User:Pranav Rathi/Notebook/OT/2012/10/01/Buffer preparation for DNA overstretching & unzipping experiments

We primarily use popping buffer and PBS for DNA-tethering, unzipping and overstretching experiments. All the other buffers consist these two. This page discusses the chemicals and process used in preparation of the buffers and solutions.

Popping Buffer

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POP we use for DNA tethering, unzipping and overstretching experiments. It is funny but true that it's called "popping buffer" because when DNA binding proteins are present they are "popped" off the DNA when it is unzipped. Popping buffer or POP is the main solution used to prepare the samples. It can be prepared in H₂O or D₂O. The primary purpose of this buffer is to maintain the PH-level and stabilize the DNA or DNA-protein complex in the solution. The popping buffer I use is 1X; which means the chemical-concentration is whatever it is in standard POP. By doubling or tripling it, 2X or 3X POPs can be made.

• The chemicals used in POP are as follows:

EDTA: Ethylenediamineteraacetic acid disodium salt dehydrate. EDTA usually binds to metal cation, such as mg+2 ions and ca+2 ions. This makes DNA-protein complex more stable and prevent protein or enzymes to cut the DNA.[1]

Sodium phosphate

monobasic: Sodium phosphate monobasic is H2Nao4P, it helps the PH-level by taking or giving OH- and H+ ions.[2]

dibasic: Sodium phosphate dibasic is HNa2O4P, it also helps PH-level maintenance. [3]

NaCl: Sodium chloride also helps with PH-maintenance. [4]

Tween 20: Polyethylene glycol sorbitan monolaurate is a detergent which prevents non-specific antibody binding and to saturate binding sites on surface. Basically it helps with DNA-tethering which uses anti-dig and dig bonding.[5]

H₂O: Primary solvent.[6]

D₂O: Primary solvent. Either one can be used depending on the experiments.[7]

Making Popping buffer (POP)

Let's say I need 100mL of 1X POP in H₂O or D₂O with the following concentration of chemicals:

• Desired Buffer-Volume: 100mL
• EDTA: 10mM in 100mL of buffer volume
• Sodium phosphate: Total concentration is 50mM in 100mL of buffer volume

Mono is 19% of 50mM + Di is 81% of 50mM =50mM in 100mL of buffer volume

• NaCl: 50mM in 100ml of buffer volume.
• Tween20: .02% of 100ml

To get these desired concentrations I convert the moles into grams and prepare the following stock in the following way:

It is always good to keep the chemicals in stock, so the weighed mass of chemicals from the bottle are based on the stock volume, but the final concentration in the buffer volume will be based on the volume which is mixed into the buffer to make final buffer;

• EDTA: 10mL is mixed.
• Sodium phosphate:

mono: 1.9mL is mixed

di: 8.1mL is mixed(this is to keep the PH at 7.5)

• NaCl: 1.25mL is mixed
• Tween20: 1mL is mixed
• H₂O or D₂O: 77.75mL is taken

The final volume is 100mL. Stock volume is completely based on my desire, so all these concentrations and volumes are needed for calculations for weighted chemical masses from the bottles.

Calculations:

I have written a simple code in LabView V9.1 with the download link:[8]

The math is following:

• Desired buffer concentration is Cb(M)
• Desired final-buffer volume is Vb(L)
• Callulated stock concentration is Cs(M)
• Volume mixed into the buffer is Vm(L)
• Mass written on the bottle is Mw(gm/mole)
• Stock volume Vs(L)
• Calculated mass to be measured is M(gm)

First to calculate the stock concentration:

[math]\displaystyle{ C_s=C_b*V_b/V_m }[/math] in moles

Now to calculate mass to be measured:

[math]\displaystyle{ M=M_w*C_s*V_s }[/math] in gm

Now this mass is weighted/measured on the scale and mix with the desired stock volume of H₂O or D₂O, then mix volume is taken from the stock to mix into the final buffer to get the final concentration. All the calculation are given in the excel spreadsheet: {{#widget:Google Spreadsheet |key=0ApjWjFYiQdkfdENGTm5kckg5dTNfN01uQjh2YUVyZWc |width=500 |height=300 }}

Procedure

1. Weight the chemicals on the scale
2. Measure and mix the chemicals in the solvent and mix using vortex
3. Measure the volume and mix into the buffer
4. Store at +3C

See the slide share for more information

Next buffer is PBS.

