTA, pH 8, 0.125% (w/v) bromophenol blue, 0.125% (w/v) xylene cyanol.
100 mM Spermidine.

Digests of genomic DNA are dramatically improved by the inclusion of spermidine in the digest mixture to a final concentration of 1 mM since the polycationic spermidine binds negatively charged contaminants.
Spermidine can cause precipitation of DNA at low temperatures, so it should not be added while the reaction is kept on ice.

Methods

Thaw all solutions, with the exception of the enzyme, and then place on ice.
Decide on a final volume for the digest, usually between 10 and 50 microliters, and then into a sterile Eppendorf tube, add 1/10 vol of reaction buffer, 1/10 vol BSA, between 0.5 and 1 micrograms of the DNA to be digested, and sterile distilled water to the final volume.

The amount of DNA to be digested depends on subsequent steps. A reasonable amount for a plasmid digestion to confirm the presence of an insertion would be 500 ng-l microgram, depending on the size of the insert. The smaller the insert, the more DNA should be digested to enable visualization of the insert after agarose gel analysis.
Take the restriction enzyme stock directly from the -20^oC freezer, and remove the desired units of enzyme with a clean sterile pipet tip. Immediately add the enzyme to the reaction and mix.

Stock restriction enzymes are very heat labile and so should be removed from -20^oC storage for as short a time as possible and placed on ice.
Incubate the tube at the correct temperature (see Note 12) for approx 1 h. Genomic DNA can be digested overnight.

Note that the incubation temperature for the vast majority of restriction endonucleases is 37^oC but that this is not true for all enzymes.
An aliquot of the reaction (usually 1-2microliter) may be mixed with a 5X concentrated loading buffer and analyzed by gel electrophoresis.

Hopefully this method can help your work.

Regards.

Reference : Number 14 on References

Agarose Gel Electrophoresis of Nucleic Acids

noreply@blogger.com (Unknown) — Mon, 11 May 2009 17:39:00 +0000

After amplify DNA template using PCR method, now you can continue your work by using gel electrophoresis in a gel composed of agarose in order to separate DNA fragments based on its molecular weight. The percentage of agarose used depends on the size of fragments to be resolved. In general a 0.8-1% gel may be used for effective separation of DNA fragments of 100-1500 base pairs.

Here are materials that you need:

Agarose (molecular biology grade).
Running buffer. There are two common types of buffer systems used in agarose gel electrophoresis, Tris-borate EDTA (TBE) and Tris-acetate EDTA (TAE).

To prepare stock solutions:

10x TBE buffer: 545 g Tris, 278 g boric acid, 46.5 g EDTA in 5 L of sterile distilled water.

50x TAE buffer: 242 g Tris, 57.1 mL glacial acetic acid, 100 mL 0.5M EDTA, pH 8.0, in 1 L of sterile distilled water.

TAE gel buffer systems are preferred to TBE systems when post-separation methods such as extraction with solid matrices are to be used. However, there is little difference between the two systems for general-purpose separation of DNA.

Sterile distilled water.
Microwave oven and boiling water bath or steamer.
Loading buffer 50% (v/v) glycerol, 50 ruM EDTA, pH 8.0, 0.125% (w/v) bromophenol blue, 1.125% (w/v) xylene cyanol. Loading buffer is at 6X concentration.
Molecular weight size marker: Numerous commercial DNA size markers are available, either as base pair ladders (e.g., 123 bp multimerladders) or predigested DNA (e.g., LambdaHindIII marker).
Ethidium bromide: 10 mg/mL dissolved in H20. Store at 4^oC in a container wrapped in tin foil.
UV transilluminator (300 nm).
Polaroid camera or gel documentation system.

And here is the method:

The precise instructions for producing a gel depends on the gel-forming apparatus used. However, it is essential to make sure the gel casting stand is leak proof and sealed correctly (e.g., with end plates or waterproof tape).
For a 1% agarose gel in TBE running buffer: Weigh an appropriate amount of powdered agarose into a conical flask. As an example, for a 1% agarose gel, add 1g of agarose and 10 mL 10X TBE running buffer and distilled water to the final volume of 100 mL, mix thoroughly by swirling.
Heat the gel mix in a microwave oven (650 W) at full power for 1 min or less.
Remove the gel mix and allow to cool to approximately 50^oC or until just cool enough to hold.
Pour the gel solution into a gel-forming tray, insert the comb template, and allow to set. This usually takes approx 25 min.
Remove the comb and tape or end plates and add sufficient 1X running buffer to the tank to cover the gel and electrodes.
To 5-10 mL of the DNA sample add 0.2 vol of loading buffer. Carefully add the sample to the well in one smooth pipetting motion.

If the samples tend to float out of the well add an additional 1-2 vol gel loading buffer and reload the sample.

Add an appropriate marker sample such as a Lambda-HindIII digest or a 123 bp ladder to the end wells of the gel.
Carry out the electrophoresis at 100 V for approx 2 hours or until the bromophenol blue dye has travelled two-thirds of the way down the gel.
Dismantle the gel apparatus and carefully place the gel in a tray. Add 100 mL of sterile distilled water containing 5 microliters of 10 mg/mL ethidium bromide (0.5 mg/mL) and allow to stain for 15 min.
Destain the gel for 5 min by replacing the solution with fresh water.

