Koch Lab:Data/Steve grad data set notes (partial)
010326.txt
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0025.dat Contains example of 0.35 um peeler. Otherwise mostly junk
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0026.dat Contains example of 0.53 um in BsaIBuffer
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0027.dat Pre-BsaI, shows some examples of artifactual data
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0028.dat Some Pre-BsaI, no good data
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0029.dat Trying to gather Pre-BsaI data. Large File.
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0030.dat BsaI--contains one good template
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0031.dat BsaI--really contains no good templates
010327.txt
010327
|=============================================================
Step 0 : Peel Buffer + Tween ; 5.000000 s.v.; 0.000000 min.
Step 1 : Peel Buffer ; 5.000000 s.v.; 0.000000 min.
Step 2 : antidig 20 ug / ml in PeelBuf ; 1.000000 s.v.; 11.000000 min.
Step 3 : BGB (old aliquots) ; 1.000000 s.v.; 5.000000 min.
Step 4 : BGB (repeat) ; 1.000000 s.v.; 5.000000 min.
Step 5 : PeelBuf ; 5.000000 s.v.; 0.000000 min.
Step 6 : 17-mer (most recent batch, 1:10 in PBS, 1:3 in PeelBuf) ; 1.000000 s.v.; 10.000000 min.
Step 7 : EcoRI Bufferf* ; 5.000000 s.v.; 0.000000 min.
Step 8 : Beads, Bangs 0.53 micron streptavidin, 1:10 into EcoRI Buffer* ; 1.000000 s.v.; 20.000000 min.
Step 9 : EcoRI Buffer* ; 10.000000 s.v.; 0.000000 min.
Step 10 : *1 sample with tween, the other without ; 0.000000 s.v.; 0.000000 min.
|=============================================================
Non-tween sample has lots of stuck beads (about 20 or more per octagon, and less than 1 tether)
Tween sample had less stuck beads. Had more things that looked like tethers, but nothing peeled! FUUCK!
Also, there is some kind of problem with the oscilloscope DAQ board, and channel-to-channel interference...probably a grounding issue
All of the data are fucking worthless
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010327\0032.dat non-tween sample. both segments are garbage. sample sucks
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010327\0033.dat tween sample...stopped early to change conditions
010328.txt
010328
|=============================================================
Step 0 : Dopes ; 5.000000 s.v.; 0.000000 min.
Step 1 : PBS ; 5.000000 s.v.; 0.000000 min.
Step 2 : antidig 20 ug / ml in PBS ; 1.000000 s.v.; 10.000000 min.
Step 3 : BGB (old aliquots) ; 1.000000 s.v.; 5.000000 min.
Step 4 : BGB (repeat) ; 1.000000 s.v.; 15.000000 min.
Step 5 : PBS ; 5.000000 s.v.; 0.000000 min.
Step 6 : 17-mer (1:10 aliquot) ; 1.000000 s.v.; 10.000000 min.
Step 7 : PBS ; 5.000000 s.v.; 0.000000 min.
Step 8 : Bangs 0.53 Streptavidin 1:10 in Dopes ; 1.000000 s.v.; 20.000000 min.
Step 9 : Dopes ; 10.000000 s.v.; 0.000000 min.
Step 10 : Delay..maybe 1/2 hour ; 0.000000 s.v.; 0.000000 min.
Step 11 : 1%BsaI in 90% dopes + 10 mM EDTA ; 5.000000 s.v.; 0.000000 min.
|=============================================================
After 10 minutes of bead incubation, before washing beads away, there are about 1 or two possible tethers per 90 micron octagon. Since I am using 3x more 17-mer than I was yesterday, this could partially explain my difficulty.
Also, I notice that the bangs beads are very clumpy. This could be a factor in reducing the tether density.
After washing away beads, seems to be about 3 beads or whatever. But I notice that they transiently stick--this is also a problem.
The sample glass is actually fairly dirty.
Added BsaI in 90%Dopes + 10 mM EDTA at 11:42PM
VIDEO of little dirt farm 12:33 AM
SO, Here is my guess as to what the secrets were to tonight:
A. Use higher DNA concentration...makes a big deal compared to background stuck beads?
B. Revert to old buffers. Same consideration as A, perhaps sticking more of a problem with the other buffers.
C. Do I have DNAse or other contamination??? It appeared to me that the tethers were going away with time, and becoming more prone to breaking. After loading in the BsaI and EDTA buffer, the problem seemed to go away, and I got lots of good data.
Also, today, I fixed a bug related to offset subtraction.
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010327\0034.dat I think wiggle array messed up
In this file, segment 006, I took video on KochVII, which shows how a peeler looks as a tether. In addition, this video shows my z=8.5 focusing method. Right after that on video is example of transient sticking problem. Even though segment 6 and 7 break soon, I think the vidoe is a good example
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010327\0035.dat I think wiggle array messed up
Some great BsaI popping. I would say about 1/3 were good. Of those, for some reason they still seemed to break early
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010327\0036.dat Just one segment, which shows that even after all of this time, there are still BsaI poppers.
I HAVE FIXED THE OFFSET SUBTRACTION FOR THE FIND TETHER CENTER NOW>
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010327\0037.dat Has some really great extremely fast pulling (where the force may indeed be incrementally higher)
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010327\0038.dat Slow popper in segment 004. Forces were all over the board (high, low). Should stare at this one for a while
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010327\0039.dat One single fast popper?
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010327\0040.dat at least four templates with extremely fast popping (50 MHz / sec). this seemed pretty awesome, but I guess you needed to be there.
010329.txt
3/29/01 2:55 AM
|=============================================================
Step 0 : Dopes ; 1.000000 s.v.; 10.000000 min.
Step 1 : 0.9xPBS 10 mM EDTA ; 5.000000 s.v.; 0.000000 min.
Step 2 : antidig 20 ug / ml in 0.9xPBS + 10 mM EDTA ; 1.000000 s.v.; 11.000000 min.
Step 3 : 10 mg / ml BGB in 0.45x PBS 5 mM EDTA ; 2.000000 s.v.; 9.000000 min.
Step 4 : 10 mg / ml BGB in 0.45x PBS 5 mM EDTA ; 2.000000 s.v.; 5.000000 min.
Step 5 : 17-mer 1:11 in 10/11 PBS + 10 mM EDTA ; 1.000000 s.v.; 14.000000 min.
Step 6 : Bangs 0.53 streptavidin 1:10 in final buffer 0.81 PBS 9 mM EDTA ; 1.000000 s.v.; 15.000000 min.
Step 7 : 0.9x Dopes + 10 mM EDTA ; 10.000000 s.v.; 0.000000 min.
Step 8 : 0.9x Dopes + 10 mM EDTA ; 10.000000 s.v.; 0.000000 min.
Step 9 : 0.9x Dopes + 10 mM EDTA ; 10.000000 s.v.; 0.000000 min.
Step 10 : 0.9x Dopes + 10 mM EDTA + 1:50 EcoRI at about 2:06 AM ; 5.000000 s.v.; 0.000000 min.
|=============================================================
after first bead wash-away, there were 1.1 "look-like" tethers per 90 micron octagon. about the same or less stuck beads (that is, stuck beads are lower than before, most probably because of the higher casein concentration)
Given the about 30% success rate, I would say actual density is about 0.3 per octagon, which is definitely very low.
after second wash, SEEMS like fewer tethers. i counted 0.8 per octagon versus 1.6 stuck. however, peelers efficiency increases!
2:06 AM, Flowed in EcoRI in 0.9x Dopes + 10 mM EDTA (1 ul enzyme in 50 ul buffer, flowed in 50 ul)
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0041.dat initial file. shows maybe 30% success rate
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0042.dat demonstrates strange increase in peeling ability by washing a second time!
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0043.dat 0.5 MHz / sec
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0044.dat 5 MHz / sec
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0045.dat 10 MHz / sec
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0046.dat 20 MHz / sec
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0047.dat 50 MHz / sec
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0048.dat free bead, maybe not configured correctly
BAD OFFSET SUBTRACTION
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0049.dat free bead 50 MHz/ sec
BAD OFFSET SUBTRACTION
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0050.dat 50 MHz / sec no bead...messed up, though
BAD OFFSET SUBTRACTION
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0051.dat 0.5 MHz / sec no bead
BAD OFFSET SUBTRACTION
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0052.dat 50 MHz / sec no bead
BAD OFFSET SUBTRACTION
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0053.dat 50 MHz / sec free bead
BAD OFFSET SUBTRACTION
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0054.dat 50 MHz / sec fixed offset correction
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0055.dat 50 MHz / sec fixed offset correction
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0056.dat 0.5 MHz / sec EcoRI...lots of popping, but seeming no full-length
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0057.dat 5 MHz / sec EcoRI pops
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0058.dat 10 MHz / sec EcoRI pops
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0059.dat 20 MHz / sec
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010329\0060.dat 50 MHz / sec
010404.txt
4/4/01 3:31 AM
|=============================================================
Step 0 : Popping Buffer(04/03/2001) ; 5.000000 s.v.; 0.000000 min.
Step 1 : Repeat ; 5.000000 s.v.; 0.000000 min.
Step 2 : Antidig (Roche) 20 ug / ml diluted in Popping buffer ; 1.000000 s.v.; 10.000000 min.
Step 3 : 10 mg / ml BGB in 0.5 PBS + 5 mM EDTA ; 1.000000 s.v.; 10.000000 min.
Step 4 : repeat ; 1.000000 s.v.; 10.000000 min.
Step 5 : Popping Buffer ; 5.000000 s.v.; 0.000000 min.
Step 6 : 1:10 17-mer (+1mM EDTA today) in PBS ; 1.000000 s.v.; 10.000000 min.
Step 7 : Popping Buffer ; 5.000000 s.v.; 0.000000 min.
Step 8 : Bangs 0.53 1:10 into Popping Buffer ; 1.000000 s.v.; 20.000000 min.
Step 9 : Popping Buffer (about 1AM) ; 10.000000 s.v.; 0.000000 min.
Step 10 : delay ; 0.000000 s.v.; 83.000000 min.
Step 11 : EcoRI, 1:100 into Popping Buffer (2:23 AM) ; 5.000000 s.v.; 0.000000 min.
Step 12 : EcoRI, 1:100 (same as last batch) at about 3:04 AM ; 5.000000 s.v.; 0.000000 min.
Step 13 : 1:5 EcoRI into popping buffer 3:14 AM ; 2.000000 s.v.; 0.000000 min.
Step 14 : Plain Popping Buffer (no enzyme) 3:22AM, ; 5.000000 s.v.; 0.000000 min.
|=============================================================
Had programming emergency for first hour. Figured out that it was a Splice12-segments(categorical) error. Resumed taking data.
15 tethers, selected by eye. 33% broke immediately, 4/15 peeled completely, and 9/15 peeled at least partially. So, I think 25% expectation for restriction enzymes is reasonable. Will now (2AM) add enzyme
Any secrets?
I think flow force may help lift good tethers off the surface. So that routine washing is a good thing.
For some reason I cannot get EcoRI to saturate
0062.dat 15 per-enzyme tethers for examples
0063.dat 0.5 MHz / sec
0064.dat 5 MHz / sec (lots of overstretching)
0065.dat 0.05 MHz / sec, several templates with popping
0066.dat 0.5 MHz / sec again, except this time surely at "room" temperature
0067.dat 0.5 MHz / sec again, with freshly added room temperature enzyme. LOTS of popping. THEORY: the flow force "rejuvinates" some stuck beads that are perfectly good tethers.
0068.dat 0.5 MHz / sec after adding 1:5 EcoRI
0069.dat same as previous
0070.dat 0.5MHz/sec after washing with plain popping buffer
010405.txt
4/5/01 12:47 AM
|=============================================================
Step 0 : Popping Buffer, room temp ; 5.000000 s.v.; 0.000000 min.
Step 1 : Antidig 20 ug / ml (1:10 in popping buffer) ; 2.000000 s.v.; 10.000000 min.
Step 2 : BGB, same as yesterday ; 2.000000 s.v.; 10.000000 min.
Step 3 : Pop Buffer ; 5.000000 s.v.; 0.000000 min.
Step 4 : BGB ; 2.000000 s.v.; 10.000000 min.
Step 5 : 17-mer +EDTA (see yesterday) ; 1.000000 s.v.; 10.000000 min.
Step 6 : Pop Buffer ; 10.000000 s.v.; 0.000000 min.
Step 7 : Bangs 0.53 streptavidin 1:10 into Pop Buffer (Diluted yesterday...when flowing in, the color is non-homogenous, probably indicating clumping from the buffer?) ; 1.000000 s.v.; 10.000000 min.
Step 8 : repeat ; 1.000000 s.v.; 10.000000 min.
Step 9 : Pop Buff ; 10.000000 s.v.; 0.000000 min.
Step 10 : BsaI 1:25 into Popping Buffer ; 1.000000 s.v.; 2.000000 min.
Step 11 : Repeat above 5 times (ended about midnight) ; 0.000000 s.v.; 0.000000 min.
