Biomod/2011/Caltech/DeoxyriboNucleicAwesome/Protocols

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 Revision as of 23:34, 2 November 2011 (view source)← Previous diff Current revision (14:07, 3 November 2011) (view source) (→DNA origami formation) (23 intermediate revisions not shown.) Line 1: Line 1: {{Template:DeoxyriboNucleicAwesomeHeader}} {{Template:DeoxyriboNucleicAwesomeHeader}} =Protocols= =Protocols= - __TOC__ - ===Quantitating Newly Ordered DNA Strands=== + === Oligonucleotides=== - :''Main article: [[Biomod/2011/Caltech/DeoxyriboNucleicAwesome/Protocols/Quantitation]]'' + Most of the sequences were from Lulu Qian’s sequences, designed as described in [1]. Modifications were added using a computer program NUPACK, and staple sequences for the rectangular origami were taken from Rothemund’s [7]. All oligonucleotides were ordered from Integrated DNA Technologies (IDT) Inc. Most of the DNA molecules were ordered standard desalted and unpurified, and modified molecules are ordered HPLC purified. Staple strands were ordered in wet plate form at 150μM in 1x TE buffer. Dried DNA was reconstituted in an appropriate amount of ddH2O, and quantified with the biophotometer. + :''More detailed article about quantification protocol : [[Biomod/2011/Caltech/DeoxyriboNucleicAwesome/Protocols/Quantitation|Quantitation]]'' - *Provided by Damien Woods + ===Buffer conditions=== + 1x TE/Mg2+ (TE=Tris-EDTA, Tris 40 mM, EDTA 2 mM, and Magnesium acetate, 12.5 mM, pH 8.0) buffer was used for all experiments. - This is a 2-step protocol where we first try to set the DNA concentration to be very roughly 100 $\mu$M, and then adjust to bring it close to 50 $\mu$M. For a faster 1-step protocol you could easily adjust the calculations and go straight for 50 $\mu$M. - The protocol assumes we will dissolve in 1x TE, some people use Milli Q H20 instead. + ===Gel Electrophoresis/ Gel Purification=== + 5% polyacrylamide gels of 1 mm in thickness were used. Native gel was polymerized by mixing 3.75 ml 19:1 40% polyacrylamide, 1 mL 10x TAE/Mg++, 5.25 ml ddH2O, 60 μL 10% APS, 6 μL TEMED. Native gels were run at 150V at 25 ºC. 1x TAE/Mg2+ was used as a running buffer. Gels were stained with SybrGold for 15 min and then scanned with Quantity 1 FX scanner. Some complexes were annealed using PCR machine and gel purified using bigger gel. Same recipe was used to make native gel. Glycerol was used as a loading dye for the modified strands. Under UV light, the bands were cut and DNA was obtained by diffusion in 1x TE/Mg2+  buffer for 48 hours. + :''More detailed article about gel puriciation protocol: [[Biomod/2011/Caltech/DeoxyriboNucleicAwesome/Protocols/GelPurification|Gel Purification]]'' + ===DNA origami formation=== + Rectangular DNA origami consists of M13 viral DNA and 202 DNA staples. Staples at the side of the origami were not included in the 202 staples to avoid the stacking problem.  Sometimes, a probe at the walker start site was intentionally left out, making a hole in the middle of the origami.  M13 scaffold DNA and staple strands were mixed in 1x TE/Mg2+ buffer with molar ratio of 1:4 between the M13 and staple strands.  For the speed of assembly, origami were annealed at 90nM, and diluted into different final concentration for the purpose of various experiments; 15 nM of M13 were used for fluorescence experiments, and 10 nM, 5 nM, or 1 nM for or AFM experiments. Origami was annealed by heating up to 90 ºC then cooling down to 20 ºC at 1 ºC/min using an Eppendorf PCR machine. After annealing, tracks were added to the annealed origami at 1:6 ratio of M13 to tracks. Walker goal and a walker start complex which is a preannealed complex “walker – walker inhibitor – track1 – probe for track 1” were also added to the origami at 1:1 ratio of M13 to strands. Mixed solution was incubated at 37 ºC overnight. - 1. Switch on biophotometer. Prepare bench, lay out the things you'll need (for consistency, do this the same way every time you do the protocol). + :''More detailed article about origami puriciation protocol: [[Biomod/2011/Caltech/DeoxyriboNucleicAwesome/Protocols/Origami Purification|Origami Purification]]'' - 2. Spin down lyophilized (dried) DNA to make sure there are no flakes on/near lid. + ===Atomic force microscopy (AFM)=== + 5 μL of sample was deposited onto a freshly cleaved mica, and 20 μL of 1x TE/Mg2+ buffer was added. 