Discard the solution containing ethidium bromide in accordance with appropriate health and safety regulations. Remember to wear gloves when handling any solution containing ethidium bromide, which is a potent mutagen and potential carcinogen.

Following electrophoresis remove the gel from the electrophoresis apparatus and view on a UV transilluminator.
Photograph the gel using a polaroid camera or make a record using a gel documentation system.

Regards and happy electrophoresis.

Reference: Number 13 on References

Bacterial DNA Extraction for Pulsed-field Gel Electrophoresis

noreply@blogger.com (Unknown) — Wed, 06 May 2009 21:30:00 +0000

Pulsed-field gel electrophoresis (PFGE) is a method used to separate large DNA fragments such as those obtained after digestion with restriction endonucleases that cut infrequently. To avoid possible risks of shearing bacterial DNA during the extraction and digestion steps, bacterial cells are immobilized prior to processing by incorporation in agarose gel. DNA extraction for PFGE is characterized by the need to prolong contact between agarose plugs and the lysis solution that must be distributed throughout the gel. However, the duration of DNA preparation has been shortened since the initial description of the method. The method described in this posting works well with Gram-positive and Gram-negative rods of clinical interest.

Materials that are needed

TE buffer: 10 mM Tris-HCl, 1 mM EDTA, pH 8.
Lysis buffer: 6 mMTris-HCl, 1M NaCl, 0.1 M EDTA, 0.5% sodium N-lauroyl-sarcosine, pH 8.

Other detergents such as sodium deoxycholate (0.2%) and Brij 58 (0.5%) (Sigma Chemicals, St. Louis, MO) may be added.
Lyzozyme solution: 85 mg/mL in sterile water.
Lysostaphine solution: 1.000 U/mL in sterile water. The stock solution is stored in aliquots frozen at -20^oC.
Low melting agarose: 2% in sterile water.
Sterile petri dishes 55 mm in diameter.
Digestion solution: EDTA 0.5 M, 1% sodium N-lauroyl sarcosine, proteinase K 2 mg/mL, pH 8.
PMSF (phenylmethysulfonyl-fluoride): 40 mg/mL in isopropanol. PMSF is harmful, so avoid contact with mucous membranes and eyes. It should not be inhaled and the solution must be handled in a chemical hood.

The Method is

Quantities are for the preparation of five plugs of 200 microliters each.

Grow bacteria in appropriate liquid medium overnight and shake if necessary.
Pellet cells by centrifugation at 2000g and wash them twice with TE buffer. The amount of cells required is equivalent to that found in 5 mL of broth culture with an optical density of 1.0 at 600 nm.
Resuspend cells in 500 microliters of TE buffer.
Add 75 microliters of lyzozyme solution. For staphylococci, also add 15 microliters of lysostaphine.
Add 750 microliters of melted (55^oC) 2% low melting agarose.
Mix thoroughly and pipet the agarose-cell suspension into the wells of a plug mold. Allow agarose plugs to solidify for at least 10 min at 4^oC.
Gently remove the plugs from the mold and transfer them into a sterile petri dish containing 10 mL of lysis buffer. Incubate for 2 h at 37^oC for Gram-negative bacteria and for 4 h for Gram-positive bacteria.
Remove the lysis buffer and replace it with 10 mL of the digestion solution. Incubate the petri dishes at 50^oC overnight. Close them firmly with parafilm to avoid evaporation of the solution.
Cool the petri dish at 4^oC. Transfer the agarose plugs into a new petri dish containing 10 mL of TE buffer. Wash the plugs three times in 10 mL of TE buffer with gentle shaking.
Remove the last 10 mL of washing buffer, add 5 mL of TE buffer.
Add 60 microliters of PMSF isopropanol solution twice 30 min apart and incubate at 50^oC. Rinse plugs three times in TE buffer.

PMSF is used to inhibit residual proteinase K activity. It is insoluble in aqueous solution where its half-life is 60 min. This step is not included in many protocols which use lower proteinase K concentrations.
Store plugs in 0.2 M EDTA at 4^oC. DNA prepared and stored is stable for several months. Plugs have to be rinsed three times for 10 min each in 10 mL of TE buffer to eliminate EDTA before digestion of DNA with restriction enzymes.

Regards.

Reference: Number 12 on References

Polymerase Chain Reaction (PCR): Basic Principle

noreply@blogger.com (Unknown) — Thu, 23 Apr 2009 07:35:00 +0000

Polymerase chain reaction (PCR) is a primer mediated enzymatic amplification of specifically cloned or genomic DNA sequences. PCR process was invented by Kary Mullis and it has been automated for routine use in laboratories worldwide. The main purpose of the PCR process is to amplify template DNA using thermostable DNA polymerase enzyme which catalyzes the buffered reaction in which an excess of an oligonucleotide primer pair and four deoxynucleoside triphosphates (dNTPs) are used to make millions of copies of the target sequence.

