User:Anthony Salvagno/Notebook/Research/2010/07/26/Catching Up and Project Planning

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Wow, my notebook for July sucks. I'm going to have to save it with a stellar week of notes. I gotta refresh myself on things I've done and prepare myself for things to do.


I need to break this down step-by-step and determine what I can do and what it affects and determine the best method. Of course this also depends on how well the new laser works. If we get power up the wazoo then I won't need to use the big beads and we have no problem with the tethers, but if I still need to tether with big beads then I'll have to figure out how to tether them reliably.

Small Bead Tethering

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  1. Clean slides - generally I do this like 50% of the time and see no results that would sway me from doing this or not doing it. We use precleaned slides so sometimes I justify not cleaning, but again I haven't had results that say clean everything all the time.
    • Diego sent me a good cleaning protocol, and I've read about a few others. Most of them involve sonicating in ethanol and then sonicating in water with some rinsing and blow drying.
  2. Dilute Anti-dig 1:10 in PBS
    • The anti-dig starts already diluted in 20ul of PBS but this is an artifact from Koch's method that has been verified in other papers.
  3. Make BGB - 5mg/ml in 1x POP
    • I store this in the fridge for weeks at a time.
  4. Dilute .5um microspheres 1:20 in BGB/POP
  5. Sonicate beads to remove clumps
  6. Make sample chamber - I have a really awesome method for doing this and it takes like 2 min to make a double chamber from scratch.

Flow Cell Technique

The sample chambers I make can hold between 6-10ul so I use 10ul as 1 sample volume

  1. Flow anti-dig and incubate for 5 min
  2. Wash with 50ul of BGB and incubate for 3 min
    • Koch's protocol calls for 2 washes, but I usually do just one since 50ul is generally much larger than 5sv.
  3. Flow DNA and incubate for 10 min
  4. wash with 50ul of BGB
  5. flow beads and incubate for 10 min
  6. wash with 50ul BGB and seal
  7. allow sealant to dry/harden.

Big Bead Problems

Almost nothing from the above protocol should change when tethering with 1um beads, and yet it doesn't work for some reason. Well my best guess at the problem is increased surface interaction between the beads and the glass due to the large bead radius. I've been mostly confirmed on this theory from experimental results but have yet to find a solution. Here are some ideas (and some comments):

  1. dilute beads in high salt BGB
    • high salt seems to get the beads closer, but the tethering efficiency is way too low. Maybe I get 1 tether every 5 field of views.
  2. dilute beads in high salt POP
  3. don't wash after bead incubation
  4. use BGB after anti-dig and use POP for all other washes
  5. try blocking with a-casein
    • I can't tell if it is this step or the next one, but I noticed that beads will stick after about 20min of incubation. I did a sample incubating for 10 min and saw regular floaters and did one incubating for 30 min and they were all stuck
  6. dilute beads with a-casein
  7. dilute beads in water
  8. long incubation times
    • I coupled this with beads in BGB. Most times I would incubate overnight and if I did the experiment during the day I would incubate for an hour at least. The overnighters would produce tethers, but nothing conclusive.

Other notes:

  • I haven't found the need to sonicate big beads. There seems to be much less sticking even after days of sitting in a tube. Gentle agitation with flicks or even pipetting up and down seems to get the job done.
  • My best guess is that the beads get close to .5um from the surface but no closer than that under normal tethering conditions. This is from the fact that tethers break very easily. Also occasionally I see beads stick to tethers, jiggle for a few seconds, and then float away. Maybe that is just the normal lifespan of biotin-streptavidin?
  • I suppose there is a lot of sticking to glass via the BGB. I see what look like tethers and then pull with the tweezers, but I get the normal stuck bead reaction (DOG graph). Like there isn't enough force to stretch or unzip.


We've ordered various parts that should make the tweezer work.

  • New Laser - hopefully it amazingly unzips everything it comes across even if it isn't unzippable and some things that actually are (pant flies anyone?)
  • objective - this was ordered a long time ago but is due when the laser is due
  • piezo - we got a z-piezo and this should make certain things easier especially the focus drift. For some reason I have jewish buyer remorse when I think about this thing.

Pranav and I need to redesign the optical path for the new laser. This new design will have the following features:

  • mapped optics - the beam expander and the beam steering optics will be conjugate planes of the beam waist
  • aom conjugate plane of beam waist
  • better use of space - right now everything is spread out and the laser covers a lot of distance
  • computer controlled stages - we have a lot of work to do on this
  • z-positioning piezo
  • water immersion objective


Koch and I discussed the possibility of altering the unzipping construct for big beads. We found some tethering success with the long 4.4kb tethers for stretching and our discussions revolved around a using this as a possible template. Here are some thoughts:

  • I would put off on this until after the laser and new objective come in. The reason for this is because we might not need to use 1um beads with the new stuff, in which case we would have no problems with the tethering portions (theoretically).
  • I would put the reverse primer just beyond the BstXI cut site and the forward primer 4kb upstream
  • Possibly use a new plasmid template, pBR322 maybe

There are lots of things to consider, and this will take considerable thinking before acting.