Koch Lab:Protocols/Microsphere-DNA tethering/Glass, dig, biotin, microsphere, 4kb DNA

Notes

1. I would call this the "Lazy protocol," or "Sufficient protocol." One could probably do better by increasing incubation times, tweaking concentrations, being careful with temperatures, being precise with incubation times, etc. But for ripping through a bunch of samples in one afternoon, it seems to work well.
2. Here is another published protocol (Appleyard et al. American Journal of Physics 75 p. 5) that is similar but slightly different.
3. Some photos I'm going to insert

Materials

Sample chamber

You can use more sophisticated sample chambers, obviously, but our most common method remains the ultra-low-tech "double-stick tape" method. For this you need:

• #1 coverglass (or whatever your microscopy requires). 60 x 24 mm is convenient. You can get Corning from Fisher.
• regular glass slides (or you can use another coverglass). 1 inch x 3 inch, about 1.2 mm thick.
  • See: thoughts on cleaning glass.
• 3M double-stick tape (like the kind you can get at Office Max).

In the picture above, you can see four strips of double-stick tape. We create these by putting a full strip onto a microscope slide and then slicing it with a razor blade. We use curved forceps to peel the tape strips off (pull up so the tape doesn't curl). Static electricity is annoying and makes it tough to make straight channels.

Anti-dig

Polyclonal sheep anti-dig from Roche (Cat. No. 1 333 089). 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 to make:

20 microgram / milliliter working solution of anti-dig
keep cold, do not freeze, store at +4C for up to a few weeks, or until you run out, or until stuff stops working

Popping buffer

This is a buffer we used for unzipping DNA experiments. It's called "popping buffer" because when DNA binding proteins are present they are "popped" off the DNA when it is unzipped.

50 mM Sodium Phosphate, 50 mM NaCl, 10 mM EDTA 0.02% Tween-20

Here is an excel recipe file: Media:010819 PoppingBuffer.xls

Blocking solution

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 differnces in purity or kinds of casein could potentially impact a sensitive assay. Often it is imagined that casein is a regular soluble protein, but Steve 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" Cat# 170-6404 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.")

Solubility is also a big problem. Even though it purportedly forms micelles, they are small micelles, and will stay colloidal. You need the BGB to be soluble in order to work well. You also need the BGB to be soluble to go through a 0.2 micron filter, and if it doesn't you may end up pushing the syringe too hard and spraying BGB all over your and your graduate student's face.

OK, so here is what you want:

5 mg / ml BGB dissolved in Popping Buffer, 0.2 micron filtered
Keep cold, do not freeze, store at +4C in 1 ml working aliquots, use as long as they are working and not growing things.

Here are more notes on how to make it: BGB Prep

DNA

This tethering protocol 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).

• Here's an example DNA construct: two DNA hairpins with complementary overhangs to a digested plasmid are ligated to the plasmid. Dephosphorylation of the 5' ends of the hairpins does not allow them to ligate to each other.

Microspheres

We have had good experiences with these two types of commercial streptavidin microspheres:

• Non-magnetic: CP01F from Bang's Labs. They are approx. half micron diameter, streptavidin coated polymer microspheres, with embedded Dragon Green dye. We have also had good luck with non-dyed streptavidin polymer half micron beads from Bangs. I think these are product numberCP01N.
• Magnetic: Dynabeads M-270 Streptavidin or M-280 from Invitrogen (used to be Dynal). Dynal by far supplied the best streptavidin coated microspheres we have ever used. M-270 and M-280 are about 2.8 micron diameter, and very monodisperse both in terms of diameter and also in maghemite content. We characterized them a bit in our [APL paper http://www.citeulike.org/user/skoch3/article/1085854].

Washing the microspheres

You should "wash" the microspheres to remove 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. Washing is easy with magnetic microspheres. It's a bit more of a hassle with non-magnetic, and you need to pick a good g-force so as to make them easy to resuspend. If you have good information on this, please post it!

Kim wipe wicks

We use twisted Kim wipes as wicks for drawing fluid out of one side of the cell. Steve likes to keep it folded in half, and twist from one corner of the fold.

Optional: Humidity chamber

Use an old pipet tip box. Fill with water to just below the plastic piece that holds the tips. You can set the sample chamber on this and close the lid and it seems to slow down evaporation. It probably also grows things.