Scope of PCR are:

Used in molecular biology and genetic disease research to identify new genes.
Viral targets, such as HIV-1 and HCV can also be identified and quantitated by PCR.
Active gene products can be accurately quantitated using RNA-PCR.
In such fields as anthropology and evolution, sequences of degraded ancient DNAs can be tracked after PCR amplification.
With its exquisite sensitivity and high selectivity, PCR has been used for wartime human identification and validated in crime labs for mixed-sample forensic casework.
In the realm of plant and animal breeding, PCR techniques are used to screen for traits and to evaluate living four-cell embryos.
Environmental and food pathogens can be quickly identified and quantitated at high sensitivity in complex matrices with simple sample preparation techniques.

PCR Process

The PCR process requires a repetitive series of the three fundamental steps that defines one PCR cycle:

Double-stranded DNA template denaturation.

The DNA sequence which is to be amplified by PCR is known as the template. The double-stranded DNA template must be denatured into two complementary single strands of DNA before the reaction can commence. DNA undergoes rapid denaturation at 94⁰ C. Five minutes are allowed for each denaturation step in a 30 cycle PCR, the enzyme will be subjected to 94⁰ C for a total time of 2.5 hours. In practice, 30 seconds to 2 minutes are allowed for the denaturation step, with 1 minute being the optimum choice for most templates.
Annealing of two oligonucleotide primers to the single-stranded template.

After denaturation, the reaction is quickly cooled, preventing immediate reannealing of long DNA strands. Due to their small size, oligos now rapidly anneal to the single strands of DNA at positions containing specified template sequence. In these positions, they act as primers for Taq polymerase. Formation of the specific primer-template complex is highly temperature dependent, so for annealing to take place, the temperature of the reaction must be lowered to a preset level calculated to maximize primer-template interaction.
Enzymatic extension of the primers to produce copies that can serve as templates in subsequent cycles.

Also known as polymerization, this is the final step of the PCR cycle in which the temperature of the reaction is adjusted to the optimum for Taq polymerase activity, which is between 75⁰ C and 80⁰ C. During this step, the polymerase enzyme incorporates nucleotides into the nascent DNA strand, producing a complementary copy of the DNA template in the region specified by the annealed primer. The new temperature is invariably above that at which annealing occurs, but does not lead to denaturation of primer-template complex, presumably because the enzyme is already active at the annealing temperature and significantly increases primer length during the annealing step, y\thus raising its denaturation temperature above that of the polymerization step.

Regards

Reference: Number 4 and 11 in References

The Measurement of Tryptophan Content by Using UV-Spectrometer

noreply@blogger.com (Unknown) — Mon, 13 Apr 2009 08:22:00 +0000

The absorption of protein solutions in the UV is the result of tryptophan and tyrosine (and to a very minor, and negligible, extent phenylalanine and cysteine). The absorption maximum will depend on the pH of the solution, and spectrophotometric measurements are usually made in alkaline solutions. Absorption curves for tryptophan and tyrosine show that at the points of intersection, 257 and 294 nm, the extinction values are proportional to the total tryptophan + tyrosine content. Measurements are normally made at 294.4 nm, since this is close to the maximum in the tyrosine curve, and in conjunction with the extinction at 280 nm, the concentrations of each of the two amino acids may be calculated. This is the method of Goodwin and Morton.
Before measure tryptophan content you have to prior hydrolyze the protein sample, see the method in my previous posting.

Here is the method of the measurement of Tryptophan content:

The protein sample is made 0.1M in NaOH.

Absorption by most proteins in 0.1M NaOH solution decreases at longer wavelengths into the region 330--450 nm, where tyrosine and tryptophan do not absorb. Suitable blanks for 294 and 280 nm are therefore obtained by measuring extinctions at 320 and 360 nm and extrapolating back to 294 and 280 nm.

Measure the absorbance at 294.4 and 280 nm in cuvets (transparent to this wavelength, i.e., quartz) in a spectrometer.
The amount of tryptophan (w) is estimated from the relative absorbances at these wavelengths
by the method of Goodwin and Morton (2) shown in Equation below:

E280 = w Ew + (x-w)Ey

Therefore:

w = (E280 – x Ey) / (Ew -Ey)

where x = total mol/L, w = tryptophan mol/L, and (x- w) = tyrosine mol/L, Ey = Molar extinction of tyrosine in 0.1M alkali at 280 nm= 1576. Ew = Molar extinction of tryptophan in 0.1M alkali at 280 nm = 5225.

Also, x is measured from E294.4 (the molar extinction at this wavelength). This is 2375 for both Tyr and Trp (since their absorption curves intersect at this wavelength). An accurate reading of absorbance at one other wavelength is then sufficient to determine the relative amounts of these amino acids.
An alternative method of obtaining the ratios of Tyr and Trp is to use the formulae derived by Beaven and Holiday.

MTyr = (0.592 K294- 0.263 K280) / 1000

MTrp = (0.263 K280 - 0.170 K294) / 1000

where MTy r and MTrp are the moles of tyrosine and tryptophan in 1 g of protein, and K294 and K280 are the extinction coefficients of the protein in 0.1M alkali at 294 and 280 nm. Extinction values can be substituted for the K values to give the molar ratio of tyrosine to tryptophan according to the formula below:

MTyr / MTrp = (0.592 E294 - 0.263 E280 / 0.263 E280 - 0.170 E294)

In this analysis, the tyrosine estimate may be high and that of tryptophan low. If amino acid analysis indicates absence of tyrosine, tryptophan is more accurately determined at its maximum, 280.5 nm.

Reference: Number 6 in References

Bicinchoninic Acid (BCA) Method: A Protein Assay

noreply@blogger.com (Unknown) — Sat, 11 Apr 2009 18:16:00 +0000

The Bicinchoninic acid (BCA) assay first was described by Smith, et al. BCA assay is similar to Lowry assay since it also depends on the conversion of Cu(2+) to Cu(+) under alkaline conditions. The Cu(+) is then detected by reaction with BCA. The reaction results in the development of an intense purple color with an absorbance maximum at 562 nm. BCA method and Lowry are of similar sensitivity, but BCA method is more advantageous compared to Lowry in a few things, here are the advantages:

BCA is stable under alkali conditions, so it can be carried out as a one-step process compared to the two steps needed in the Lowry assay.
BCA is more tolerant to the presence of compounds that interfere with the Lowry assay.
it is not affected by a range of detergents and denaturing agents such as urea and guanidinium chloride, although it is more sensitive to the presence of reducing sugars.

Materials that you need:

Standard Assay

Reagent A: sodium bicinchoninate (0.1 g), Na2CO 3.H2O (2.0 g), sodium tartrate (dihydrate) (0.16 g), NaOH (0.4 g), NaHCO3 (0.95 g), made up to 100 mL. If necessary, adjust the pH to 11.25 with NaHCO3 or NaOH.
Reagent B: CuSO4 . 5H2O (0.4 g) in 10 mL of water.

Reagents A and B are stable indefinitely at room temperature. They may be purchased ready prepared from Pierce, Rockford, IL.

Standard working reagent (SWR): Mix 100 vol of regent A with 2 vol of reagent B. The solution is apple green in color and is stable at room temperature for 1 week.

Microassay

Reagent A: Na2CO3 . H2O (0.8 g), NaOH (1.6 g), sodium tartrate (dihydrate) (1.6 g), made up to 100 mL with water, and adjusted to pH 11.25 with 10M NaOH.
Reagent B: BCA (4.0 g) in 100 mL of water.
Reagent C: CuSO4 . 5H20 (0.4 g) in 10 mL of water.
Standard working reagent (SWR): Mix 1 vol of reagent C with 25 vol of reagent B, then add 26 vol of reagent A.

The Bicinchoninic Acid method are:

Standard Assay

To a 100-1aL aqueous sample containing 10-100 ~g protein, add 2 mL of SWR. Incubate at 60 Celcius degree for 30 min.

The sensitivity of the assay can be increased by incubating the samples longer. Alternatively, if the color is becoming too dark, heating can be stopped earlier. Take care to treat standard samples similarly.

Cool the sample to room temperature, then measure the absorbance at 562 nm.

Following the heating step, the color developed is stable for at least 1 hour.

A calibration curve can be constructed using dilutions of a stock 1 mg/mL solution of bovine serum albumin (BSA).

Note, that like the Lowry assay, response to the BCA assay is dependent on the amino acid composition of the protein, and therefore an absolute concentration of protein cannot be determined. The BSA standard curve can only therefore be used to compare the relative protein concentration of similar protein solutions.

.
Microassay

To 1.0 mL of aqueous protein solution containing 0.5-1.0 ~tg ofprotein/mL, add 1 mL of SWR.
Incubate at 60 Celcius degree for 1 h.
Cool, and read the absorbance at 562 nm.

Regards

Reference: Number 8 in References

Protein Assay using UV-Spectrophotometer at 280 nm

noreply@blogger.com (Unknown) — Tue, 31 Mar 2009 18:56:00 +0000

It is possible to estimate protein concentration in a solution by using simple spectrometer. Absorption of radiation in the near UV (280 nm) by proteins depends on the Tyrosine and Tryptophan content (also to a very small extent on the amount of Phenylalanine and disulfide bond).

The advantages of using this method are:

It is very simple.
The sample can be recovered.

While the disadvantages are:

Interference from other chromophores.
The specific absorption value for a given protein must be determined.
The extinction of nucleic acid in the 280-nm region may be as much as 10 times that of protein at their same wavelength, and hence, a few percent of nucleic acid can greatly influence the absorption.

Here is the step by step method:

Prepare a reliable Spectrophotometer.
The protein solution must be diluted in the buffer to a concentration that is well within the accurate range of the instrument.

It is best to measure absorbance in the range 0.05-1.0. At around 0.3 absorbance, the accuracy is greatest.
Bovine serum albumin is frequently used as a protein standard; 1 mg/mL has an A-280 of 0.66.

The protein solution to be measured can be diluted in a wide range of buffers.