Step 12 : seal nail polish ; 0.000000 s.v.; 0.000000 min.
|=============================================================
tether density is very low (less than 1/2 per 90 micron)
I believe even more in the theory that good tethers transiently stick to the surface (I even grabbed one off and peeled it perfectly)
0071.dat 0.5 MHz / sec, lots of popping, and perhaps full poppers
0072.dat same as previous. lots of segments shows the problem with biased force
0073.dat 5 MHz / sec, lots of popping
0074.dat same as previous
0075.dat 10 MHz / sec Done for the night
010509.txt
5/9/01 2:23 AM
|=============================================================
Step 0 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 1 : PBS ; 5.000000 s.v.; 0.000000 min.
Step 2 : antidig (Roche) 20 ug / ml in PBS ; 2.000000 s.v.; 7.000000 min.
Step 3 : BGB 10 mg / ml in ? (same as last time) ; 2.000000 s.v.; 10.000000 min.
Step 4 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 5 : DNA [either pRL574 peeling template from december, or 17 pM dsDNA PCR) ; 1.000000 s.v.; 14.000000 min.
Step 6 : Dopes (ran out of thawed popBuf) ; 5.000000 s.v.; 0.000000 min.
Step 7 : Bangs 1:10 in PopBuf 0.45 micron streptavidin, freshly diluted ; 1.000000 s.v.; 22.000000 min.
Step 8 : PopBuf ; 10.000000 s.v.; 40.000000 min.
Step 9 : EcoRI, 1:50 in PopBuf (LifeTech) (about 11:43PM) ; 2.000000 s.v.; 20.000000 min.
Step 10 : EcoRI, 1:50 in PopBuf (LifeTech) (about 12:04PM) ; 2.000000 s.v.; 26.000000 min.
Step 11 : EcoRI, 1:5 in PopBuf (LifeTech) (about 12:30PM) ; 0.000000 s.v.; 0.000000 min.
|=============================================================
Data set 082, segment 005 through xxx, I was in some weird tether farm in the middle of the sample. It showed lots of wiggly beads (at least 10x higher conc. than other areas) and high probability of non-specific tethering, or double peeling tethering
12:30 AM added 1:5 Eco in PopBUf
also tonight, made 4 molal sucrose by adding .4 moles of sucrose to 100 ml of H2O (ends up about 200 ml). i then made 2 molal sucrose 50% popping buffer by mixing equal volumes.
0079
0080 EcoRI...peeling and some popping?
0081 EcoRI...peeling and some popping?
0082 0.5 MHz / s
freshly added Eco after 1st segment
0083 5 MHz / s
0084 0.5 Mhz / s
10x more EcoRI (1:5)
THIS IS THE BEST FILE YET...at least 3 popping events, which look to be at the correct site
0085 5 Mhz / s
EcoRI 1:5
This is an awesome data file!
NOTE: Mysterious fluctuations...possibly overstretch related...examine the shearing sequence for possible EcoRI binding...this gave me idea for using shearing to study protein binding!
0086 4.4 kb double tagged template in popBUf only. EVERYthing seems to break right at overstretch
0087 just for fun, added EcoRI 1:5...has a few overstretches
0088 overstretchin in 2 Molal sucrose in 50% popping buffer
0105092.txt
5/9/01 5:49 AM
3:10 added the sucrose
about 3:38 AM added 1:20 BsaI into 45% pop + 2 m Sucrose (45 ul popBuf + 50 ul 4 m Sucrose + 5 ul BsaI)
4:10 AM 1 m + 1:20 BSAI
4:29 AM 0.5 m + 1:20 BSAI
4:52 AM 0.25 m + 1:20 BsaI
5:12 AM 0.125 m + 1:20 BsaI
5:22 AM 0.0625 m + etc.
5:36AM 1:20 BsaI in popBuf (NO SUCROSE)
0089 0.5 MHz / sec
naked 17-mer in popBuf
0090 0.5 Mhz / sec
naked 17-mer in 2 m Sucrose
0091 0.05 Mhz / sec
naked 17-mer in 2 m Sucrose
0092 5 Mhz / sec
naked 17-mer in 2 m Sucrose
0093 0.5Mhz / sec
BsaI...lot's of overstretching!!!!
0094 0.05Mhz /sec
BsaI...still overstretching, but this time popping too!
0095 0.005 MHz / sec
BsaI...now down to reasonable forces...i think these three previous files prove that the sucrose is drastically increasing binding strength
0096 0.05 MHz / sec
BsaI in 1 m Sucrose
0097 0.005 Mhz / sec
BsaI in 1 m Sucrose
0098 0.5MHz / sec
BsaI in 1 m Sucrose
0099 0.5 MHz / sec
BsaI in 0.5 m Sucrose
0100 0.05 MHz / sec
BsaI in 0.5 m Sucrose
0101 0.005 MHz / sec
BsaI in 0.5 m Sucrose
I am now getting distressed that I am seeing too much weird shit...i think part of the stuff i have been seeing may have to do with multiple tethering?
0102 0.5 MHz / sec
BsaI in 0.25 m Sucrose
0103 0.05 MHz / sec
BsaI in 0.25 m Sucrose
0104 0.5 MHz / sec
BsaI in 0.125 m Sucrose
0105 0.05 MHz / sec
BsaI in 0.125 m Sucrose
0106 0.5 MHz / sec
BsaI in 0.0625 m Sucrose
0107 0.05 MHz / sec
BsaI in 0.0625 m Sucrose
0108 0.5 MHz, BsaI only (no sucrose)
0109 0.05 MHz, BsaI only (no sucrose)
010510.txt
5/10/01 3:31 AM
|=============================================================
Step 0 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 1 : PBS ; 5.000000 s.v.; 0.000000 min.
Step 2 : Antidig from yesterday (20 ug / ml) ; 2.000000 s.v.; 5.000000 min.
Step 3 : BGB 10 mg / ml (same tube at +4C for weeks) ; 2.000000 s.v.; 10.000000 min.
Step 4 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 5 : 17-mer, 1:5 dilution of freezer stock in PopBuf ; 1.000000 s.v.; 12.000000 min.
Step 6 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 7 : Bangs 1:10 in PopBuf ; 1.000000 s.v.; 35.000000 min.
Step 8 : PopBuf ; 10.000000 s.v.; 0.000000 min.
Step 9 : see data notes for rest of protocol (BsaI and washes) ; 0.000000 s.v.; 0.000000 min.
|=============================================================
1:05 AM flowed in 1:20 BsaI 2 s.v. in PopBuf
2:17 AM 5 s.v. popping buffer
2:27 another 5 s.v.
2:32 another 5 s.v.
3:25 AM another 5 s.v. after waiting this 1 hour
0111 0.5 MHz / s
Naked 17-mer
6 out of 14 (43% showed peeling) but only 3 / 14 (21%) showed complete peeling. Only 1 (7%) showed overstretch behavior
0112 0.5 Mhz / s
BsaI 1:20 popping buffer
First two segments STILL show overstretching. IDEA: Last night and tonight I have NOT turned off the room fan. This would lower the temperature of the sample, and perhaps affect the binding energy this significantly?
This overstretching went away after a few minutes and overall, I had lots of popping (very high percentage success). This is consistent with previous observations that fresh washes tend to revive good tethers.
0113 0.05 MHz / s
BsaI 1:20 popping buffer
4 segments, first and last show very nice low force popping
0114 1.85 V Force Clamped Pop
BsaI 1:20 popping buffer
segment is likely huge
0115 higher force than last file, still doesn't budge
0116 2.1 V force clamped
BsaI 1:20 popping buffer
higher force than last file, finally budges
this file has lots of great force clamp results
0117 2.2 V FC
BsaI 1:20 popping buffer
0118 0.05MHz/sec
BsaI 1:20 popping buffer
to show that there is still enzyme binding
0119 0.5Mhz / sec
BsaI after 5 s.v. wash
0120-0122, after 10 s.v., 15 s.v., 20 s.v. 0.5MHz / sec
0123 0.5Mhz / sec
After waiting an hour even...these guys just won't go away
0105102.txt
5/10/01 5:39 AM
|=============================================================
Step 0 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 1 : PBS ; 5.000000 s.v.; 0.000000 min.
Step 2 : Antidig from yesterday (20 ug / ml) ; 2.000000 s.v.; 4.000000 min.
Step 3 : BGB 10 mg / ml (same tube at +4C for weeks) ; 2.000000 s.v.; 12.000000 min.
Step 4 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 5 : 17-mer, 1:5 dilution of freezer stock in PopBuf ; 1.000000 s.v.; 11.000000 min.
Step 6 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 7 : Bangs 1:10 in PopBuf ; 1.000000 s.v.; 25.000000 min.
Step 8 : PopBuf ; 10.000000 s.v.; 0.000000 min.
Step 9 : see data notes for rest of protocol (BsaI and washes) ; 0.000000 s.v.; 0.000000 min.
|=============================================================
4:11 2 S.V. of BsaI 1:20,000 (this is the second so far)
4:19 AM, 2 s.v. BsaI 1:5,000
4:39 AM 2 s.v. BsaI 1:2,000
4:57 2 s.v. more of 1:2,000
5:05 2 s.v. 1:500
0124 0.5MHz / sec
BsaI 1:20,000, 2 s.v.
only 1 segment has pops
0125 0.05MHz / sec
BsaI 1:20,000 2 s.v. so far
0126 0.05MHz / sec
BsaI 1:20,000 4 s.v. so far
0127 0.05MHz / sec
BsaI 1:5,000 2 s.v.
0128 0.5MHz / sec
BsaI 1:5,000 2 s.v.
I am noticing a propensity to stick right at the END of template
0129 5 MHz / sec
BsaI 1:5,000 2 .s.v.
There are three complete peeler segments, which make me feel comfortable that nothing too freaky is going on (I hope)
0130 5 MHz / sec
BsaI 1:2,000 2 .s.v.
0131 0.5 MHz / sec
BsaI 1:2,000 2 .s.v.
0132 0.05 MHz / sec
BsaI 1:2,000 2 .s.v.
0133 0.05 MHz / sec
BsaI 1:2,000 2 .s.v. (second wash)
0134 0.5 Mhz / sec
BsaI 1:500 2 s.v.
0135 0.05 MHz / sec
BsaI 1:500 2 s.v.
010513.txt
5/13/01 6:24 AM
|=============================================================
Step 0 : Note, the sample was rather small, probably 80% of what i call s.v. below ; 0.000000 s.v.; 0.000000 min.
Step 1 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 2 : PBS ; 5.000000 s.v.; 0.000000 min.
Step 3 : antidig, thawed last week ; 2.000000 s.v.; 4.000000 min.
Step 4 : 10 mg / ml BGB (same as last week...is very old) ; 2.000000 s.v.; 1.250000 min.
Step 5 : repeat above 2 times ; 0.000000 s.v.; 0.000000 min.
Step 6 : 17-mer, thawed and diluted 1:5 last week ; 1.000000 s.v.; 6.000000 min.
Step 7 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 8 : Bangs 1:10 in PopBuf, diluted last week or earlier ; 1.000000 s.v.; 5.000000 min.
Step 9 : repeat above once ; 0.000000 s.v.; 0.000000 min.
Step 10 : Bangs 1:10 freshly into PopBuf ; 1.000000 s.v.; 5.000000 min.
Step 11 : Repeat above once (a total of 4 bead washes) ; 0.000000 s.v.; 0.000000 min.
Step 12 : PopBuf ; 10.000000 s.v.; 0.000000 min.
Step 13 : BsaI 1:2,000 in PopBuf ; 1.000000 s.v.; 0.000000 min.
|=============================================================
Second protocol:
|=============================================================
Step 0 : wash glass with water and kim wipes ; 0.000000 s.v.; 0.000000 min.
Step 1 : antidig 20 ug / ml freshly diluted ; 2.000000 s.v.; 9.000000 min.
Step 2 : BGB 10 mg / ml ; 2.000000 s.v.; 9.000000 min.
Step 3 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 4 : 17-mer freshly diluted 1:5 ; 1.000000 s.v.; 8.000000 min.
Step 5 : popBuf ; 5.000000 s.v.; 0.000000 min.
Step 6 : Bangs 1:10 PopBuf (fresh today) ; 1.000000 s.v.; 10.000000 min.
Step 7 : repeat last step 1 time ; 0.000000 s.v.; 0.000000 min.
Step 8 : PopBuf ; 10.000000 s.v.; 0.000000 min.
Step 9 : BsaI, 1:2,000 in PopBuf ; 2.000000 s.v.; 0.000000 min.
|=============================================================
5:23 AM BsaI 1:2,000 2 s.v.
0136 Naked DNA...the protocol did not produce very good tethers. Lots of non-specific jiggly beads...but I can find just enough peelers that I suppose it's worth trying the 1:2000 BsaI next file
0137 trying force clamp on naked DNA...abandoning sample after this
BEGIN NEW SAMPLE
0138
0139 Lots of popping...unfortunately, looks like I again have too much enzyme...or at least as time goes on?
0140 more of the same
0141 the first segment in this file is the same tether as last segment of last file. the popping is gone, but still pins at the same site
010704.txt
7/4/01 6:21 AM
|=============================================================
Step 0 : Begin with Bert's specially cleaned coverglass and slide, no further pre-rinsing ; 0.000000 s.v.; 0.000000 min.