40+ μL of 1x TE/Mg2+ buffer was added to the fluid cell and the sample was scanned in a tapping mode with DNP tips.  When streptavidin was used, 1 μL of streptavidin was added to the mica, and we waited 15 or more minutes before imaging to ensure the streptavidin and biotin could bind.  The exception to this was when we used the Walker Lock-Down Protocol. - 3. First, shoot for 200 $\muM: If IDT claim to give us [itex]x nanomoles, we add [itex]5x [itex]\muL 1x TE, to the dried DNA, to get roughly [itex] 200 \mu$M, explained as follows: + ====Walker Lock-Down Protocol==== - + To see what this protocol was used for see [[Biomod/2011/Caltech/DeoxyriboNucleicAwesome/AFM_Experiments#Walker Lock-Down Protocol|the AFM experiments page]].  (When we tried this protocol, the concentration of origami was too high, so we tried using 2 μL of sample instead of 5 μL, but in the future we will dilute the sample before using it, and we will stick to using 5 μL of sample.)  The sample, buffer, and streptavidin are mixed on mica as above, but after waiting 15 minutes, we do the following: - $\frac{x \times 10^{-9}}{5x \times 10^{-6}} \quad \frac{moles}{litre} \quad + - =\quad \frac{2x \times 10^{-9}}{x \times 10^{-5}} \quad \frac{moles}{litre} + - \quad + - =\quad 2\times 10^{-4} \quad \frac{moles}{litre}$ + - $\quad=\quad 200 \times 10^{-6} \quad \frac{moles}{litre} + 1. add 20 μL of 1x TE/Mg2+ buffer to mica, then remove as much liquid as possible (40+ μL) from the mica without touching the mica. - \quad + - =\quad 200 \quad \frac{micro moles}{litre} \quad = \quad 200 \, \mu M + - 4. Vortex (mix) for a few minutes. Spin down using microcentrifuge to remove any droplets form the lid/top of test tube. Vortex again. Spin. It is important to get the DNA completely into solution, so mix well. Finish by spinning down. + 2. add 40 μL of 1x TE/Mg2+ buffer to mica, then remove as much liquid as possible. Repeat this 3 times. (In the future we may repeat this more times to remove as much excess streptavidin as possible.) - 5. Put 98 [itex]\mu$L of solvent (1X TE buffer) into a plastic cuvette. Place in biophotometer with (upwards pointing) triangle facing you. Push down in a specific pattern (top-left-bottom-right, then bottom-left-top-right, then top left). (Insert the cuvette in exactly same direction, and using the same technique, every time.) Set large pipettor to one side (do not let the tip touch anything, I leave it over the side of the bench). + 3. add 20 μL of 1x TE/Mg2+ buffer to mica. - 6. Hit "blank". + 4. add biotinylated† walkers at a 1-to-1 ratio to the tracks or higher (excess should not affect the results, and when implemented a 1-to-1 ratio would require a volume that is too small to be pipetted, so excess is likely necessary.) - 7. Add 2 $\mu$L of DNA. Mix with small pipettor (suck up and down). Mix with large pipettor (suck up and down very carefully). Try not to introduce bubbles, nor splash on side of pipettor. + 5. wait for 15+ minutes before imaging sample. - 8. Insert cuvette, using the same technique as before. Hit "sample", and record the value. + †We have only seen the walkers when 5'-end biotinylated walkers were used, but 3'-end biotinylated walkers were only tried at a low concentration. - 9. Mix with large pipettor (stir, while in biophotometer), record value. + Based on our preliminary results using this protocol, streptavidin binds to mica, and if it is ever lifted off of the mica, it can bind to walkers on