If the solution is turbid, the apparent A-280 will be increased by light scattering, so it will need filtration (you can use 0.2 micrometers Millipore filter) or clarification by centrifugation. For turbid solutions, a convenient approximate correction can be applied by subtracting the A-310 (proteins do not normally absorb at this wavelength unless they contain particular chromophores) from the A-280.
At low concentrations, protein can be lost from solution by adsorption on the cuvet; the high ionic strength helps to prevent this. Inclusion of a nonionic detergent (0.01% Brij 35) in the buffer may also help to prevent these losses.

Measure the absorbance of the protein solution at 280 nm, using quartz cuvets or cuvets that are known to be transparent to this wavelength, filled with a volume of solution sufficient to cover the aperture through which the light beam passes.
The actual value of UV absorbance for a given protein must be determined by some absolute method, e.g., calculated from the amino acid composition, which can be determined by amino acid analysis. The UV absorbance for a protein is then calculated according to the following formula:

A280 (1 mg/mL) = (5690Nw + 1280Ny + 120Nc)/M

where Nw, Ny, and Nc are the numbers of Trp, Tyr, and Cys residues in the polypeptide of mass M and 5690, 1280 and 120 are the respective extinction coefficients for these residues.

The presence of nonprotein chromophores (e.g., heme, pyridoxal) can increase A-280. If nucleic acids are present (which absorb strongly at 260 nm), the following formula can be applied.

Protein (mg/mL) = 1.55 A280 -0.76 A260

This gives an accurate estimate of the protein content by removing the contribution to absorbance by nucleotides at 280 nm, by measuring the A-260 which is largely owing to the latter. Other formulae (using similar principles of absorbance differences) employed to determine protein in the possible presence of nucleic acids is the following:

Protein (mg/mL) = (A-235 - A-280)/2.51
Protein (mg/mL) = 0.183 A-230 - 0.075.8 A-260

Regards

Reference: Number 5 on References

Surfactant Properties: How to Quantitatively Measure

noreply@blogger.com (Unknown) — Sun, 29 Mar 2009 17:22:00 +0000

In order to measure the properties of surfactant, we can use the surface tension, emulsification activity, and hemolytic activity as the parameters. Here are the methods:

Surface tension analysis was carried out using the method of Makkar et al.

Aqueous solutions of MC and PPGF were prepared at concentrations of 0.1, 0.5, 1, 2.5 and 5 mg/ml.
Triton X-100 well known chemically synthesized surfactant was employed as a control.
The surface tension of each sample was measured five times using a Du Nouy ring equipment and the average was calculated.

Assay for emulsification activity was carried out according to the modified method of Navon-Venezia et al.

One hundred microliters of each hydrocarbons and olive oil, was added to 10 ml of 20 mM sodium phosphate buffer (from pH 5.0 to 7.5) or 20 mM Tris-HCl buffer (from pH 7.0 to 9.0) containing Surfactant, or Triton X-100 at 1 mg/ml in a 50 ml grass tube.
Shaking reciprocally in a shaker at 30oC for 1 h and standing for 10 minutes.
The lower phase was removed and absorbance was measured at 620 nm.

Assay for hemolytic activity was carried out according to the modified method Morán et al.

Fresh human blood was mixed with an incomplete medium (1:1 v/v) containing (per liter): 10.4 g of RPMI 1640 Media, 5.95 g of HEPES, 0.5 g of L-glutamine, 2 g of NaHCO3, and 105 units of penicillin-G.
The mixture was centrifuged at 3,000 rpm for 20 min at 4 Celcius degree and the supernatant was discarded.
This procedure was repeated twice and the precipitate was resuspended in the medium (1:1 v/v).
One hundred microliters of an aqueous solution of PPGF and protein fractions purified by electrophoresis was added into a mixture of 80 microliters of 20 mM sodium phosphate buffer containing 40 mM NaCl and 20 microliters of the red blood cell suspension described above containing approximately 18,000 cells in a 96-well microplate having a u-bottom.
After incubation at 37ºC for 30 min, the minimum concentrations showing hemolytic activity were determined.

Regards

DNA Extraction Using Prepman Ultra

noreply@blogger.com (Unknown) — Wed, 25 Mar 2009 06:37:00 +0000

In my previous posting I’ve slightly given the bacterial DNA extraction method using Instagene matrix. In this particular posting, I want to give you the more simple method to extract DNA, which is using Prepman Ultra. According to the producer, Applied Biosystem, Prepman ultra is applicable to successfully preparing DNA template from bacteria, yeast, filamentous fungi, both from a plate or from tissue smears.

Here is the method:

Picking out of 1 ose of the bacteria colony from the plate into 200 microliters of Prepman ultra reagent (placed in the sterile tube).
Incubate the reagent+colony in 100 Celcius degree water-bath for 10 minutes.
Cool the tube at the room temperature for 2 minutes.
Centrifuge the tube at 10000 rpm for 2 minutes at 4 Celcius degree.
Taking out the supernatant and leave the cell in the tube.
Now you have 200 microliters of DNA template.
You can transfer 5 microliters for the assay.

It is just as simple as that to get the DNA template, thanks to Applied Biosystem for this helpful kit.