Step 1 : Antidig 20 ug / ml in Popping Buffer ; 1.000000 s.v.; 5.000000 min.
Step 2 : BGB, 10 mg / ml in 50% popping buffer. Note, this BGB is maybe 1 or 2 months old at +4C...However, it has been left out at room temperature 2 or 3 times for over 24 hours each time. Oh well, I'm not going to remake it now. ; 5.000000 s.v.; 5.000000 min.
Step 3 : PopBuff ; 5.000000 s.v.; 0.000000 min.
Step 4 : New 17-mer peeling template, estimated 29 pM (Diluted 1:10 from stock into Popping Buffer) ; 1.000000 s.v.; 8.000000 min.
Step 5 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 6 : Bangs, freshly diluted 1:10 into popping buffer, and then sonicated (100 ul volume) for about 1 minutes by hand ; 1.000000 s.v.; 8.000000 min.
|=============================================================
5:47 added BsaI, 5 s.v. 1:50 in PopBuf
Tethering stats:
stuck: N/A ; 19 ; 20 ; 16; 25
jiggle; 19 ; 17; 10; 9; 13 (see transient stickign)
0165 naked DNA, 0.05 MHz / sec
0166 0.05MHz / s
single tether, lots of data, forgot to increase the speed
(but is a double tether)
0167 0.5 MHz / s naked
SHould be several good peelers for comparison
0168 naked
peeled a long way, and then reversed. repeated same tether
0169 Bsai added...a reversal file
0170 BsaI added...quick reversals...a few good ones to examine
0171 BsaI, reversing before a pop (waiting 4,000 data points)
0172 some more like the previous
010705.txt
7/5/01 11:58 AM
|=============================================================
Step 0 : Begin with Bert's specially cleaned coverglass and slide ; 0.000000 s.v.; 0.000000 min.
Step 1 : Antidig (same as two days ago) in Pop BUf ; 1.000000 s.v.; 5.000000 min.
Step 2 : BGB (see notes from two days ago) ; 2.000000 s.v.; 5.000000 min.
Step 3 : PopBUf ; 5.000000 s.v.; 0.000000 min.
Step 4 : Today's Ligation reactions (either HindIII or XhoI), 1:2 into Pop Buf (unpurified ligation reaction) ; 1.000000 s.v.; 6.000000 min.
Step 5 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 6 : Bangs freshly 1:10 into PopBuf, sonicated for roughly 1 minute (100 ul volume) ; 1.000000 s.v.; 8.000000 min.
Step 7 : PopBuf ; 10.000000 s.v.; 0.000000 min.
|=============================================================
0173 0.5 MHz / s, HindIII cap
only 1 out of 18 segments showed the cap...not so good
0174 0.5 MHz / s, XhoI cap
Much better than HindIII cap...about 50% show capping (6 out of 13)
0175 0.5 Mhz / s XhoI cap, reversing at 0.05MHz / s
only first segment is good.
0176 0.5 Mhz / s XhoI...a couple good reversals
0177 0.5 Mhz / s HindIII...no good segments, really, but perhaps is a couple capped (but no reversal caps)
010710.txt
7/10/01 3:54 PM
Super-fast tethering protocol (no wait times: antidig; 5 s.v. BGB; 5 s.v. Pop; 1 s.v. DNA (30 pM, new 17-mer); 5 s.v. Pop; 1 s.v. Beads (about 3-5 minutes); 10 s.v. Pop, observe:
90 micron octagon fields:
stuck: 3;0;2;2;3;5;9;3;6;5
wiggle 0;2;0;2;5;5;7;1;0;3
added XhoI at 2:24 AM
197 trying to focus exactly on tether. This is producing stuck beads. This is either (A) because I changed the nomarski compenation and am now lower than previously tonight or (B) the sample is stickier due to the fast application of BGB...Actually, I just checked, and it seems that my eye sees "in focus" much higher when the compensation is set for zero (when the bead image is symmetric, as opposed to the usual 3-D thing). therefore, I will switch back to this method in next file
198 z=9.5, shows no evidence of sticking
199 still playing with z height
200 still playing with z height
201 z=10.25, first segment looks perfect
202 one peeler after adding fresh beads
203 0.5 MHz / sec Added XhoI in 1:50 Pop Buffer
Are there hints of force-mediated cutting?
204 0.05 MHz / s XhoI
0107102.txt
7/10/01 3:55 PM
tethering before adding any ligase buffer:
looking at TV screens (which are smaller than the actual video)
stuck: 0; 1; 0; 1; 1;1;0;0;1;0
wiggle: 3;0;3;1;3;4;3;2;1;2
Added 2 s.v. T4 DNA LIgase in LIgase buffer (no DNA, though) at 6:48 PM...Tethers did not stick immediately.\
TV screens:
stuck: 2;0;1;1;1;2;0;1;0;0
wiggle:2;1;0;3;1;2;5;5;1
7PM added all components
8PM added: 2 s.v. (1 ul T4 DNA ligase--progema; 2 ul 10x Ligase buffer; 2 ul pCP681 EarI sticky from Jan. 27, 2001; 15 ul h2o)
11 PM, flowed 10 s.v. of 1x Ligase BUffer (Promega) through what I'll call sample "B" (made at the same time as A, but haven't added anything after the bead wash at 7PM). File 190 will be a pre-ligation reaction control in the 1x Ligase buffer
12:25 AM flowed more beads into sample A and washed out after 10 minutes
178 anchoring
179 Added s.v. T4 DNA ligase (no DNA) in 1x Ligase buffer and also 0.02% Tween-20.
This file should have good statistics for control (don't really see much difference, except 20% of tethers may have higher shearing / peeling force)
180 added complete reaction at 7PM...this file shows no peeling in the first 5 minutes
181 no peeling after 10 minutes
182 no peeling after 15 minutes
183 no peeling after 1/2 hour
184 no peeling after 45 minutes...i give up
185 still no peeling
186 first 5 minutes after adding second reaction (8PM)
shows two peelers
187 10 minutes after reaction...no peelers
188 ?
189 10 PM, having left reaction going for 2 hours. See 2 complete peelers out of 24 segments, for about 8% efficiency. Not great, but not terrible.
190 control sample B
191 Added P18-cap-High LIgation reaction..no clear peelers
192 After about 25 minutes, there is still no good evidence of successful ligation (even though cap concentration is extremely high (10 micromolar)
193 Some more of the same
195 Took 15 segments back in Sample A, and there were NO peelers. THis is pathetic
196 Sample A, after adding fresh beads, and in fact getting plenty of loose tethers. Only the last segmetn (out of 16) peels, showing pitiful ligation efficiency. There could be something wrong with the method
010710_2.txt
7/10/01 9:04 PM
Used Richard's coverglass, and tethering is not at all as good.
about 2 or 3 times as many stuck beads as wigglers. about 5 wigglers per 90 micron octagon
4:24 PM, XhoI 1:200 in Pop Buff, 2 s.v.
205 Pre-XhoI, 0.5 MHz / sec, 30 tethers
206 XhoI, 0.5MHz / s, 30 tethers
207 XhoI, 5 MHz / s, 30 tethers
208 XhoI, 0.005 Mhz / s (VERY SLOW) 18 tethers
Watching the oscilloscope, I notice that at the very beginning, there are very many kinds of flip-flopping events (time scales and length scales) it would be interesting to take non-decimated early data...
209 XhoI, 0.05 Mhz / s 30 tethers
I would say sample is degrading
210
211 XhoI, after washing many sample volumes(15 to 20) with popping buffer...lots of breaking, probably no popping
212 XhoI washed, 0.5MHz / s 10 segments
After having washed with 15 s.v. of 1 mM Tris + 0.01% Tween, and the 5 s.v. PopBuf
There is still evidence of popping at the early non-specific site, and things are still breaking
213 XhoI washed with many buffers (finally with 2 M NaCl), 0.5 MHz / s, 30 segments
010711.txt
7/11/01 5:40 AM
|=============================================================
Step 0 : Start with Karen's (stolen from Richard, cleaned by Brent) slides...not so clean ; 0.000000 s.v.; 0.000000 min.
Step 1 : antidig 20 ug / ml in Pop Buf ; 1.000000 s.v.; 3.000000 min.
Step 2 : BGB ; 2.000000 s.v.; 7.000000 min.
Step 3 : Pop Buf ; 5.000000 s.v.; 0.000000 min.
Step 4 : New 17-mer, 1:2 (already 1:10 earlier) in Pop Buf, estimated 15 pM ; 1.000000 s.v.; 3.000000 min.
Step 5 : Pop Buf ; 5.000000 s.v.; 0.000000 min.
Step 6 : Bangs 1:10 Pop Buf, sonicated earlier today ; 1.000000 s.v.; 10.000000 min.
Step 7 : Pop Buf ; 10.000000 s.v.; 0.000000 min.
Step 8 : One sample only (for now) BamHI, 1:200 in Pop Buf (Life Tech) ; 0.000000 s.v.; 0.000000 min.
Step 9 : Stretch For a While ; 0.000000 s.v.; 0.000000 min.
Step 10 : Other Sample, EcoRV at about 4:34 AM (1:50 in PopBUf) ; 5.000000 s.v.; 0.000000 min.
|=============================================================
Added Eco RI
214 0.5 MHz / s A few BamHI files...NOTEThere are extra sites!!!
215 0.5 MHz / s
216 0.5 MHz / s in 10 mM CaCl2
EXCEPT AT END! FORGOT TO SWITCH FILES!
217 EcoRV i nPop Buf 0.5 MHz / s
Maybe a couple non-specific bindings here and there, but for the most part, either non-detectable sliding, or no binding
218 EcoRV 5 MHz / s
219 EcoRV 0.05 MHz / s
220 EcoRV Calcium + PBS 0.05 MHz / s
1 segment only
221 EcoRV Calcium 10 mM in PBS 0.5 MHz / s
Sample is completely fucked up (precipitate) all beads stuck in a matter of minutes. Maybe got one pop
010712.txt
7/12/01 4:55 PM
|=============================================================
Step 0 : Bert's slides ; 0.000000 s.v.; 0.000000 min.
Step 1 : antidig, fresh, 1:10 in PopBUf (20 ug / ml) ; 1.000000 s.v.; 3.000000 min.
Step 2 : BGB 10 mg / ml, ; 2.000000 s.v.; 10.000000 min.
Step 3 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 4 : 17-mer, 1:4 in PopBuf ; 1.000000 s.v.; 6.000000 min.
Step 5 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 6 : Bangs, soncicated, 1:10 PopBUf ; 1.000000 s.v.; 12.000000 min.
Step 7 : PopBuf ; 10.000000 s.v.; 0.000000 min.
Step 8 : BsoBI (one sample only), 1:50 in PopBuf ; 2.000000 s.v.; 0.000000 min.
|=============================================================
3:57 PM 1:10,000 BsoBI in Sample B
4:40 PM Washed with 5 s.v. PopBUf
222 BsoBI 0.5 MHz/ s
Just a couple files, shows popping
223 BsoBI 0.005 MHz / s
A single segment with peeling
224 BsoBI 0.05 MHz / s
Segment 8 possibly shows sliding
225 BsoBI 0.5 MHz / s, 1:10,000
226 BsoBI 5 MHz / s 1:10,000
30 segments
227 forgot to check reversal box
228 BsoBI 0.5 MHz / s, stop after 4000 points, reversal
Same tether 4 times
229 BsaBI 0.5 MHz / s, washed 5 s.v. (sample A from before)
Looks like enzymes diss. very quickly?
230 BsaBI 0.5 MHz / s sample A, washed 5 times (same as last file)
Shows some zero poppers, but also many poppers
010713.txt
7/13/01 10:58 AM
This sample is for studying z-height (will do many z telescope settings)
|=============================================================
Step 0 : Karen's Slides & cover ; 0.000000 s.v.; 0.000000 min.
Step 1 : antidig, fresh 1:10 popBuf ; 1.000000 s.v.; 3.000000 min.
Step 2 : BGB (I left it at room temperature AGAIN!...probably like 6 hours) ; 2.000000 s.v.; 8.000000 min.
Step 3 : New 17-mer (1:4 dilution of previous 1:10 dilution...was also at room temperature last night) (Forgot to wash before this step) ; 1.000000 s.v.; 3.000000 min.
Step 4 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 5 : Bangs 1:10, freshly in popBuf, sonicated 1 minute ; 1.000000 s.v.; 10.000000 min.
Step 6 : PopBuf ; 10.000000 s.v.; 0.000000 min.
Step 7 : Seal ; 0.000000 s.v.; 0.000000 min.
|=============================================================
With z = 8.5, it seems like I have to focus about 500 nm +/- 200 nm into solution in order to make reflection of laser beam look small. This would imply that trapping center is about 500 nm above beam waist?