origami. Thus, we will try using the above protocol, but using 1x TE/Mg2+ with 100 μM Na+ instead to minimize the amount of streptavidin that binds to mica. - 10. Mix with large pipettor (remove from biophotometer, suck up and down), record value. Discard cuvette. + ===Spectrofluorometer (SPEX)=== + After cuvettes are cleaned with ddH2O and 70% ethanol, 10 nM - 1 uM sample was loaded in 1.5mL 1x TE/Mg2+ buffer into each cuvette for kinetics experiment in solution. For random walking experiment on origami, 200 ul of 15 nM samples were used. Fluorescent level of each cuvette was observed in real time. - 11. If the 3 values are close to each other (within a few percent), then record the first one. If not, try again. + :''More detailed article about SPEX protocol: [[Biomod/2011/Caltech/DeoxyriboNucleicAwesome/Protocols/SPEX|SPEX]]'' - + - 12. Make a note of the fact you have removed 2 $\mu$L from the DNA. + - + - 13. Repeat at least 3 times, by going back to step 5 on each iteration. Usually I do 3 to 5 samples. If the results are not consistent, re-mix, and do more. Let $A_{260}$ be the average of all samples. (If there is one outlier I ignore it.) + - + - 14. Use the following formula to find the concentration: []$\mu$M = dilution-factor $\times \frac{A_{260}}{\epsilon} \times 10^6$ + - + - Here dilution-factor = 50 (i.e. you put $2 \, \mu$L of DNA into  $98 \, \mu$L 1x TE), $A_{260}$ is the absorbance value, and $\epsilon$ is the extinction coefficient of your oligo usually given in /M/cm (supplied by IDT with your oligo, or else use the in-house DNA group code for double stranded complexes). This comes from: $A_{260} = C \times L \times \epsilon$ where C is the concentration, L is the path length (1 cm for the Biophotometer). And this in turn comes from the Beer-Lambert Law. [http://www.idtdna.com/analyzer/applications/Instructions/Default.aspx?AnalyzerDefinitions=true#ExtinctionCoefficient| See IDT's Page] + - + - 15. Shoot for 100 $\mu$M. We will calculate how much 1x TE to add in order to get as close as possible to 100 $\mu$M. To get the current volume of DNA in solution, subtract $2y \, \mu$L from the initial volume ($5x \, \mu$L), where $y$ is the number of DNA samples you took. Then apply the following formula to find out how much 1x TE to add to get the desired concentration (100 $\mu$M in our case): + - + - $\textrm{volume\_to\_add} \, (\mu L) = \frac{\textrm{current\_concentration} \, (\mu M) \times \textrm{current\_volume} \, (\mu L) }{\textrm{desired\_concentration} \, (\mu M)} \,\, - \,\, \textrm{current\_volume} \, (\mu L)$ + - + - 16. Add $\textrm{volume\_to\_add} \, (\mu \textrm{L})$ to the solution. Mix, spin, mix spin. + - + - 17. Do steps 4-14 with the new sample. If everything went well, you should get a value close to 100 $\mu$M. + - + - === Gel Purification === + - + - 1. Anneal the complex using a PCR machine + - + - 2.  Wash the plates and dry for 45 minutes + - For 300ul sample, add 50ul BB_ND dye + - 170 μl of sample will be loaded per lane. Each sample will take 2 lanes, and one blank lane should be placed between different samples. + - + - 3. Make a 12% non-denaturing (ND) gel for purification + - Lulu Qian’s recipe for 40ml of 12% ND gel which works for one gel + - - 12ml 40% Acrylamide + - - 4ml 10x TAE/Mg++ + - - Fill with MQ water to 40ml total volume (add 24 ml) + - - 240ul 10% APS + - - 24ul TEMED + - Note: APS should be used within 10 days + - + - 4. Run the gel under 150V for roughly 4.5 hours ~ 6 hours (when the dye nearly runs off the gel) depending on the size of the sample. For example, if the length of DNA strand is 122nt~176nt, run for 6 hours. Change the buffer every 1.5 hours. + - + - === SPEX=== + - *Provided by Lulu Qian. + - + - 1. Turn on the lamp and wait for 30 min for the lamp to warm up. + - + - 2. Launch the software: Instrument control center + - + - 3. Temperature set up + - Click the third square icon “Visual