Regards

PCR Components

noreply@blogger.com (Unknown) — Tue, 24 Mar 2009 09:05:00 +0000

The Polymerase Chain Reaction (PCR) created a revolution in molecular biology research and its applications. PCR is an in vitro method that enzymatically amplifies specific DNA sequences using oligonucleotide primers that flank the region of interest in the target DNA. The principle involves a repetitive series of cycles each of which consist of template denaturation, primer annealing, and extension of the annealed primers by a DNA polymerase to create the exponential accumulation of a specific fragment whose ends are determined by the 5’ ends of the primers. The PCR is so named because it involves a polymerase and the products synthesized in each cycle can serve as templates in the next so the number of DNA copies approximately doubles at every cycle to create a chain reaction similar to the principles in a nuclear reactor.

Polymerase enzyme used in the PCR reaction is isolated from Thermus aquaticus, therefore it is named Taq polymerase. Taq polymerase is a thermostable enzyme that enabled the amplification reaction to be carried out by cycling the temperature within the reaction tube after mixing all the reaction components.

The taq polymerase enzyme works with high efficiency in relatively simple buffer systems. Thus the essential components of the polymerase chain reaction include: the DNA template, two defined oligos to act as primers, the four deoxynucleotide triphosphates (dNTPs) for incorporation into the new DNA strands, Taq polymerase and magnesium, which is the cationic cofactor of the enzyme, in a Tris buffer of appropriate pH and salt concentration. The salt most frequently used is potassium chloride. A layer of light mineral oil is always placed on top of each reaction to prevent evaporation which would otherwise stop the reaction within a few cycles.

Those basic components remain largely unchanged for most applications of PCR, but their concentrations and the buffering pH of the reaction vary according to the type of experiment, the nature of the primers and/or template, and the experience of the experimenter. The reaction mix of the PCR (the concentrationand pH ranges) can see in the following table.

You can also use the standard PCR component, here is the list:

10 mM Tris-HCl pH 8.4

50 mM KCl

1 mM Magnesium chloride

0.05% (v/v) Tween 20

0.2 mM dNTPs

100 pmol each primer

1 U Taq polymerase

Regards

Reference: Number 4 on References

Bradford Method: Colorimetric Protein Assay

noreply@blogger.com (Unknown) — Sun, 22 Mar 2009 10:59:00 +0000

Bradford method is a common colorimetric method to determine protein concentration in a sample solution. The Bradford method of protein determination is based on the binding of a dye, Coomasie Blue G, to the protein. This binding shifts the absorbtion maximum of the dye from red to blue. The absorbance of the solution is measured at 595 nm and is proportional to protein concentration when compared to a standard curve. Two types of assay are described here: the standard assay, which is suitable for measuring between 10 and 100 microgram of protein, and the microassay, which detects between 1 and 10 microgram of protein.

Materials

Coomasie Blue G

Preparation of the 5x Bradford Reagent

Dissolve 100 mg of Coomasie Blue g in 50 ml of ethanol and 100 ml of phosphoric acid (85%). The mixture is stirred for approximately 10 minutes. The solution is diluted to 200 ml with distilled water and filtered.

Ethanol

Phosphoric acid, 85% - Caution! Phosphoric acid can cause burns!

Test Tubes

Protein Standard

Protein standard. Bovine gamma-globulin at a concentration of 1 mg/mL (100 micrograms/mL for the microassay) in distilled water is used as a stock solution. This should be stored frozen at –20^oC. Since the moisture content of solid protein may vary during storage, the precise concentration of protein in the standard solution should be determined from its absorbance at 280 nm. The absorbance of a 1 mg/mL solution of gamma-globulin, in a 1-cm light path, is 1.35. The corresponding values for two alternative protein standards, bovine serum albumin and ovalbumin, are 0.66 and 0.75, respectively.

Spectrophotometer

Micropipettes and tips

Method-Protein Determination by the Bradford Method

Standard Assay Method

Pipet between 10 and 100 microgram of protein in 100 microLiter total volume into a test tube. If the approximate sample concentration is unknown, assay a range of dilutions (1, 1:10, 1:100, 1:1000). Prepare duplicates of each sample.
For the calibration curve, pipet duplicate volumes of 10, 20, 40, 60, 80, and 100 microliter of 1 mg/mL gamma-globulin standard solution into test tubes, and make each up to 100 microliter with distilled water. Pipet 100 microliter of distilled water into a further tube to provide the reagent blank.
Add 5 mL of protein reagent to each tube and mix well by inversion or gentle vortex mixing. Avoid foaming, which will lead to poor reproducibility.
Measure the A-595 nm of the samples and standards against the reagent blank between 2 min and 1 h after mixing. The 100 μg standard should give an A-595 value of about 0.4. The standard curve is not linear, and the precise absorbance varies depending on the age of the assay reagent. Consequently, it is essential to construct a calibration curve for each set of assays.

Microassay Method

This form of the assay is more sensitive to protein. Consequently, it is useful when the amount of the unknown protein is limited.