Arbitrary estimation of when beam waist hits surface, when I am focused on surface (multiple measurements, but will undoubtedly be biased): 10.55 mm; OFF-SCALE...HMMMMM I was "wrong" on the first measurement (or at least I changed my mind as to how it should look when the beam hits the surface of coverglass). Video on Koch VII, 9 AM today...shows dust and laser, with z = (offscale positive, estimated 13.5 mm)
Laser = 1V, 14 amps. Estimation of parfocal bead: 9.35; 9.14; 9.13; 8.95; 9.36; 9.08; 9.19; 9.25; 9.01; 9.34 (9.18 +/- 0.144 mm) fairly insensitive to depth into solution, but these measurements were taken close to surface, but far enough away so that trapped bead could drop below focus without hitting surface.
Laser = 4V, 14 amps. Adjusted DIC. parfocal bead: 9.08; 9.14; 9.34; 9.14; (refocused objective) 9.18; 8.99; 9.02; 9.18; 8.90; 9.18 (avg = 9.12 + / - .123 mm)
Laser = 4V, focusing on small dust particles, trapped free bead. Lower z telescope, until I think bead is stuck (and I am right, because it is stuck); Therefore, this is a lower limit for when a trapped bead sticks, when focused on the surface. Multiple measurements: 6.95 mm; 8.95 mm; [start to notice change in signal at 10.25 mm] 9.20; 9.38; 9.92
[Note, this gives an estimate of dust focusing error]
Same as last, but now am just stepping the z-telescope down, so I can get an upper and lower bound...And I am saying it is not stuck if it flies out of trap momentarily when I cycle beam off / on quickly. multiple measurements: 9.1 to 9.2 mm; 9.9 to 10.0 mm; 9.7 to 9.8 mm; 9.5 to 9.6
[Note, this gives an estimate of laser focusing method error]
Same as last, but am using laser pattern at z = 10.5 mm to focus on surface. 8.6 to 8.7 mm; 8.8mm to 8.9 mm; [VIDEO KOCH VII 9:54 AM, focusing method; bead flew away at 9.3 mm, though]; 8.9 mm to 9.0 mm; 9.5 mm to 10 mm; 9.0 to 9.1 mm
230 z = 11 mm
DO NOT LIKE THIS FOCUSING METHOD...WILL ABANDON
231 z = 13 mm
Focusing on laser beam, as discussed in journal to data file (calibrated image at z = 13 mm, and very small dust particle...that is, a DIM dust particle)
232 z = 12 mm
233 z = 11 mm
234 z = 10.25 mm
not even sure if i got a single full peeler, but this method is much too much of a pain in the ass.
010713_2.txt
7/13/01 1:41 PM
|=============================================================
Step 0 : Karen's Slides & cover ; 0.000000 s.v.; 0.000000 min.
Step 1 : antidig, fresh 1:10 popBuf ; 1.000000 s.v.; 3.000000 min.
Step 2 : BGB (I left it at room temperature AGAIN!...probably like 6 hours) ; 2.000000 s.v.; 3.000000 min.
Step 3 : BGB ; 5.000000 s.v.; 0.000000 min.
Step 4 : New 17-mer (no dilution of previous 1:10 dilution...was also at room temperature last night) (Forgot to wash before this step) ; 1.000000 s.v.; 3.000000 min.
Step 5 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 6 : Bangs 1:10, freshly in popBuf, sonicated 1 minute ; 1.000000 s.v.; 10.000000 min.
Step 7 : PopBuf, supplemented with 1mM biotin, which also results in 1% DMSO ; 10.000000 s.v.; 0.000000 min.
|=============================================================
PLEASE NOTE THE BIOTIN! (And DMSO)
Washed again with popping buffer, because tethers were falling off surface when i looked after step 7. This was probably due to DMSO, in my opinion.
At 1:33 PM Added BsoBI 1:2,500 in:
10% PopBuf in addition to:
10 mM EDTA (so, final is 11 mM)
0.02% Tween (so, final is 0.022%)
235 z = 10.25
This can go with the last group of files...I replaced the DMSO buffer with popping buffer, so now should be the same peeling forces.
236 high data rate, for early fluctuations
237 ?
238 One peeler after adding the BsoBI...the only one i could find. All tethers either stuck or detached, i cannot tell which. Unexpected result of the low-salt buffer.
010714.txt
7/14/01 6:47 PM
V:\Aatte\koch\data\Popping_VC_BsoBI
|=============================================================
Step 0 : Karen's Slides & cover ; 0.000000 s.v.; 0.000000 min.
Step 1 : antidig, yesterday 1:10 popBuf ; 1.000000 s.v.; 1.000000 min.
Step 2 : BGB same tube ; 5.000000 s.v.; 2.000000 min.
Step 3 : BGB ; 5.000000 s.v.; 0.000000 min.
Step 4 : New 17-mer (no dilution of previous 1:10 dilution.) ; 1.000000 s.v.; 2.000000 min.
Step 5 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 6 : Bangs 1:10, freshly in popBuf, sonicated 1 minute ; 1.000000 s.v.; 8.000000 min.
Step 7 : PopBuf, supplemented with 1mM biotin, which also results in 1% DMSO ; 5.000000 s.v.; 0.500000 min.
Step 8 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 9 : BsoBI, very roughly 1:100 in PopBuf, added at about 5:30 PM, at least 10 minutes before first stretching ; 2.000000 s.v.; 0.000000 min.
|=============================================================
About 5 tethers per 90 micron octagon, and about 10 stuck beads
DataFile003: Washed with 5 s.v. PopBuf (no Enzyme at 6:27 PM)
002 0.5 MHz / s; 50 segments
003 0.5 MHz / s; 30-ish segments in no-enzyme buffer. The purpose of this file is to get information about what kinds of things happen during dissociation. For example:
A. Do certain sites dissociate earlier?
B. What is the kind of rate?
C. Do the enzymes seem to dissociate from the ends of the template?
010715.txt
7/15/01 4:16 PM
V:\Aatte\koch\data\Popping_VC_BsoBI
|=============================================================
Step 0 : Karen's Slides & cover ; 0.000000 s.v.; 0.000000 min.
Step 1 : antidig, yesterday 1:10 popBuf ; 1.000000 s.v.; 1.000000 min.
Step 2 : BGB same tube ; 5.000000 s.v.; 1.000000 min.
Step 3 : BGB ; 5.000000 s.v.; 0.000000 min.
Step 4 : New 17-mer (no dilution of previous 1:10 dilution.) ; 1.000000 s.v.; 2.000000 min.
Step 5 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 6 : Bangs 1:10, freshly in popBuf, sonicated 1 minute ; 1.000000 s.v.; 4.000000 min.
Step 7 : PopBuf, supplemented with 1mM biotin, which also results in 1% DMSO ; 5.000000 s.v.; 0.500000 min.
Step 8 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 9 : BsoBI, 1:100 in PopBuf, added at about 5:30 PM, at least 10 minutes before first stretching ; 2.000000 s.v.; 0.000000 min.
|=============================================================
Much fewer beads than yesterday...probably about 1 tether per octagon at most
0004.dat 5 MHz / s, 50 tethers
010719.txt
7/19/01 4:28 AM
V:\Aatte\koch\data\Popping_VC_BsoBI
|=============================================================
Step 0 : Karen's Slides & cover ; 0.000000 s.v.; 0.000000 min.
Step 1 : antidig, fresh 1:10 popBuf ; 1.000000 s.v.; 1.000000 min.
Step 2 : BGB same tube ; 5.000000 s.v.; 1.000000 min.
Step 3 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 4 : New 17-mer (1:4 dilution of previous 1:10 dilution.) ; 1.000000 s.v.; 2.000000 min.
Step 5 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 6 : Bangs 1:10, diluted fresh in popBuf, sonicated 1 minute today ; 1.000000 s.v.; 5.000000 min.
Step 7 : PopBuf, supplemented with 1mM biotin, which also results in 1% DMSO ; 5.000000 s.v.; 0.500000 min.
Step 8 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 9 : BsoBI, 1:100 in 90% popbuf, 400 m Sucrose ; 2.000000 s.v.; 0.000000 min.
|=============================================================
Tethering not so great...difficult to find loose tethers
0006 0.05 MHz / s; 400 m sucrose
0007 0.05 MHz / s; 10 mM Tris only (no enzyme, but for what remained during / after wash)
0008 0.05 MHz / s CalTris (50 mM CaCl, etc.)
MOst of the tethers stuck (99% ish)
A couple peelers clearly show higher peeling force...certainly than the 10 mM Tris sample
010719_2.txt
7/19/01 8:53 AM
V:\Aatte\koch\data\Popping_VC_BsoBI
|=============================================================
Step 0 : Karen's Slides & cover ; 0.000000 s.v.; 0.000000 min.
Step 1 : antidig, fresh 1:10 popBuf ; 1.000000 s.v.; 4.000000 min.
Step 2 : BGB same tube ; 5.000000 s.v.; 1.000000 min.
Step 3 : New 17-mer (1:4 dilution of previous 1:10 dilution.) ; 1.000000 s.v.; 4.000000 min.
Step 4 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 5 : Bangs 1:10, diluted fresh in cal5Tris, sonicated 1 minute today ; 1.000000 s.v.; 7.000000 min.
Step 6 : Cal5Tris, supplemented with 1mM biotin, which also results in 0.5% DMSO ; 5.000000 s.v.; 0.500000 min.
Step 7 : Cal5Tris ; 5.000000 s.v.; 0.000000 min.
Step 8 : BsoBI, 1:100 in Cal5Tris ; 2.000000 s.v.; 0.000000 min.
|=============================================================
Tethering actually looks pretty good.
0009 0.05MHz / s, auotJ, cal5Tris BsoBI
FORGOT TO SET ALL OF THE AUTOJ PARAMETERS!
It is possible that slide was not firmly attached
0010 0.5 MHz / s, autoJ, cal5Tris BsoBI
30 segments
It is possible that slide was not firmly attached
0011 2 MHz / s autoJ, cal5Tris BsoBI
31 segments
It is possible that slide was not firmly attached
0012 0.2MHz/s autoJ, cal5Tris BsoBI
30 segments (I fixed slide, I think...if there were a problem before at all, which i don't really know)
0013 0.05 MHz / s autoJ, cal5Tris BsoBI
25 segmetns
010829.txt
8/29/01 10:59 PM
|=============================================================
Step 0 : This protocol is inexact, but pretty close to the usual ; 0.000000 s.v.; 0.000000 min.
Step 1 : Bert's glass & Karen's slide cover, rinsed with water ; 0.000000 s.v.; 0.000000 min.
Step 2 : antidig, from last week 1:10 PBS ; 1.000000 s.v.; 4.000000 min.
Step 3 : BGB: from a couple weeks ago: 20 mg / ml in H20 and then diluting 1:2 into PBS, then adding 4 M NaCl to 100 mM. I thin k the BGB precipitated, because the solution was cloudy, and it would not go through the 0.2 micron filter. I used this anyway. ; 2.000000 s.v.; 4.000000 min.
Step 4 : BGB ; 2.000000 s.v.; 5.000000 min.
Step 5 : BGB ; 2.000000 s.v.; 0.500000 min.
Step 6 : New 17-mer, 1:20 dilution of the rest of the gel extraction from the first week of July ; 1.000000 s.v.; 6.000000 min.
Step 7 : Bangs 1:10, diluted fresh in popBuf, sonicated 1 minute today ; 1.000000 s.v.; 8.000000 min.
Step 8 : PopBuf, supplemented with 1mM biotin, which also results in 1% DMSO ; 5.000000 s.v.; 0.500000 min.
Step 9 : PopBuf ; 5.000000 s.v.; 0.000000 min.
Step 10 : EcoRI, from 10x Concentrated, 1:100 into PopBuf (100 ul) added at 10:30 PM ; 5.000000 s.v.; 0.000000 min.
|=============================================================
Added the EcoRI (Concentrated, 1:100) at about 10:30 and sealed
0001 Strange results
0002 Strange results
011119.txt
11/19/01 11:19 PM
Short Capped Template Unzipping
NOTE: Even though this slide has VERY low tether density, the ratio of stuck to tethered is also very low. I would put the ratio at about 1:1, with about 1/4 tether per octagon (or maybe less).
So, something about Karen's blocking method is in fact very successful. Perhaps it is because I have incubated the beads for sometime in BGB buffer.
0001 Low cap template, 0.1 Mhz / s
0002 Low cap template, 0.01 MHz / s
0003 Again, but later on. There may be a couple interesting data sets here, but overall the capping is not working at all.
011201.txt
12/1/01 3:34 PM
Short Capped Template Unzipping
Low tether density again!
"Internal / Low" sample:
stuck / tether: 4/0; 4/0; 6/0
Many fewer tethers than "high" sample...I would say about 1 per five 200 micron fields
"Internal / High" sample:
stuck / tether: 6/ 1; 4/0; 5/0
It is overall pretty easy to find tethers, probably about 1 per 200 um field
0004 High capped template
0005 Low Capped template
0006 repeating a tether (with reversal)
0007 Low cap, reversal still on
011203.txt
12/3/01 10:35 PM
Today will be normal uncapped unzipping template (not gel extracted).