Procedure

Create a sample chamber

Details later (it's tough to describe, but easy to learn from someone)

1. Clean glass surfaces
2. Make chambers
   • We find that making a bunch of chambers ahead of time doesn't save that much time, and they may not stay clean, and may losing sticking strength. So we make them on demand.

Form tethers

This assumes you have a typical sample chamber made out of #1 coverglass, double stick tape, and microscope slide (or another coverglass). Assumes sample volume about 10-20 microliters. Assumes glass is already cleaned, and dry.

1. Coat the glass surfaces (and presumably the sticky tape walls) with anti-dig. The sticking is non-specific, and presumably some antibodies will denature or not be able to bind dig. The fraction that stick is unknown.
   • Because the sample is dry (and clean/hydrophilic), the liquid will flow in without the need for a wick on the opposite side. If it doesn't flow in easily, and bubbles form, it's probable that your glass isn't clean enough.
   • During this step, measure the volume of your sample chamber, which will be determined by the width of the channel. You can make a good guess by eye, dial it in on your pipetman, and then make a good estimate based on how under- or over-filled it gets. From here on out, we will call this the "sample volume" or s.v.
   • Flow in 1+ s.v. of 20 microgram/ml antidig. (By 1+, I mean let the liquid pool on the sides, so that evaporation won't be too much of a problem over 5 or 10 minutes.
   • Let sit at room temperature for 5 to 10 minutes.
     • Optional: let sit in a "humidity chamber" to minimize evaporation. This can be constructed as mentioned above.
2. Optional: If you are worried about unbound anti-dig in later steps, and don't want to waste blocking solution. Wash away un-bound antidig with PBS
   • Flow through 5 s.v. of PBS buffer. (Draw liquid through with kim wipe wick.) Repeat if you want to.
3. Block glass surface with casein. This is one of the troublesome things, because everyone says "casein" and there are a whole slew of different casein products out there, and Sigma stopped selling one of the popular ones. See "Materials" above for discussion. The purpose of this step is to prevent DNA or microspheres from sticking to glass that wasn't blocked with the relatively small amount of anti-dig.
   • Flow through 1.5 s.v. of blocking solution (see above; often 5 mg / ml BGB in popping buffer). (Again, leave solution piled up on edges to deal with evaporation.)
   • Incubate for 2 - 5 minutes (usually 2 minutes) at room temperature.
   • Repeat for total of 3 times. Waiting longer on the final step would make more sense, if casein solution is being depleted by sticking to walls.
   • Variation If you have a lot of blocking solution, you can skip the above optional washing step, and use 5 s.v. of blocking solution here, repeating 3 times.
4. Bind DNA to surfaces via dig/anti-dig bonding.
   • Flow in 1.5 s.v. of DNA solution (typically 20 picomolar dig, biotin labeled dsDNA in buffer or blocking solution).
   • Incubate at room temperature for 5 to 10 minutes.
5. Wash away free DNA (optional if you think DNA binding is complete). The goal is to prevent free DNA from sticking to beads in solution, but this is probably not a problem.
   • Flow through 5 s.v. of buffer or blocking solution
   • Repeat for a total of two times
6. Bind microspheres
   • Flow through 1.5 s.v of microspheres (concentration? current guess: 1:10 dilution of stock beads, diluted in popping buffer).
   • Tethering can begin immediately, if you have a dilute solution of microspheres, you can observe before the next step (washing away free beads).
   • Incubate for as long as you wish...10 minutes at room temperature is sufficient usually.
7. Wash away free beads
   • Flow through 5 s.v. of buffer.
   • Repeat for total of 2 times.
8. Seal sample, if desired
   • Regular nail polish works well
     

Here is a sample tethering experiment recipe:
{{#widget:Google Spreadsheet |key=t70tuJVfe2-TSghF7uwTUKg |width=600 |height=300 }} Here is a picture of someone changing buffers through the sample chamber:

Observation

Here's a cartoon of the tethering setup (NOT DRAWN TO SCALE).

It includes the DNA, streptavidin-coated fluorescent microspheres, anti-dig antibodies, dig molecules, biotin molecules, and casein blocking agent (in peach).