setup” + - Temperature control icon: Turn on and Set to 25 degreeC + - Choose T bath (internal). + - + - 4. Wash the cuvettes. + - Distilled Water (DW) (10 times) 70% ethanol (twice) DW (5 times) 70% ethanol (once) + - Air dry the cuvettes for 1 h. + - + - 5. Clean the cuvettes using lens paper. + - + - 6. Click second square icon “real time display”. + - High voltage: HV on, S 950, T 950 + - Bandpass : 2 + - + - 7. Click the last icon “run CWA”. + - Open the file “ROX_CWA” + - Optional mode: kinetics, Integration time: 10s + - Define: Repeat acquisition: 1 min (minimum) for a total of ~ min, depending on how long you would like to run the experiment. + - + - 8. Mixing: Mix the sample with 1X TE/Mg++ buffer inside cuvettes by pipetting for 30 times. + - + - 9. Start acquisition. Run sample. + - The data should not exceed 1 million (1000,000). + - + - 10. To add more samples (for example, walker triggers) in the middle of SPEX experiment + - When the status is “waiting”, click “stop”. + - Add samples and mix. Replace cuvettes into the SPEX machine. + - + - 11. Save the data + - When the status is “waiting”, click “stop”. + - Click “yes” + - Save data in two forms (DWA datafile, txt file) + - Check if the data is properly saved. + - *If you want to see the reference signals, go to options: view option and display raw R. + - + - === Origami Purification === + - + - *Provided by Nadine Dabby and Sungwook Woo. + - + - We suggest using the [http://www.millipore.com/catalogue/item/42413| Millipore Microcon YM-100] (blue) spin filters. The times below are estimates for how long to spin your sample (to err on the safe side I would check on the sample after 30 minutes and then estimate how much longer one needs to spin it so as to preserve a small amount of liquid on top of the filter but being VERY careful not to allow it to dry out completely. + - + - this protocol should drastically reduce the presence of excess strands. (the more rounds of adding buffer and filtering you do, the cleaner your sample, but the lower the yield. + - + - Admittedly the yield will be quite low (we haven't figured out exactly how low) -- supposedly each round of filtration should preserve 95% of the origami according to the filter spec but one problem here is dilution (since I have almost always ended up with a sample that is 3X more dilute than I started with). The dilution can be fixed by vaccuum centrifuging at 30 degrees (this does not hurt the origami at all according to Sungwook, but I haven't tried this with our substrate-laden samples. + - + - 1. Mix 100uL of your sample with 300uL of 1X buffer of your choice. + - + - 2. Vortex mixture and pipette it into a YM-100 spin filter. + - + - 3. Spin at 100g for ~30 minutes at 4°C. (Spinning at this low speed prevents the origami from being ripped apart) + - + - 4. Remove sample for centrifuge, leaving the machine at 4°C because we’ll be using it again in a moment. The spin filter should have only a few tens of microliters of your sample left in it, just enough to cover the filter. Remove the filter and get rid of any liquid in the tube below. Now reinsert your spin filter and add 200uL of 1X buffer, then pipette up and down just over the filter (five to six times) to recover any bound origami on the filter. Be very careful not to puncture the filter during this step. Finally, add another 200uL of buffer to your sample to bring the final volume to ~400uL. + - + - 5. Spin at 100g for ~30 minutes at 4°C. + - + - 6. Remove sample for centrifuge, leaving the machine at 4°C because we’ll be using it again in a moment. The spin filter should have only a few tens of microliters of your sample left in it, just enough to cover the filter. Remove the filter and get rid of any liquid in the tube below. Now reinsert your spin filter and add 200uL of 1X buffer, then pipette up and down just over the filter (five to six times) to recover any bound origami on the filter. Be