First I am comparing heated and unheated samples
Files will be stored in Popping_VC_Osmolite\
Heated sample (stuck / tether): 4 / 0; 0 / 0; 4 / 0; 0 / 0; (don't see any tethers in like 10 full-fields)
non-heated sample: 3 / 0; 1 / 0; 2 / 0; 3/ 0; 3 / 0
|===========
THERE ARE NO TETHERS IN EITHER SAMPLE
011203_2.txt
12/3/01 11:18 PM
This is trying tethering (according to the usual recent protocol) with the very diluted 17-mer from July. There are still NO tethers (I stretch the 3 I could find that perhaps looked like tethers, but no unzipping).
I wonder if something is truly fucked up with my buffers or other supplies?
0001 No tethers
011204.txt
12/4/01 10:03 PM
These are the end-labeled templates. At first I am hoping to see a very good tethering efficiency, given what I have learned lately about the failure of the internal reactions lately.
Yay!
This is a great control (unfortunately, somewhat). First, I see about 1 tether per 90 micron octagon in the bottom sample (high-cap, I think), and I see about 0.5 per 90 micron field in the top sample (low-cap, I think)
Now for stretching.
0008 top sample (low I think)
(With reversal)
0009 same sample (top), with reversal turned off
0010 bottom sample
011204_3.txt
12/4/01 12:28 AM
Trying another sample, with:
New antidig
New PopBuf
Beads are not in blocker but are just in popbuf
None of the biotin quenching
17-mer from July
And, I see NO tethers. Well, actually I see about 1 or less per big (200 um) field...I stretched about 7 of them, and only saw unzipping twice.
So, I mixed 10 ul of beads with some PCR-tagged DNA that has been sitting in the fridge for MONTHS...I flowed in, and tethers started forming IMMEDIATELY. Huh.
Files are in Fluctuations/Overstretch/
PoppingVC_osmolite/0002 No tethers
Fluctuations\Overstretch\0001 Some overstretch
Fluctuations\Overstretch\0001 Some overstretch
011214.txt
12/14/01 1:07 AM
This is the new pBR322 construct.
The protocol can be found in \PaperI\EQ constant\011213\*.ini
There are VERY FEW tethers. But I could easily find a few. There are less than 1 tether per 10 90 micron fields, depending on whether the slight jigglers are tethers. There are about 2 stuck beads per 90 micron
Flowed in cold 10xEcoRI 1:10 in Kk Buffer at 11:55 PM.
After taking EcoRI data, I flowed in old beads in BGB popping buffer, and then washed away after about 10 minutes (1:05 PM). I did not see a significant increase in tethers, which may lead one to believe that there is little DNA on the surface, or that it is laying down stuck.
0005 naked pBR322
0006 naked pBR322
0007 EcoRI, 0.5 MHz / sec
0008 EcoRI, 0.3 MHz / sec, with proportional to 300
0009 EcoRI, 0.1 MHz / sec, with proportional to 300
011214_2.txt
12/14/01 4:12 AM
THis is the second sample of the night.
I put in 10 times as much DNA, but there seems to be even FEWER tethers! Argghhh!
Enzyme added at 3:40 AM.
0010-0012 10xEcoRI 1:1000, 0.1 MHz / s proportion v-clamp referenced to 100
020122.txt
1/22/02 3:21 PM
Using the popping buffer tethering protocol...the tethering efficiency is much better than when i tried the Kk buffer last month.
2:12 PM, added EcoRI (10x), 1:1000 in popping buffer + ~10 nM hindIII capping oligo
2:33 PM...I found a tether farm in the middle of my sample!!! tether density increased from about 1 to 50 tethers per octagon
Popping_VC_EcoRI\020122\0013 Naked DNA
0014-0015 Data files for Richard
0016-0019, for measuring EQ constant of EcoRI in popping buffer on pBR322 site Files 0017 & 18, I am in what is called the "tether farm"...
File, 20, 21 pre-shining laser onto tether before stretching
FIle 22 Being VERY careful not to pre-shine laser before stretching (and thus centering is pretty bad)
020122_2.txt
1/22/02 10:30 PM
About 9:09 PM, added EcoRI 1:2,500 (10x stock) in Kk buffer (+ ~ equimolar companion = 3:2,500 of 1:80 dilution of P15 / P17 duplex, which has EcoRI site). The tethering protocol was the popping variety, but I flowed 4 sample volumes of Kk buffer through before adding EcoRI. I will start taking data immediately after adding EcoRI.
For the second batch of data (0026 ->), I added EcoRI, 10x 1:2500 in Kk buffer plus 100 mM extra NaCl, plus 1:10 of 5 mg / ml BGB in Hepes. The dilution was performed 1:50 twice.
before 0026 first batch of data
0026 directly after adding the second batch of (high salt) ecoRI
020123.txt
1/23/02 11:44 PM
Eco added at 10:08:30
Tethering protocol is the same as the popping buffer protocol from yesterday. I did not switch to the Kk buffer until I added the EcoRI, at which point, I flowed in 4 sample volumes of the EcoRI (I think it was 1:800 instead!!)
10:46 PM, added the following (1:400)
1:20 was 1 ul Eco added to 16.6 ul Kk plus 0.4 ul 5 M NaCl plus 2 ul 5 mg / ml BGB in Hepes
1:400 was 1 ul of the 1:20 added to 35.2 ul Kk plus 0.8 ul 5M NaCl plus 4 ul 5 mg / ml BGB in Hepes (NO! I think I screwed up the second dilution and used 1:800 instead!)
An important note with the second set of samples (with BGB), is that I am moving towards the entrance of the sample as time goes on. It seems like the popping frequency is increasing with time, but it's hard to say first whether that's even true and second, whether it's due to time or spatial...
Added the 150 mM NaCl at 11:18:40 PM. This one was made by diluting:
2 ul EcoRI 10x stock
0.536 5 M NaCl
4 ul BGB
33.5 ul Kk buffer
And then repeating this 1:20 dilution. I expect there to be > 50% occupancy.
NOTE: Above I was actually using 1:800 dilution!!!
Sheesh, the new 1:400 in 150 mM salt had TONS of occupancy. I wonder if now it's going to be too high of a Ka?
0029 One naked tether
0030 9 tethers, directly after adding the EcoRI
0031 After giving the sample a bit more time to equilibrate
<=0035 THese are with the BGB sample
0036 150 mM NaCl (this is more exact than the previous) plus 0.5 mg / ml BG in Kk derived buffer
020124.txt
1/24/02 6:18 PM
Typical popping buffer tethering protocol, following which, I added
EcoRI 1:80 into Kk based 250 mM NaCl buffer
Before adding the EcoRI, there are plenty of tethers, at least two per 90 micron
Added the EcoRI at about 4:40 PM
Flowed in 4 more s.v. of 1:80 EcoRI at 4:46:15 PM...I saw very low popping before, compared with expected. After a few more data sets, I will in fact up the stretch rate, because maybe the expected popping force is a bit too low?
At 5:15:15 PM, added 225 mM Kk based 1:80 EcoRI
NOTE: I think I may have used the WRONG BGB for the last sample (that is, popping based, instead of Hepes based). Therefore, I will repeat.
Added the repeat with the correct Hepes BGB at 5:32 PM
6:04 PM, added the 165 mM sample (1:400)...this sample looked a lot better
0040 250 mM NaCl, saw very little popping, possibly too early???
0046 225 mM NaCl
0048 New batch of 225 mM
after 6:04 PM...the 165 mM sample
020125.txt
1/25/02 4:14 PM
Repeating the 225 mM Data point, in hopes that the higher stretch rate will reveal more popping.
Added the enzyme at 3:03:30 ish
3:37:30 ish, added the 125 mM sample
4:01:30 ish, added the 100 mM sample
0054-0056 225 mM NaCl in Kk + 0.5 mg / ml BGB 10 MHz / s
0057-0059 125 mM NaCl in Kk + 0.5 mg / ml BGB 10 MHz / s
0060-0062 100 mM NaCl in Kk + 0.5 mg / ml BGB 10 MHz / s
020127.txt
1/27/02 8:20 PM
Popping buffer tethering protocol with 17-mer from July.
Then adding EcoRI 1:100 in 150 mM Kk buffer + 500 ug / ml BGB
Trying to do close site popping first, and then perhaps also complete template mapping
Good tethering before EcoRI, about 1 stuck and 1 tether per 90 micron octagon
EcoRI flowed in about 6:19 PM
Flowed in 5 s.v. more of the EcoRI at 6:56 PM
0063-0065 1 MHz / s 10 decimation
0066-0068 0.2 MHz / s 10 decimation
020127_2.txt
1/27/02 8:59 PM
New sample at roughly 8:30PM.
THis is using a 1:10 dilution of the "internal biotin / 17-mer / hindIII capped" unzipping construct from December 05, 2001 (which I have minimal notes on in my notebook). I am guessing it's about 200 pM stock concentration. The tethering protocol is the same as with the others lately: popping buffer protocol.
FLowed in EcoRI at about 8:37:30
Hmmm...somehow my sample turned really ugly after flowing in EcoRI. I don't know what happened. Perhaps the 150 mM NaCl is bad for the 17-mer? (Doubt it)
Perhaps I got immersion oil into the sample? That is possible, since I had too much on the condenser side.
0069 0.2 MHz / s 10 decimation; new sample; naked DNA
0070 0.2 MHz / s 10 decimation; new sample; 1:400 EcoRI
bubble in oil
0071 got rid of bubble
020128.txt
1/28/02 1:01 AM
New sample at roughly 11:45 PM
THis is using a 1:10 dilution of the "internal biotin / 17-mer / hindIII capped" unzipping construct from December 05, 2001 (which I have minimal notes on in my notebook). I am guessing it's about 200 pM stock concentration. The tethering protocol is the same as with the others lately: popping buffer protocol.
THis time, I got rid of the biotin quenching step
11:51 PM Sample did not have enough tethers
12:35PM Very, very few tethers in the 1:10 end-biotin sample
AND, there aren't any tethers in the undiluted internal biotin sample
There must be some other sort of tethering problem (dig or biotin?)
Tried flowing in freshly diluted beads in BGB, and no more tethers formed
0073 0.2 MHz / s 10 dec; naked 17-mer
0074 End-biotin version
020128_2.txt
1/28/02 3:52 PM
1:5 dilution of the internal anchoring 17-mer I tried yesterday. I added 200 ug / ml to the stock DNA tube. Also, I am using fresh antigid and fresh beads. I deleted the biotin quench step.
Tethers are about a couple per 90 micron ocatagon. THere are more stuck beads then yesterday. I stretch 9 naked tethers (file 75) and got 2 good stretches. Lots of weird shit happens. I wonder if I'm too close to the surface?
About 2:50 PM, flowed in 1:400 EcoRI in 125 mM NaCl...I saw beads going by the screeen as I flowed.
At 3:23 PM, changed to 1:3600 EcoRI in 125 mM NaCl by flowing in 6 s.v. of the new concentration. I will also switch back to 2 MHz / s
Indeed, the 1:3600 sample looked a lot better...I may have even gotten a good Figure 3
0075 naked DNA
0076 naked DNA, focusing intentionally high. I don't think the strangeness is reduced at all.
0077 A few stretches at 0.2 MHz / s
0078 Deciding to switch to 0.1 MHz / s
0080-0083 1:3600 EcoRI, 2 MHz / s
020129.txt
1/29/02 2:51 PM
Flowed in EcoRI 1:3600 in Kk 125 mM + 0.5 mg / ml BGB at about 2:21:30PM
0084 naked pBR322 in poppping buffer
0085-0090 125 mM EcoRI pBR322, 0.5 MHz / s VC
020129_2.txt
1/29/02 8:59 PM
Trying two new pBR322 samples, both 1:10 dilutions, but one I sonicated for about 30 seconds after dilution.
sonicated: stuck / tether: 2/2; 0/1; 0/1; 0/3; 0/2 (overall, looks pretty nice)
no sonicated: s / t: 2/7; 0/1; 3/1; 0/ 2; 2/1 (overall, can't say is different than sonicated sample)
Added EcoRI at about 8:15 PM
0090 Naked DNA
0091-0094 EcoRI 1:3600 125 mM pBR322 1MHz / s (about 10 pops)
0095-0097 same, but 2 MHz / s (about 11-13 pops...some overstretch)
0098-0100 same, but 1.5 MHz / s
020131.txt
1/31/02 5:48 PM
New capped 17-mer; Popping based protocol
Added 1:3600 EcoRI (125 mM Kk + 0.5 BGB) at about 5:34:30 PM
Before 0106 Various conditions of the new Proportional (J&F) clamp on naked 17-mer
106 With EcoRI; F & J enabled
107 EcoRI, F only
020206.txt
2/6/02 11:26 PM
Popping based tethering of pBR322. Antidig and Beads were diluted at least 1 week ago.