Pipet duplicate samples containing between 1 and 10 microgram in a total volume of 100 microliter into 1.5-mL polyethylene microfuge tubes. If the approximate sample concentration is unknown, assay a range of dilutions (1, 1:10, 1:100, 1:1000).
For the calibration curve, pipet duplicate volumes of 10, 20, 40, 60, 80, and 100 microliter of 100 microgram/mL gamma-globulin standard solution into microfuge tubes, and adjust the volume to 100 microliter with water. Pipet 100 microliter of distilled water into a tube for the reagent blank.
Add 1 mL of protein reagent to each tube and mix gently, but thoroughly.
Measure the absorbance of each sample between 2 and 60 min after addition of the protein reagent. The A-595 value of a sample containing 10 microgram gamma-globulin is 0.45. Following Figure shows the response of three common protein standards using the microassay method.

Variation in the response of proteins in the Bradford assay. The extent of protein–dye complex formation was determined for bovine serum albumin (Square), gamma-globulin (Circle), and ovalbumin (Triangle) using the microassay.

The Bradford method doesn’t measure the presence of peptide bonds but detects specific amino acids, which is believed to be responsible for the binding of the dye to the protein. The dye does not bind to free arginine or lysine, or to peptides smaller than about 3000 Da. The Bradford assay is not suitable for quantifying the amounts of such compounds.

Regards

Reference: Number 2 and 10 on References

Quantitative Estimation of DNA Concentrations

noreply@blogger.com (Unknown) — Fri, 20 Mar 2009 16:09:00 +0000

DNA, RNA, and protein strongly absorb ultraviolet light in the 260 to 280 nm range. UV spectroscopy can be used as a quantitative technique to measure nucleic acid concentration and protein contamination. Nucleic acids strongly absorb at 260 nm and less strongly at 280 nm while proteins do the opposite. The general rules for determining the concentrations of nucleic acids at 260 nm are:

1 Optical Density (OD) unit of double-stranded DNA is 50 micrograms/ml.
1 OD unit of single-stranded DNA is 33 micrograms/ml.
1 OD unit of single-stranded RNA is 40 micrograms/ml.

Proteins absorb strongly at 280 nm where 1 OD unit is 1 mg/ml. When using UV spectroscopy for estimating DNA concentrations, it is very important to remove all protein and RNA from the DNA solution. Good estimations can only be made on clean preparations.

An estimate of the purity of a DNA preparation can be made by measuring the absorbance at both 260 nm and 280 nm. Pure solutions of nucleic acid will absorb approximately twice as much at 260 nm as at 280 nm. Experimentally, the ratio of 260 nm/280 nm of a pure DNA solution is between 1.8 to 2.0. As protein contamination increases, the ratio decreases. Additionally, the presence of contaminating oranic solvents, such as phenol, can affect estimations of concentration and purity.

Materials you need are:

UV Spectrophotometer

Quartz or UV compatible cuvettes

TE buffer

DNA template

Method:

Fill the cuvette with water or TE buffer. Zero the spectrophotometer at 260 nm with this blank.
DNA from plasmid and genomic preparations is typically at a concentration exceeding 1 micrograms/microliter. Consequently, DNA is usually diluted before measuring its absorbance. An unfortunate result of this measurement is that the DNA is expended as a result of the dilution. Be sure these is adequate DNA to waste. Start by diluting the DNA sample 1 microliter : 999 microliters of TE buffer (the dilution can be done directly in the cuvette). Mix the dilution thoroughly.
Measure the optical density (OD). Multiply the resulting OD by 50 micrograms/ml. For a 1:1000 dilution, the mass of DNA is equal to micrograms/microliter.
Similarly, the same sample can be measured at 280 nm. A ratio of the OD-260nm/OD-280nm is an indicator of DNA purity. A ratio of 1.8 or higher indicates minimal protein contamination.

Measuring DNA concentration is not difficult, is it?

Regards

Reference: Number 2 on References

Estimation of DNA Concentrations

noreply@blogger.com (Unknown) — Thu, 19 Mar 2009 15:39:00 +0000

The determination of the concentration of DNA or RNA in solution is a fundamental task in molecular biology. DNA is usually the limiting reagent in most experiments; therefore, the knowledge of its concentration is critical. Determination of the DNA concentration can be estimated either by qualitatively comparing the fluorescence of DNA bands in an agarose gel to a standard or by spectrophotometric means.

Qualitative Estimation

DNA fluorescence in the presence of ethidium bromide, and the intensity of the fluorescence is proportional to its concentration. As such, comparison of relative fluorescence of unknown DNA to known standards can be used as a rough estimation of DNA concentration. This comparison is usually made following the electrophoresis of the standard (for example, 1 microgram of Lambda DNA cleaved with the restriction endonuclease HindIII) and the unknown. The mass of 1 microgram of a Lambda-HindIII standard provides convenient references for comparison.

An alternative method for estimating DNA concentration relies on the lower limit for visually detecting DNA in a gel. It has been estimated that a band of DNA below 5 nanograms is not detectable by the human eye. Serially diluting DNA to extinction, followed by electrophoresis, can also be used to measure DNA concentration. This technique is particularly if there is more than one species or fragment of DNA in a sample. In this manner, the concentration of individual bands can be estimated.