PBS Buffer

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PBS is Phosphate Buffer Saline. It is a buffer solution commonly used in biological research. It is a water-based salt solution containing sodium chloride, sodium phosphate, and, in some formulations, potassium chloride and potassium phosphate. The buffer's phosphate groups help to maintain a constant pH. The osmolarity and ion concentrations of the solution usually match those of the human body (isotonic). We use PBS to make Anti-digsolution for DNA-tethering.[9]

• The chemical used in POP are as follows:

Salt	Concentration	Concentration
(—)	(mmol/L)	(g/L)
NaCl	137	8.01
KCl	2.7	0.20
Na2HPO4 • 2 H2O	10	1.78
KH2PO4	2.0	0.27
pH	7.4	7.4

Making PBS

The simplest way to prepare a PBS solution is to use PBS buffer tablets (see slides). They are formulated to give a ready to use PBS solution upon dissolution in a specified quantity of H₂O or D₂O. They are available in the standard volumes: 100, 200, 500 and 1000 mL.

Procedure

Dissolve 1 table in 200mL of H₂O or D₂O.

Using these two buffers (POP & PBS) all other solutions used in DNA- tethering, unzipping and overstretching are prepared.

Blocking Solution (BGB)

The purpose of blocking solution is to block exposed glass surfaces after binding anti-dig. Various kinds of casein are typically used, which I think evolves from the common practice of using non-fat dried milk (NFDM) in standard wet-lab protocols, such as western blotting. NFDM is predominantly casein, and so people use NFDM and casein interchangeably, usually ignoring the fact that differences in purity or kinds of casein could potentially impact a sensitive assay. Often it is imagined that casein is a regular soluble protein, but Dr.Koch found in the past that casein forms polydisperse micelles, probably. He doesn't know whether these polydisperse micelles are important for it's blocking capabilities, but he did find some references that said they are (small micelles fill gaps in big micelles). Brent Brower-Toland, being a good biologist, ignored the anlaysis paralysis of physicists and just ordered cheap good blocker from Bio-Rad, called "Blotting-Grade Blocker" [10] at Bio-rad.com. This worked very well and we continue to use it. Bio-Rad calls it "non fat dried milk," so I'm not sure if it's the same stuff you get at the supermarket. We'll call this BGB from now on (which can also read as "Brent's Good Blocker.")

• Why use BGB; blotting grade blocker: BGB is casein (α, β and κ) which is hydrophobic and like to be clustered in water. It is used to block the surface where anti-dig is not present (coat that part with casein). Casein make small hemisphericalballs (micelle [11]), these balls fillup the gaps around the anti-dig preventing the beads sticking to the surface.

Procedure

To make 5mg/ml BGB solution in popping buffer:

1. First weight out 15mg of BGB poweder
2. Add 1.5 ml of 1x pop and mix by vortexing
3. Run through .2μM syringe filter using 3ml syringe.
4. Now store 5mg/ml BGB in 1x pop buffer (minus amount of protein that stuck to filter) at 3C.

This should be good for few weeks.

Anti-Dig Solution

Polyclonal sheep anti-digoxigenin from Roche [12]. This is shipped as a lyophilized powder. We always resuspend entire 200 microgram bottle with 1 ml of ice-cold PBS, and then make 20 microliter aliquots which are stored at -80C. An aliquot can be extracted from freezer, and diluted with 180 microliters of cold PBS.

procedure

1. Add 1mL of ice-cold PBS into 200μgm of Anti-dig and mix it.
2. Make 20μgm/ml of aliquots.
3. Nitrogen flash-freeze them and store them at -80C.
4. When use for sample; take out an aliquot from freezer and mix 180mL of cold PBS and store at +3C.

