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 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.
• 3M double-stick tape (like the kind you can get at Office Max).

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, 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

Blocking solution

We use: 5 mg / ml "Blotting Grade Blocker" from Bio-rad, dissolved in "popping buffer."

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
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 picture of someone changing buffers through the sample chamber: