Deborah, On 6/11/02 we performed the experiments described below. The images taken this day are all reachable from http://www.cs.caltech.edu/research/bio/Images/deborah/index.html Both flattened DI files and tif files are available. Because of name constraints in the stupid DI software, all of the tif files have the name rseXXX.tif rather than the more informative names of the DI files. All of the DNA stocks that I gave you were 10 uM in water. A previous email details the strands and how they are hybridized to compose any of 5 tiles, RE, SE, UE, VE, or SE15J where SE15J differs from SE by the addition of 2 extra hairpins. In the experiments we did, tiles RE, SE15J, UE, and VE were prepared separately. We made 100 ul (microliters) of each by mixing: 4 ul of each of 5 strands 10 ul of 10XTAE/Mg 70 ul milli-Q H20 10X TAE/Mg is just 10X TAE with .125 molar Magnesium Acetate. Diluted to 1X there will be 12.5 mM Mg++ in the buffer. Samples were prepared in PCR tubes, mixed by flicking, and spun down. Then each of the 4 tile samples were subjected to the following conditions in an eppendorf gradient PCR machine: 105 degree lid. 90 for 5 minutes then 85 to 25 at 1 degree/min. Preparation of the samples: In general 5 ul of sample was pipetted onto freshly cleaved mica (roughly 1cm octagons cut from 1cmx 4xcm strips) mounted on a large (15 mm) steel puck. (the mica was attached to the puck with hot glue). A petri dish cover was placed over the sample and it was allowed to adsorb for 3 minutes. Then 30 ul of of 1X TAE/Mg buffer was added and the sample was imaged in tapping mode under buffer (more buffer is added because the fluid cell is wet before it is positioned over the sample, more details later). We looked at 5 samples over the course of our session. Initially we made two sample tubes: 10 ul of RE and 10 ul of SE15J (sample 1) and 10 ul of UE and 10 ul of VE. We never looked at the UE/VE mixture. After mixing sample 1 ten times with the pipettor, we immediately adsorbed it to mica and imaged it. Images rse.001 to rse.014 are of this sample. They show A) that the mica surface is entirely covered with small domains of DAE lattice B) there is an occasional tube between the lattices. Deborah's notes indicate that rse.002 is at 0 degrees scan angle and rse.003 is 90 degrees CCW. Also rse.004 is at 0 degrees and rse.005 is rotated 90 degrees. After 1 hour 45 minutes we took another 5 ul sample out of sample 1 and imaged it. These are files rse1h45.015 to rse1h45.036 As you noted, for the first 10 micron scan, the sample appears to be mostly many micron tubes (with some very small pieces of tube/lattice). **Our current hypothesis is that at early times, (as in the first imaging experiment at essentially 0 time) few tubes have had an opportunity to nucleate and grow in solution. Thus, when the sample is incubated over mica during the adsorption step, monomers nucleate lattices on the mica, and the mica becomes covered with small sheets. After 1h45 tubes have had suitable time to nucleate and grow. Thus we see almost entirely tubes. From rse1h45.019 to rse1h45.020 there is a 90 CCW rotation. in image rse1h45.023 the scan angle was rotated back. After 4 hours and 30 minutes we took another sample and let it adsorb to mica for 3 minutes. Then we added the customary 30 ul of buffer and let the puck sit out on the lab counter for 45 minutes before imaging. Upon imaging, (rse4h30.037 to rse4h30.040) we were suprised that we saw rather than tubes, we seemed to see lots of long narrow sheets (but no small sheets nucleated on mica) that seemed to participate in some larger sheets. This was weird---one interpretation is that leaving the sample out in the open air, subject to evaporation, may have promoted tube opening---or perhaps that particular sample of mica was particularly sticky and this promoted tube opened (or something.) Deborah did float the hypothesis that the micas stickiness for DNA lattice was promoting opening of the tubes. (Mica certainly does this with cationic lipid vesicles. They start life as little globules that fuse flat to a mica surface). Surprised and disturbed by this, we took our last 5 microliters of sample and looked at it immediately (rse4h30b.041 to rse4h30b.045) and found that the sample was still tubes. rse4h30b.045 is one of the highest res images (showing individual DX) that we took that day. We felt a little better. Finally, interested in seeing the effects of concentration, we diluted RE and SE15J to .02 uM (from .2 uM) and mixed 10 microliters of each. After 1h 18 minutes we observed the pictures in rsed1h18.046 to rsed1h18.051 We see mostly things that we interpret as either short (~200 nm) tubes or small lattices and a few longer (a couple-few microns). AFM tips: What follows is not a detailed AFM protocol but rather some of the parameters we use and some suggestions for getting the best possible images. Imaging was performed on a DI multimode in tapping mode. We used DI's NPS tips (nitride sharpened tips.) There are 4 cantilevers on each NPS chip. We use the short (100 micron) cantilever with skinny legs. We used a vertical-engage J-scanner. (100 micron scanner) We start with an integral gain of .4 and a proportional gain of .6 I generally engage with a scan size of 10 microns. We use the 9 kHz resonance of the NPS tips. We feel comfortable imaging with an tapping amplitude of anywhere from .5 to 1.5 although I have imaged with oscillation amplitudes of .3. We try to perform imaging with a drive amplitude of less than 500 mV. We assume that the lower (and less violently the tip is driven) the better. Often 200-300 mV is all that is necessary to get a reasonable oscillation amplitude (say 1V). **We put the tip into the AFM fluid cell _dry_ so that it can be positioned easily and then add 30 ul of TAE/Mg buffer on and around the chip (making sure that the top of the chip and cantilever is wetted). This is important so that 1. There will be enough buffer when this is combined with the 30 microliters that is on the sample. 2. There is no air bubble on top of the chip---if one gets formed then the laser will be refracted by going through the interface and no signal will be observed! We do not use the silicone O-ring. Instead, if the fluid drop is/becomes too small we sneak a small round gel pipette tip in the side and add TAE/Mg buffer (or milli-Q H20) Also, be careful installing the chip in the fluid cell. The weak point in the system is the damn spring/arm that holds the chip in place. Make sure to depress the spring with a thumb and get the arm that holds the chip up off the surface of the fluid cell before rotating it to secure the chip. If you attempt to rotate the arm while the spring is relaxed it may bind and break the arm and/or the spring. This is a known bug---be careful! Also, be careful when putting the fluid cell in or out of the AFM head--- the right arm (of the two that secure the fluid cell) will catch on the spring/arm of the fluid cell and move it causing either A) the chip to come loose B) perhaps in an extreme case the arm/spring to break. Also we put a small saran wrap cover over the end of the J-scanner to avoid getting buffer into the piezo and shorting it out. We use a small (few cm on a side) square of saran wrap. We are unsure if this causes resolution problems (by making the mechanical coupling of the sample to the piezo sloppier) but it doesn't seem to and makes us feel better. (We know people who appear to have shorted out a piezo with buffer). We use the vertical engage scanner to avoid crashing the tip into the sample. This seems important for achieving the highest resolution imaging. A tip crash almost always abolishes one's ability to see individual double-crossovers. Rather than manually bringing the tip to the surface with the up/down switch on the microscope until a deflection can be observed (say by a slight change in the tapping signal), we try to rely on the computer entirely. We also do not engage in contact mode as many people do before changing to tapping mode. I try to be overly conservative, often starting very far away from the surface. If imaging degrades and doesn't improve with with a little fiddling, do not waste time---try changing the tip. Sometimes when the tip appears to have degraded (it is doubled or resolution has decreased) we attempt to 'clean' it by withdrawing the tip 20-50 microns from the surface and putting the instrument in 'tuning mode'. There we set the frequency sweep from 5-50 kilohertz (we have not studied whether a larger sweep to say 300 khtz might be better or whether a smaller sweep would do as well) and let the tip shake for a while---sometimes we increase the drive amplitude to 500 mV for this operation. Sometimes this seems to ameliorate tip doubling or low resolution, we assume by shaking crap off the tip.