Microspheres Solution

We use two different size (1μM and .5μM) streptavidin coated microspheres.

• 1μM streptavidin coated polymer microspheres are from Bang's Labs.[13]. They are approximately 1.04μM.
• .5μM streptavidin coated polymer microspheres are from Bang's Labs.[14]. They are approximately .53μM.

Both types work fine. In my experience .5μM works better with tethering. 1μM works better with FTC (fine tether center) and calibration. I am trying to increase tethering efficiency with 1μM beads.

Procedure

It is important to wash the beads before put them into the sample to remove the free streptavidin. Free streptavidin can kill SM experiments, because it will diffuse much more quickly than the microspheres, and will quench the small amount of biotin. Washing also lets you pick a usage buffer.

Wash Procedure

Dr.Koch Method

1. Aliquot 50uL beads from stock and put in eppi.
2. Add 950uL 1x popping buffer
3. Centrifuge at 6,600g for 5 min
4. discard supernatant
5. Repeat steps 2-4 for a total of 3-5 times
6. After discarding of supernatant, I measure the Volume and add BGB 4X of it to get 1:5 stock-concentration.

Bangs Labs Method

1. Suspend 50ul beads in 950ul 1x popping buffer.
2. Centrifuge at 2,200g for 15 min (change acceleration depending on bead size).
3. Discard supernatant.
4. Repeat steps 2-3 for a total of 3-5 times
5. After discarding of supernatant, Same as above.

Destroying Clumps

Clumps of beads can easily be seen after washing. It is recommended one destroy these after washing.

It is important to get beads to be suspended individually because clumpiness can really ruin a sample. Below are two ways that this can be achieved. With proof.

Sonication Method

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You will want to sonicate beads for a decent amount of time. The time it takes depends on the size of the beads and the level of clumpiness. Also adding buffer that reduces the hydrophobicity of the bead surface may be of use. I usually sonicate the beads for 180 seconds for both sizes.

Vortexer Method

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In tests done, it seems this method works as well (or as poor depending on your perspective) as sonicating. Vortex in pulses of 5 seconds for however long you deem worthy depending on your volume and level of bead clumps.

• 

  This image was taken after sonication.

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  This image was taken after vortexing (instead of sonicating). Seems as clumpy as sonicating.

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  This is a view of one surface of the sample in vortexing, but the same can be seen after sonication.

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  Instead of washing and sonicating/vortexing I just diluted the stock beads in water. Not dilute enough, but no clumps.

Based on these images, it seems like not washing is the best possible choice because the beads exist in solution as single entities.

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1. Takeout 50μL from bottle to keep in stock
2. Take 5μL and add 45μL of BGB to make 50μL (1:10) bead solution
3. This can be further diluted per requirement.

DNA solution

The tethering protocol (discussed here) has been used for a variety of dig & biotin labeled DNA constructs. One specific fragment is 4.4 kb PCR-labeled DNA. Another is 17-mer unzipping DNA. The protocol will have to be adjusted for very short DNA tethers--one reason being the increased importance of surface charge interactions. Other departures from these types of DNA may also require modifying the protocol. A typical concentration of DNA to use is 20 picomolar (pM).

Procedure

Dilution depends on the DNA concentration and it is done in 1x POP. A quick example of dilution can be like this:

1. 2 μL dna from stock (4.4 kb; stretching;TpAls)+ 18 μL of 1x POP= 20μL of DNA (1:10)
2. 1 μL dna of the above diluted dna (1:10; 4.4 kb; stretching;TpAls) + 9 μL POP(1X) = 10μL dna of (1:100).

This page discussed the general procedure used to make different buffers and solutions. Normally solutions depends on the specific sample, but the buffers;POP and PBS remains similar as discussed.

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