Then adding EcoRI at 11:01 PM, 1:3600 in 150 mM
0109?-0113 All EcoRI 1:3600 in 150 mM Kk
020207.txt
2/7/02 11:42 PM
Popping based tethering of pBR322. Antidig and Beads were diluted at least 1 week ago.
Then adding EcoRI at 10:45:30 PM, 1:6400 in 125 mM NaCl Kk
There is a weird spot in the Bertrand lens, somewhere between the condenser iris and the polarizer / 1/4 wave plate i think.
Flowed in EcoRI 1:50 in 250 mM NaCl at 11:15 PM
0113-0116 All EcoRI 1:6400 in 125 mM Kk
0117-0122 EcoRI 1:50 in 250 mM Kk
020209.txt
2/9/02 4:54 PM
Popping based tethering of pBR322. Antidig was diluted at least 1 week ago.
Beads are fresh dilution in 5 mg / ml BGB today
Then adding EcoRI at 3:40:30 PM, 1:4900 in 100 mM NaCl Kk (1x Kk plus 500 ug / ml BGB) (7 s.v.)
Added 6 s.v. 1:3600 EcoRI in 205 mM NaCl at 4:13PM, after washing with 5 s.v. of Popping Buffer.
Added 4 s.v. of 1:400 EcoRI at 4:32:30 PM (the above conc. was a mistake)
0123-0126 100 mM; 1:4900
0127-0130 205 mM 1:400
020301.txt
3/1/02 2:57 PM
Tethering is popping based with pBR322 1:10. There are a lot of tethers before adding EcoRI.
EcoRI added (No Ca or EDTA, "125 mM" NaCl) @ 1:31:30 PM
@1:57 PM, washed with 10 s.v. of 1x Kk Metal buffer
2:29:30 PM, added same EcoRI as above, except with 92 micromolar of CaCl2 added (so, EcoRI is 1:484 in 125 mM NaCl, 0.092 mM CaCl2)
0131 125 mM NaCl 0 Ca
0.1 MHz / s, jref = 600; Fref = 15 pN
0132-0133 Same as last, except changing initial (pre) stretch rate lower, to prevent overstretch
0134-0136 EcoRI plus Ca, stretch rate reduced to 0.025
020305.txt
3/5/02 2:14 PM
Tethering is popping based with pBR322 1:20. (Difference in protocol is that instead of washing beads away with 5 mg / ml BGB in popping buffer, I washed with 0.5x Kk Metal Buffer) Tethering is sort of crappy (moderate amount of stuck, very non-uniform tether density)
0137-0140 EcoRI 0.025 MHz / s (prop F & J) 0 mM Ca 125 mM NaCl
020308.txt
3/8/02 4:11 PM
With this new washing procedure (windex), stuck beads are not too plentiful. Seems to be comparable number of tethers and stuck beads. I haven't looked around yet to see whether there are still tether farms. After looking around a bit more, still no tether farms. I would say about 1 tether / 90 micron, and fewer than that for stuck beads.
After a while (3:56 PM), I did find a small tether patch (about 20 microns y dimension, 175 microns wide (x)).
At 4:01 PM, found a much bigger tether farm. There is a tall piece of gunk (say 20 microns diameter, infinitely tall), and in the patch around it, there is a tether farm about 100 microns in y dimension, and several hundred microns wide.
Another trend I seem to notice is that the tethers seem to like to form near or on bumps. I notice this, because I do not stretch tethers that have visible gunk associated with their area of the slide (where I think the bead is interacting with the gunk).
0141-0144 EcoRI 1:484 in 1x Kk Buffer "125 mM" NaCl. 100 kHz / s j600 F15
020309.txt
3/9/02 1:05 AM
@ 10:38:30PM, I added the EcoRI in 0 mM Ca 0 mM EDTA "125" mM NaCl.
Protocol is same as earlier today (windex wiped glass). Following beads, I washed with 2 s.v. 5 mg / ml BGB in Popping buffer, followed by 5 s.v. of 0.5x Kk 125 buffer (w/ EDTA), followed by 0.5x Kk metal 125 buffer (minus EDTA). Then I flowed in the 4.s.v of 1:484 EcoRI (5x NEB)
12:08AM, Flowed in 5 s.v. of 1x Kk Buffer supplemented to 50 mM EDTA. The hope here is to leach all of the metal ions out of the BGB and whatever else is in the sample. I'll let sit for a few minutes, and then re-try the 0 Ca 0 EDTA 0 BGB experiment
12:27AM, FLowed in 5 s.v. of 0.5x Kk metal buffer (0 mM EDTA)
12:29AM, another 5 s.v.
12:31AM, 4.4 s.v of EcoRI 1:484 in Kk metal 125 (0 Ca, 0 EDTA)
0145-0148 EcoRI 1:484 in Kk metal buffer 125 mM NaCl, 0 mM Ca, (0 mM EDTA is implied by "metal" label); 100 kHz / s, j600 F15
0149-0150 After the leaching attempt (50 mM EDTA). There are very few tethers left, so I didn't keep taking data.
020311.txt
3/11/02 1:19 AM
Sample was prepared with typical popping based protocol. Again, like recently, I scrubbed the slides with windex; sprayed windex off with building air; rinsed with barnstead water; sprayed water off with building air. pBR322 was diluted 1:20.
Following typical popping based tethering, I washed sample with 2 x 5s.v. of "1x Kk 125 Minimal", which is 10 mM Hepes, 125 mM NaCl, and nothing else. Then I added the 4.4 s.v. of EcoRI 1:484 in the same buffer.
In the first group of samples, it appeared to have lots of high force popping, as if I still had contamination.
At 11:55 flowed in EcoRI in 1x Kk 125 Min 920Ca (which is the same with 920 micromolar of Calcium Chloride)
Found a small tether farm at 12:04 AM
12:14AM flowed in EcoRI 1:484 in Kk 125 min 9200Ca (9.2 mM Ca)
0151-0153 EcoRI 1:484 in Kk 125 minimal buffer (Hepes and NaCl only); 100 kHz / s reference
0154-0156 EcoRI 1:484 in Kk 125 min 920Ca
0157-0160 EcoRI 1:484 in Kk 125 min 9200Ca
020312.txt
3/12/02 6:56 PM
Top sample is Bsa-popping buffer protocol (replacing BGB with NEB BSA 10 mg / ml)
Bottom sample is BGB-popping based.
Preliminary analyses: BSA sample has superior tether to stuck ratio, with about the same number of tethers. That is, it appears to have worked beautifully.
5:47 PM, Flowed into the BSA sample 4.4 s.v. of EcoRI 1:484 in Kk 125 min 0 Ca, after washing with 2 x 5s.v. of 1x Kk 125 minimal. AFter flowing in new buffer and EcoRI, the tether density seemed to go down DRASTICALLY. IN contrast, the BGB sample retained tethers later on.
[Note added a week later: I would guess that the decrease in tether density was a mistake. The repeat of this experiment showed a lower tether density to begin with.]
6:36PM, flowed 1:484 EcoRI in Kk 125 buffer (full compenents, except BGB) into the BGB sample. I did not pre-rinse with Kk buffer before flowing in the 4.4 s.v. of EcoRI
0161-0162 BSA sample; 1:484 EcoRI in Kk 125 min 0 CA
100 kHz reference
0163 same sample, switching to 1 MHz ref.
I think I am just seeing no binding.
0164-0166 1:22 EcoRI
0167-0169 BGB sample; Still did not see much popping, but tether density was reasonably high (indicating that RE cutting was not a problem, at least in this sample.
020315.txt
3/15/02 12:32 AM
Both samples on same glass, cleaned with windex scrub / h2o rinse method (building air). Both sampels I flowed through popping buffer before antidig.
Sample A "Bsa BLocking": Like normal sample, except BGB 5 mg / ml replaced with 10 mg / ml BSA from NEB. The DNA is 1:20 pBR322 in NEB BSA
Sample B "BGB Blocking": Like normal, except pBR322 diluted in 5 mg / ml BGB.
Tethering efficiency:
BSA Sample, large (200 micron) field of view:
Stuck / tether: 1/2; 4/3; 6/1; 3/0; 5/1; 1/0; 0/1; 4/0; 0/2; 11/1
BGB Sample:
s / t : 5/ 7; 3/8; 2/10; 5 / 5; 5 / 2
11:19PM Flowed the EcoRI 1:484 into the BSA Sample
11:48PM Flowed in EcoRI 1:484 in Kk Metal 920micromolar Calcium
12:20 AM Flowed into BGB Sample EcoRV, 1:50 in 1x Kk buffer. THe EcoRV is Gibco Lot # 1069588, which is 10,000 U / ml stock
0170-0171 BSA Sample: EcoRI 1:484 in Kk buffer (no BGB or extra BGB supplementation).
Still shows no occupancy!!!
0172-0173 Same sample, 50 times faster stretching
0174-0175 EcoRI 1:484 in Kk metal 920 Ca.
Still very low occupancy, although I probably DID see some binding (and high force even)
EcoRV/0002 low stretch rate
EcoRV/0003 high stretch rate
020318.txt
3/18/02 11:45 PM
BGB Tethering protocol.
Flowed in EcoRI 1:484 in Kk 125 (no BGB) supplemented to 10 mM EDTA. However, I made a pipeting error such that I put in a total volume of 46 ul instead of 44 ul. So, the dilution is a little higher, and the salts etc. are a little off. I flowed in at about 10:43:30 PM.
I found a small tether farm right away (indicating that windex treatment was not 100%??? Anyway, it was also easier to flow the initial PBS into the sample, which would say glass wasn't "super clean")
11:17 PM, FLowed into the same sample 4.4 s.v. of 1:484 EcoRI in Kk 125, including 500 ug / ml BGB (so, the same buffer I used to take data for the paper).
The BGB sample definitely seemed to have very high occupancy at the beginning. However, it subsequently seemed to drop. Then I moved back towards the center, and it may or may not have increased again.
I WONDER: Are the narrower samples (e.g., slides with two chambers) more prone to non-specific sequestering of the EcoRI at the edges. For example, sticking to the double-stick tape edges?
NOTE: This slide was actually a dual sample chamber slide (and even particularly skinny), even though I only prepared the bottom sample.
0176-0179 EcoRI 1:484, 10 mM EDTA
0180-0181 EcoRI 1:484, 1 mM EDTA 500 ug / ml BGB
0182-0183 Same as previous two segments, except that I ensured that I am more near the center (most flow) of the sample.
020319.txt
3/19/02 6:20 PM
Double-wide sample (more than 20 microliters, probably). Normal tethering protocol with volumes doubled. (BGB blocking)
Flowed in 1:484 at 5:32PM in Kk 125 supplemented with 500 ug / ml final of BGB
In the center of the sample, occupancy seemed fairly high, although perhaps slightly low (but definitely much higher than 50%)
0184-0187 EcoRI In the middle of the sample
0188-0189 Near the top edge of the sample
0190-0191 Back in the center again (seemed like less occupancy...there is danger that I was "re-mowing" the same grass.
020322.txt
3/22/02 2:08 AM
This will be the first data set with Jen-Jacobson EcoRI. The source is the HCDB (a small dilution into 50% glycerol). It will be 1:6400 dilution.
The sample is normal popping / BGB based. I did the windex scrub / building compressed air cleaning method, and I must say that the sample at first (before Eco) looks very nice.
Flowed EcoRI in (1x Kk 150 buffer (no BGB or Ca) at 1:01:30AM
For whatever reason, clearly not enough protein. Will try higher concentration.
1:08AM, replaced with 1:80 EcoRI (note, will have a higher proportion of the storage buffer)
1:33AM, flowed in 1:80 in Kk 150 plus 0.02% tween-20
It does not look like the Tween helped at all.
0194 1:6400
0195-0196 1:80 in Kk 150
0197-0200 1:80 in Kk 150 plus 0.02% tween
020326.txt
3/26/02 2:24 PM
Popping based protocol, 1:20 of pBR322 in popping buffer.
Beads and antidig are from last week dilution.
Added 1:3600 JJ EcoRI in Kk150 (with BGB) at about 1:33:30PM.
At 2:04 PM, replaced the JJ EcoRI by flowing in 6 s.v. of 1:3600 NEB 5xEcoRI in Kk 150 (with BGB)
0201 FJ 10MHz/s; JJ 1:3600
0202 VC 10MHz/s; JJ 1:3600
0203-0204 Fadjust 10MHz/s; JJ 1:3600
0205-0208 Fadjust 10MHz / s; NEB 1:3600
020326_2.txt
3/26/02 5:33 PM
Trying to as closely resort to old protocol as possible. Changes from last sample:
Slides cleaned with water only, no scrubbing (I even used polypropylene gloves while cleaning). New double-stick tape as well (on water-only cleaned slide)
I did the biotin quenching step (about 30 seconds with popping buffer bioton (so it has a little DMSO).
I also note that the coverglass are a new batch.