Reference: Number on References

Measurement of Biosurfactant Activity

noreply@blogger.com (Unknown) — Wed, 18 Mar 2009 06:52:00 +0000

Biosurfactant activity was measured by an oil displacement test. This method is so sensitive that only a small amount of sample is required to measure the surfactant activity. Here is the method:

A sample is solubilized or suspended in 10 micro liters of 10% acetonitrile or 0.1 M Tris-Cl (pH 8) buffer.
After solubilized or suspended , a sample was put on the center of an oil membrane which was formed on the surface of water in a large size petri dish (15 cm diameter).
The size of the resultant oil-displaced circle area reflects the activity of a surfactant.
The larger the size is, the higher the activity of a surfactant is.
One BS unit was defined as the amount of surfactants forming 1 cm square of oil displaced area.
The surface tensions were measured using a ring tensiometer (K6; Kruss, Germany).

Regards

Reference: Number 3 on References

Advertise with Us

noreply@blogger.com (Unknown) — Wed, 11 Mar 2009 16:29:00 +0000

If you intent to advertise your products or services with us, please send your request or an email to this email address:

biotechmethods@gmail.com

Email should contain these following things:

Your personal information.
Your bid (this is negotiable and reasonable).
Your request for ads placement.

As soon as possible I will respond your request. Thanks for your cooperation.

Regards

Isolation of Yeast Genomic DNA

noreply@blogger.com (Unknown) — Thu, 26 Feb 2009 04:43:00 +0000

Isolating genomic DNA from yeast involves culturing the microbe, harvesting the cell, enzymatically removing the cell wall, lysing the protoplast, and finally separating the DNA from the other cell debris.
Materials that you need are:

Yeast culture-prepared previously

Spectrophotometer with cuvettes

50 mM EDTA, pH 8-ice cold

50 mM Tris, pH 9.5, 2% 2-mercaptoethanol

1.2 M sorbitol, 50 mM Tris, pH 7.5

Lyticase solution-500 U/ml in 50 mM Tris, pH 7.5

10% Sodium Dodecyl Sulfate (SDS)-used for checking protoplast formation

Lysis buffer-100 mM Tris, pH 7.5, 100 mM EDTA, 150 mMNaCl, 50 micrograms/ml RNase A

Lysis buffer with 2% SDS

95% Ethanol-stored at minus 20 degree Celcius

TE buffer-10 M Tris, pH 8, 1 mM EDTA

3 M potassium acetate, pH 5.5

Here are the step by step methods:

The yeast can be cultured for as long as 48 hours at 30 degree Celcius. The optical density of a 1:10 dilution of the culture in water can be as high as 1.0 at 520 nm.
Harvest 5 ml of cells by centrifugation (5 minutes at 5000 rpm). Resuspend the yeast in 1 ml of cold 50 mM EDTA, pH 8, and transfer to a 1.5 ml microfuge tube. Centrifuge for 1 minute, decant, and resuspend again in 50 mM EDTA.
Pellet the cells as before and suspend the cells in 1 ml of 50 mM Tris, pH 9.5, 2% 2-mercaptoethanol. Incubate for 10 min at room temperature. Centrifuge and decant.
Resuspend the cells in 800 micro liter of 1.2 M sorbitol, 50 mM Tris, pH 7.5. The sorbitol act as an osmotic support and prevents rupture of the cells as the wall is removed. As the yeast cell walls degrade, membranes can easily overextend and rupture.
Add 200 micro liter of Lyticase (500 U/ml in 50 mM Tris, pH 7.5). Place the cells on a rocker and incubate at 37 degree Celcius for one hour. Lyticase is a yeast cell wall degrading enzyme isolated from the bacteria Arthrobacter luteus.
Examine the suspension under a microscope to ensure protoplast formation. As the yeast wall is degraded, the cell membrane can ooze out of the sack. Viewed with phase contrast microscopy, yeast protoplasts are characteristically refractile (or bright) spheres, and yeast cell wall shells appear as gray ghosts (cell walls without membrane and cytosol). Combine 10 micro liter of 10% SDS with 10 micro liter of yeast protoplasts. Examine the cells under the microscope. The absence of refractile yeast indicates the protoplasts were lysed by the SDS.
Pellet the protoplasts by centrifuging at 10000 rpm for five minutes. Resuspend the cells in 1 ml of 100 mM Tris, pH 7.5, 100 mM EDTA, 150 mM NaCl (lysis buffer). Transfer the cells to a 5 ml polypropylene tube. Add 1 ml of lysis buffer with 2% SDS. Mix and incubate at 30 Celcius egree for 30 minutes. Check the cells under a microscope for lysis.
Centrifuge the lysate at 5000 rpm for 15 min to pellet cellular debris. Decant the upper phase containing the DNA.
Using a pipet, determine the volume of the DNA solution. Add 1/10th volume (e.g., 100 micro liter for every ml) of 3 M potassium acetate to the solution. In the presence of ions Na and K, DNA precipitates if mixed with either ethanol or isopropanol. Incubate the DNA at -20 Celcius degree for 30 min (or overnight if possible. Centrifuge the solution at 7000 rpm for 20 minutes. The DNA appears as white pellet. Decant and remove as much moisture as possible, but do not allow the pellet to dry. Once genomic DNA drys, it can be very difficult to resuspend.
Resuspend the DNA in 100 micro liter of TE buffer and freeze.

Reference: number 2 on References