The 500 ul tubes are also a new container (don't know about batch)
Flowed in NEB EcoRI 1:3600 in Kk150 (BGB) at 5:06:30PM
5:27PM, Flowed in the same kind of EcoRI as before, except prepared in sterile tubes. (And dilution was 1:60 x 1:60, instead of 0.5:30 x 1:60 as before)
OK: I could swear that I saw something swimming (like backteria, except maybe too small)? In any case, that is terrible. So I'll go re-make everything, or at least most things. It's probably the BGB that is offending.
0209-0210 1:3600 NEB EcoRI
0211 1:3600 of the new batch
020327.txt
3/27/02 6:50 AM
Top Sample: BGB BLocking (Fresh BGB see lab notebook today)
Bottom Sample: Casein Blocking (Fresh)
Both samples popping buffer tethering, 1:20 pBR322
90 micron octagons:
Top: Stuck / Tether: 5/3; 7/2; 7/2; 8/2; 7/1
Bottom: S/T: 6/2; 3/1; 3/5; 8/1; 5/0
Both samples look similar, with extra stuck beads in both samples (compared to before the fresh BGB). Tether density is perhaps a little higher also.
Notes about sample prep: Glass cleaned only by squirting water and spraying with building air (no scrubbing or windex). I did not quench with biotin.
5:23:30AM, FLowed EcoRI (NEB 5x) 1:3600 into the top (BGB) sample.
About 5:48AM, Flowed EcoRI (NEB 5x) 1:3600 into the bottom (Casein) sample.
6:31:30AM, Flowed in the rest of the 1:60 NEB EcoRI into the casein sample. There may be a large bubble preventing good flow, however. (BTW, by this time, most of the tethers seem to have gone. I would guess detached (or cut), as opposed to stuck, since the overall density seems lower of stuck + tether)
0212-0214 BGB Sample, EcoRI 1:3600; Fast
0215-0218 Casein Sample, EcoRI 1:3600; Fast
0219 Casein Sample, EcoRI 1:3600, DFS
0220-0221 Casein Sample, EcoRI 1:60 DFS
020330.txt
3/30/02 5:53 PM
Aptamer project
In the 240 pM 2:1 sample (see lab notebook), I see no obvious "tethers"...perhaps some jiggling,...
90 micron:
(Stuck or Jiggling) / (really jiggling): 5/0; 1/0; 3/0; 8/0; 2/0
10:1 sample: 1/0; 4/0; 7/0; 4/0; 3/0
After adding 10x more complexes to the 10:1 sample, I did not really see an appreciable increase in bead surface density. So, I conclude that these samples don't work at all
020330_2.txt
3/30/02 8:28 PM
8:13 PM, flowed in 10 s.v. of BsoBI 1:100 in Kk150 (+BGB)
File Popping_VC_BsoBI/today
The occoupancy is just too low with these particular conditions. I can either try Kk 100, or resort to the old popping buffer.
0033 0.1MHz/s reference
0034 0.5 MHz / s reference
020331.txt
3/31/02 4:54 PM
Typical Popping based tethering protocol, 1:20 of HindIII-capped internal biotin. I included the biotin quench step. I forgot to sonicate the beads, which I diluted freshly today.
4:29PM, flowed in 1:50 BsoBI in popping buffer
Wow. I forgot to sonicate the beads. Is that why there are no tethers? There are very, very, very few tethers (I found 3), and many of the stuck beads seem to be double beads or more. Hmmm.. I flowed in sonicated beads for about 10 minutes. WHile more stuck beads appeared (and single ones...not clumped), I didn't really see more tethers. Strange. I will just have to re-try the whole prep.
0035 0.5MHz / s reference; only 3 segments because tether density is so low
020331_2.txt
3/31/02 8:33 PM
6:34 PM flowed in 1:50 BsoBI popping buffer
Through 0040, Same conditions 500 ref
020331_3.txt
3/31/02 8:45 PM
8:32 PM, flowed in AvaI
through 0002 AvaI 500 ref
020401.txt
4/1/02 4:18 PM
3:32:30 added the AvaI 1:50 in pure popping buffer.
3:47 FLowed in AvaI 1:50 in popping buffer supplemented with 50 mM extra NaCl
4:06:30 AvaI 1:50 in Popping buffer plus 500 ug / ml BGB
Tether density is annoyingly low.
0003 forgot to turn on "steve params"
0004 10 segments, AvaI 1:50 in pure Pop Buf
0005 AvaI 1:50 in Pop plus 50 mM NaCl
0006 AvaI 1:50 in Pop plus 500 ug / ml BGB
020401_2.txt
4/1/02 6:03 PM
This is a new sample, prepared just like the previous, except replacing BGB with Casein.
The tethering density is still pretty low, but seems significantly better than the last sample. However, it's about as good as the best from yesterday (which was BGB based), so it's likely that the difference is random rather than casein versus BGB.
5:15:30 Flowed in AvaI 1:50 in Pop plus 500 ug / ml Casein
5:32:30 FLowed in Ava1 1:50 in Kk metal 920 uM Ca
5:46 Flowed in AvaI 1:10 in Pop plus 500 ug / ml Casein
0007-0008 AvaI 1:50 in Pop plus 500 ug / ml Casein
0009 AvaI 1:50 KkMetal 920 uM Ca
0010 Same as previous, 5x stretch rate (still seems to be low occupancy
0011 AvaI 1:10 in Pop plus casein, 500 Khz ref
0012 same as previous, 3x faster
020402.txt
4/2/02 2:01 PM
Typical popping based tethering, BGB replaced with casein. (HindIII capped 17-mer diluted 1:20 in popping buffer).
Tethering denisty is pretty low, but doable. Stuck beads also very low.
1:44 PM, Flowed in 1:50 BsoBI popping buffer
0040-0041 BsoBI 1.5 MHz/s reference
020402_2.txt
4/2/02 6:38 PM
Typical popping based tethering, BGB replaced with casein. (HindIII capped 17-mer diluted 1:20 in popping buffer). I flowed in 4 extra sample volumes of antidig, to see if that would increase the tether density. It seemed to have no effect. May want to try reducing the casein before the DNA
Tethering denisty seems lower than the previous sample today.
0042-0044 BsoBI 1500 reference
020409.txt
4/9/02 4:31 PM
Typical popping based tethering, BGB replaced with casein. (HindIII capped 17-mer diluted 1:20 in popping buffer). double the time on DNA and bead incubation steps (was talking)
Added AvaI (concentrated) 0.5ul in 25 ul of popping buffer at about 4:21PM
0017 and previous naked DNA, screwing around with dF / dt clamp parameters
0018 AvaI, 1.5MHz/ s reference.
Even though using 5x concentrated, still seems to be very low occupancy.
020409_2.txt
4/9/02 5:01 PM
Same sample as the AvaI earlier
Flowed BsoBI in at about 4:41PM
0045 Data just for testing the dF / dt with max force
Hmmm...second segment did not seem to work with maxF
0048 re-trying at 60 pN. Problem is probably when the bead moves out of trap further, so that force is higher than we think, based on the setpoint.
0047 demonstrating that it works for maxF = 30 pN
0048 5MHz/ s
0049 3MHz / s
020508.txt
5/8/02 3:40 PM
Protocol can be found in data directory.
J-J EcoRI 1:50 in Kk200 added @ 3:05 PM
@ 3:25 PM, switched to J-J 1:50 in Kk150
0222 1:50 EcoRI in Kk200 dF / dt clamped
0223 1:50 EcoRI in Kk150 dF / dt clamped
0224 same sample, one data set VC clamping only
0225-0226 same sample, back to dF / dt clamp, 0.1 reference
020513.txt
5/13/02 4:42 PM
EcoRI this morning on 17-mer.
Protocol is all casein based, popping buffer, except for last step. I used casein in ALL steps at 5 mg / ml in popping buffer except EcoRI step, where I used 500 ug / ml in specified buffer. Even the DNA was diluted in 5 mg / ml casein (popping buffer, 1:20 17-mer HindIII capped)
J-J EcoRI 1:200 in Kk150 casein & tween; added @ 1:39 PM
0227-0229 EcoRI, 1 MHz reference
020514.txt
5/14/02 1:58 PM
J-J EcoRI on 17-mer.
Protocol is pretty much same as yesterday. Protocol is all casein based, popping buffer, except for last step. I used casein in ALL steps at 5 mg / ml in popping buffer except EcoRI step, where I used 500 ug / ml in specified buffer. Even the DNA was diluted in 5 mg / ml casein (popping buffer, 1:20 17-mer HindIII capped)
J-J EcoRI 1:400 in Kk150 casein & tween; added @ 1:13PM
By "Casein and Tween", I mean 500 ug / ml casein plus 0.02% tween
0230-0235 EcoRI, 0.3 MHz reference
0236 Some Restretching files, just to see how the 17-mer
behaves
0237 EcoRI, 0.3 MHz reference
020515.txt
5/15/02 5:28 PM
"Secret Peeking Software" was accidentally running during files 239 to 244 (or maybe 240-244)
J-J EcoRI on 17-mer.
Protocol is pretty much same as yesterday. Protocol is all casein based, popping buffer, except for last step. I used casein in ALL steps at 5 mg / ml in popping buffer except EcoRI step, where I used 500 ug / ml in specified buffer. Even the DNA was diluted in 5 mg / ml casein (popping buffer, 1:20 17-mer HindIII capped) The bead incubation step was over 1/2 hour long due to distractions.
J-J EcoRI 1:2000 in Kk150 casein & tween;
238-244 J-J EcoRI (1:2000) 0.3 MHz / s reference
020521.txt
5/21/02 6:12 PM
The first sample today, I forgot to store a "data set notes." The first sample was like the second one. They both had bubbles trouble. The first sample I forgot the casein wash after the beads, so I had to re-ad the eznyme later.
5:45 PM, second sample of the day.
Protocol is pretty much same as last week. Protocol is all casein based, popping buffer, except for last step. I used casein in ALL steps at 5 mg / ml in popping buffer except EcoRI step, where I used 500 ug / ml in specified buffer. Even the DNA was diluted in 5 mg / ml casein (popping buffer, 1:20 17-mer HindIII capped). Like before, the EcoRI is added in the wet lab, before mounting the sample. So, there is at least a 3 minute delay before the data set begins.
J-J EcoRI 1:2000 in Kk150 casein & tween
THis enzyme was diluted earlier today (the first of the two step 1:2000 dilution)
I don't know why, but this sample had a much lower density of stuck and tethers (sort of like the earlier one today). It may have been related to bubbles going through the sample. I should be more careful for that tomorrow. Maybe that also accounts for the high density of "crappy" tethers that I seemed to have stretched in this second sample today.
0245-0248 J-J EcoRI 1:2000 in Kk150 plus tween and casein
300 kHz reference; first sample
0249-0250 J-J EcoRI 1:2000 in Kk150 plus tween and casein
300 kHz reference; second sample
020525.txt
5/25/02 6:57 PM
Protocol is pretty much same as last week. Protocol is all casein based, popping buffer, except for last step. I used casein in ALL steps at 5 mg / ml in popping buffer except EcoRI step, where I used 500 ug / ml in specified buffer. Even the DNA was diluted in 5 mg / ml casein (popping buffer, 1:20 17-mer HindIII capped). Like before, the EcoRI is added in the wet lab, before mounting the sample. So, there is at least a 3 minute delay before the data set begins.
EcoRI added at about 6:18 or so
Overall, this sample very low tether density and crappier tethers. I incubated extra long on DNA and bead steps, don't know if that accounts for anything.
0251-0254 J-J EcoRI 1:2000; Kk150 plus casein and tween
020526.txt
5/26/02 1:20 PM
Protocol is pretty much same as last week. Protocol is all casein based, popping buffer, except for last step. I used casein in ALL steps at 5 mg / ml in popping buffer except EcoRI step, where I used 500 ug / ml in specified buffer. Even the DNA was diluted in 5 mg / ml casein (popping buffer, 1:20 17-mer HindIII capped). Like before, the EcoRI is added in the wet lab, before mounting the sample. So, there is at least a 3 minute delay before the data set begins.
EcoRI added at about 12:55 or so
Sample has a lot of "gunk" floating around, as if one of my solutions has gone bad. Gun has various forms, long skinny "sticks" (0.5 micron by 10's of microns) and spherical stuff (several micron diameter). None of it seems to be swimming.
0255-0258 J-J EcoRI 1:2000 Kk150 plus tween and casein
020526_2.txt
5/26/02 6:02 PM
Protocol is pretty much same as the All Casein protocol used for the past few weeks. However, I have replaced all Casein steps with freshly made BGB.
EcoRI added at about 5:35 or so
So, for this BGB sample, it did not seem to improve the number of "wacky" tethers. It seems that the majority of the tethers cannot go full length before going into a "weird" mode. One thing that I often see is the tether _almost_ break, followed by catching on something, and then a steady, jerky increase in force. I would imagine I have seen this at least 5% of the time, definitely more than that today, and I would expect it to show up in the data: force will be like unzipping / popping, then drop close to zero, followed by continuation of weirdness.
Since BGB and casein are likely similar, I can't really rule them out as the culprit. Although it does not seem to be the freshness that matters, since I just made fresh BGB. Perhaps I should compare also with NEB enzyme, and also with pBR322 construct?
One other thing to note, I suppose is that I could be using fairly old antidig, as I am almost running out, and only use 20 ul for each sample.
0259-0261 J-J EcoRI 1:2000 in Kk150 plus tween and BGB
020526_3.txt
5/26/02 8:45 PM
This is a dual sample, doing side by side comparison of pBR322 and 17-mer HIndIII capped. Both are with BGB and without EcoRI (last step is simply 5 mg / ml BGB popping buffer).
|===================
It is completely clear to me that these two samples behave very differently. The 17-mer is dominated by freaky tethers and crappiness. Conversely, the pBR322 sample has a large percentage (probably >50%) that stretch completely without any obvious weirdness. I did see a small 5-10% amount of weirdness in the pBR322 construct.
It is possible that it is simply a tether density effect. Since the 17-mer had slightly fewer tethers, perhaps I am biased towards finding weird stuff. Or it could be something intrinsic to the 17-mer construct.
In any case, it's probably not worth fooling around with the 17-mer under the conditions that I have been using.
|==========
8:31 PM, added J-J EcoRI 1:2000 (from older 1:44 dilution) to the pBR322 sample
0262-0263 17-mer; naked in 5 mg / ml BGB in popping buffer
0264-0265 pBR322; naked in 5 mg / ml BGB in popping buffer
0266-0268 J-J 1:2000 in Kk150 plus BGB plus tween; pBR322
020603.txt
6/3/02 6:19 PM
Popping Buffer / Casein protocol, similar to recent past.
5ul old capped 17-mer plus 5ul of popping buffer is what i'm tethering with.
BsoBI added at 5:35PM
With the 5x extra DNA, I do indeed get more "non-crappy" tethers. Probably the tether density was reduced somewhat by using old beads.
0050 1MHz / s
0051 2 MHz / s
0052-0053 5 MHz / s
0054 2 MHz / s
020621.txt
6/21/02 1:30 PM
Popping Buffer / BGB protocol, similar to recent past.
DNA is P1-dig-F and P13-bio-R PCR product (double tether)
Purpose is to do a DFS control, and show that other linkages are stronger than the DNA / protein interaction.
Oops, there are about 200 tethers per 90 micron octagon. So, I should use even a 1:50,000 dilution of my DNA (something weird is going on?).
0014 Just some practice data sets, I will need lower tether density to take the real control data.
020628.txt
6/28/02 1:02 PM
This sample sucked. Probably bad beads. I was trying to do the typical Popping / BGB 17-mer protocol. (2.5 ul of 17-mer in 7.5 ul of popping buffer). I will flow in fresh beads to see if I get better tethers.
So, I flowed in fresh 1:20 dilution (into 5 mg / ml BGB popping buffer) of bangs beads. I think, indeed, more tethers formed. So, I will remake the sample and take data (probably tomorrow).
0015 A naked DNA tether. The computer (Aatte) was behaving strangely. FTC was taking twice as long as normal, and then the system locked up when I tried to run Secret Peeking Software.
0016 more naked DNA after rebooting.
FIGURED OUT THE PROBLEM: My AOD Hard Limits frequency range had somehow gotten switched to 10 MHz. That was slowing down the FTC
020701.txt
7/1/02 10:04 AM
Typical Popping / BGB based. Diluting the HIndIII capped DNA 1:4 in 5 mg / ml BGB popping buffer. There is no biotin quench step. Stretching buffer is 1:100 BsoBI in straight popping buffer.
0018-0020 2 MHz / s df / dt
0021-0023 0.2 MHz / s dF / dt; Fresh BsoBI before hand
0024-0026 0.5 MHz / s dF / dt; Fresh BsoBI before hand
0027-0029 5 MHz / s dF / dt; Fresh BsoBI before hand
020701_2.txt
7/1/02 10:42 AM
Typical Popping / BGB based. Diluting the HIndIII capped DNA 1:4 in 5 mg / ml BGB popping buffer. There is no biotin quench step.
Previously, was the BsoBI sample today.
Stretching buffer is 1:100 AvaI (concentrated) in straight popping buffer.
0019-0020 Naked DNA (Same BsoBI sample, have washed with 5 s.v. Popping buffer (wait 10 minutes), then 5 s.v. again). This file shows that BsoBI is still hanging around somewhat.
0021 Added 1:100 AvaI (concentrated) at about 10:32 AM.
This file shows that AvaI does not bind strongly at this concentration (no detectable binding, really)
020708.txt
7/8/02 9:32 AM
0030 BsoBI 20 kHz / s reference. Deleted file, forgot to tape slide
0031-0033 50 kHz; fresh BsoBI added
0034-0036 2000 kHz; fresh BsoBI added before 0034
0037 10000 KHz / s ; fresh BsoBI added before 0037
020709.txt
7/9/02 12:53 PM
BsoBI 1:100 in Popping buffer
0038-0041 10 MHz / s
0042-0045 500 KHz / s; fresh BsoBI added before hand
0046-0048 100 kHz / s; fresh BsoBI added before hand
020906.txt
9/6/02 4:18 PM
XhoI 1:100 in popping buffer added before file 0049. The sample is sealed with nail polish
0049-0054 0.5MHz/s reference
0055-0057 0.05MHz/s reference
0058 5MHz / s reference
020909.txt
9/9/02 12:10 PM
XhoI 1:100 in popping buffer added before file 0059 at about 10:55 AM. The sample is sealed with nail polish
0059-0064 2 MHz / s
0065-0070 0.05 MHz/ s
New Directory: "Repetitive experimentation"
File 0071 same tether as file 0070, last segment in other directory
Testing how DFS would look on repetetive stretching
020910.txt
9/10/02 11:03 AM
XhoI 1:100 in popping buffer added before file 0059 at about 10:00 AM. The sample is sealed with nail polish.
(Incidentally, I looked at the sample from yesterday (24 hours old), and there were no viable tethers remaining. This means that the sample is indeed degrading fairly rapidly.)
Also as a note, all of these files lately (even including June and July) are with the HindIII-capped 17-mer. So, some of these data sets can actually be used for calibration as well.
0072-0073 200 KHz / s (Arthur and visitor were in the room talking)
0074-0077 200 KHz / s
0078-0082 5000 KHz / s
020911.txt
9/11/02 11:20 AM
BsoBI 1:100 added before file 0083. Sample is sealed with nail polish. (Note: this sample actually has a little less DNA than previous many weeks. I ran out of DNA today, so today used about 1.5 ul of DNA added to 7.5 ul of buffer (for 1.5 ul in 9 ul total).
0083-0088 200 KHz / s
0089-0091 50 KHz /s
021103.txt
11/3/02 2:34 PM
BsoBI 1:100 added before first file. BsoBI was in about 5-10 minutes before that. Sample is sealed with nail polish.
Protocol is normal Popping / BGB protocol. DNA is from July 2001, and is the "unpurified HindIII ligation attempt", diluted 1:2 today in 5 mg / ml BGB.
0092-0097 BsoBI loading rate clamp 500 reference. There is a strand of fuzz in the BFP, not sure which segments it may affect, if any.
0098-0100 5,000 reference
0101-0103 50 reference
021112.txt
11/12/02 12:43 PM
BsoBI 1:100 added before first file. BsoBI was in about 10 minutes before that. Sample is sealed with nail polish.
Protocol is normal Popping / BGB protocol. DNA is from July 2001, and is the "unpurified HindIII ligation attempt", diluted 1:2 today in 5 mg / ml BGB.
NOTE: The BGB is the same tube that is a couple weeks old. Also, I forgot to sonicate the beads today. For one or both of those reasons (or something else), the sample today is not nearly as nice as last Sunday. I see many more strange things, and the tethers seem to break earlier. Also, there are many more clumped beads stuck / tethers around. THis may be good to keep in mind, if the data end up looking strange.
104-109 500 kHz / s
110-112 10,000 kHz / s
021112_2.txt
11/12/02 4:03 PM
BsoBI 1:100 added before first file. BsoBI was in about 5-10 minutes before that. Sample is sealed with nail polish.
Protocol is normal Popping / BGB protocol. DNA is from July 2001, and is the "unpurified HindIII ligation attempt", diluted 1:2 today in 5 mg / ml BGB.
NOTE: The BGB is the same tube that is a couple weeks old. I remembered to sonicate the beads this time.
After taking all the data with the second sample, I wouldn't say the sample was any better than the first (except fewer clumped beads). So, perhaps it still is the BGB being old (as opposed to the lack of sonication)
113-115 5,000 KHz / s
116-118 200 KHz / s
119-121 100 KHz / s
122-124 2000 KHz / s
021120.txt
11/20/02 11:02 AM
XhoI 1:100 added before first file. XhoI was in about 5-10 minutes before that. Sample is sealed with nail polish.
Protocol is normal Popping / BGB protocol. DNA is from July 2001, and is the "unpurified HindIII ligation attempt", diluted 1:2 today in 5 mg / ml BGB.
The BGB was made yesterday with Dan (A difference is that I used room temperature popping buffer to make the BGB, whereas I usually use cold).
The antidig is several weeks old, I think.
0125-0131 2,000 kHz / s reference
0132-0137 5,000 kHz / s reference
0138-0142 50 kHz / s reference
0143-0145 500 kHz /s reference
030115.txt
1/15/03 4:36 PM
Protocol is normal Popping / BGB protocol. However, Popping buffer is made from DSJ 5x Popping buffer July 2002. So, THE pH IS NOW 7.5!!!
DNA is end-labeled from December 5 2001. I diluted 1:10 in BGB 5.
BGB made fresh today. Antidig diluted fresh today. Beads diluted fresh today.
Today will be Betaine studies. All buffers based on 1x Popping (DSJ), and all enzyme (if present) will be 1:100 dilutions of the BsoBI (same as from Papers)
The sample doesn't have very many tethers.
NEXT TIME use 1:2 dilution!!!
Will give up on this sample
0146 In naked Popping buffer
030121.txt
1/21/03 12:54 PM
Protocol is normal Popping / BGB protocol. However, Popping buffer is made from DSJ 5x Popping buffer July 2002. So, THE pH IS NOW 7.5!!!
DNA is end-labeled from December 5 2001. I diluted 1:2 in BGB 5.
BGB made fresh last week. Antidig diluted fresh last week. Beads diluted fresh today.
Today will be Betaine studies. All buffers based on 1x Popping (DSJ), and all enzyme (if present) will be 1:100 dilutions of the BsoBI (same as from Papers)
Sample has more tethers than with 1:5 dilution. However, the tethers don't seem to be full length (as if nicked)
0149? (Check time) In 5 mg / ml BGB / popping buffer (DSJ)
0149-0150 Added BsoBI (5 s.v.) (1:50 in 1x DSJ Popping buffer) at about 11:46 AM, after washing with 5 s.v. of 1x DSJ Popping buffer
0151 Flowed in 5 s.v. more of BsoBI at 11:55 AM
0152 Re-trying with new buffer and enzyme. (Re-diluted 5x DSJ Popping buffer and re-diluted BsoBI in new 1x Pop) Flowed in 5 s.v. at about 12:10 PM
0153-0154 BsoBI 1:50 in SJK Popping buffer, flowed in 5 s.v. at about 12:19 PM
0155-0157 2/6 M Betain (1:6 dilution of 2M Betain in Popping) plus 1:50 BsoBI @ 12:30 PM
030124.txt
1/24/03 5:49 PM
Protocol is normal Popping / BGB protocol. However, Popping buffer is made from 2x Popping buffer January 2003. BGB is actually 10 mg / ml ish.
DNA is end-labeled from December 5 2001. I diluted 1:2 in BGB 10.
BGB made fresh today. Antidig diluted fresh last week. Beads diluted fresh today.
Today will be Betaine studies.
I will try probably only 1 M betaine 1x popping buffer plus 1:1000 BsoBI
16 amp laser, sum signal at 1.5 V AOD is 243 units.
0158-0159 1M betaine 500 kHz / s; 1:1000 BsoBI added at 4:19 PM
0160 repeating same tether
0161 repeating same tether with reversal
0162-0163 same as 0158-0159; added fresh enzyme before hand
0164 refolding last tether from 0163
0165 back to normal
0166 michelle data
0167-0168 michelle data, 10,000 reference
0169 same tether, michelle trying reversal at 10,000 initial
0170 same tether, michelle trying reversal at 500 initial
Untitled.txt
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0025.dat Contains example of 0.35 um peeler. Otherwise mostly junk
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0026.dat Contains example of 0.53 um in BsaIBuffer
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0027.dat Pre-BsaI, shows some examples of artifactual data
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0028.dat Some Pre-BsaI, no good data
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0029.dat Trying to gather Pre-BsaI data. Large File.
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0030.dat BsaI--contains one good template
D:\Aatte\koch\data\Project 3 -- Overstretch and Y Templates\010326\0031.dat BsaI--really contains no good templates