very careful not to puncture the filter during this step. Finally, add another 200uL of buffer to your sample to bring the final volume to ~400uL. + - + - 7. Spin at 100g for ~30 minutes at 4°C. + - + - 8. Remove sample for centrifuge, leaving the machine at 4°C because we’ll be using it again in a moment. The spin filter should have only a few tens of microliters of your sample left in it, just enough to cover the filter. Remove the filter and get rid of any liquid in the tube below. Now reinsert your spin filter and add 200uL of 1X buffer, then pipette up and down just over the filter (five to six times) to recover any bound origami on the filter. Be very careful not to puncture the filter during this step. Finally, add another 200uL of buffer to your sample to bring the final volume to ~400uL. + - + - 9. Spin at 100g for ~20 minutes at 4°C. This should leave you with ~70-100uL of sample retained in the spin filter. Perform the above pipetting step again to recover as many origami as possible from the spin filter. + - + - 10. Remove the spin filter, and place it upside down in a fresh tube (being careful not to dump your sample on the ground!). You can now spin your sample in a microcentrifuge for 12 minutes to collect everything. + - + - + - *Paul W.K. Rothemund's additional notes: + - + - A similar spin protocol appears necessary for binding DNA origami to sticky patches on silicon. Without it, the staples seem to interfere in binding. This protocol was originally taken from Rob Barish's protocol for origami/ribbon constructs. I think that some parts of the protocol may not be necessary for regular origami. Notably, I'm not sure that pipetting up and down is necessary, except perhaps at the last step to maximize recovery. Similarly the use of a 4C centrifuge seems unnecessary, the centrifugation may go much faster at room temperature. If you use a different centrifuge or temperature, calibrate the passage of buffer through the filter before you try purifying any origami to get an idea of how much buffer is retained in the filter after a spin. Fewer spins will be required if a smaller excess of staples is used. You should be able to get away with 10X fewer staples than we typically use according to my standard origami protocol, easily. That is, a 5-10X stoichiometric excess of staples over scaffold is probably fine and will result in cleaner origami faster. + - + - ===Opening SPEX Data in MATLAB=== + - *Open up MATLAB and use File->Import Data + - *Navigate to the raw-text (.txt) data file and open it + - *The table on the right should show eight rows, where the odd ones are the times associated with the signal from the row under them. The top set is sample 1; the next is 2, the next is 3, the bottom set is sample 4. Press "next." + - *Optionally right click on "data" and rename it to whatever you want. This isn't necessary unless you're afraid of your data getting overwritten eventually when you import another set. + - *Click "finish." + - *Your data should be visible in the "workspace" window with the name "data" or whatever you might have renamed it to. Right click on it -> "open selection" to view it. + - *To plot a single sample's data, use this, where "data" is the name of your data variable: + - :plot(data(1, :), data(2, :)) + - *To plot all four samples' data on the same plot, use this: + - :plot(data(1, :), data(2, :), data(3, :), data(4, :), data(5, :), data(6, :), data(7, :), data(8, :)) + ===Normalizing SPEX Data in MATLAB=== ===Normalizing SPEX Data in MATLAB=== - *Upon opening the data, use the following code, replacing "start" with the beginning of the experiment (for origami experiments, this will usually be immediately after the walker trigger is added), and replacing "finish" with the data point corresponding to the reaction reaching its completion level. + :''More detailed article: [[Biomod/2011/Caltech/DeoxyriboNucleicAwesome/Protocols/MATLAB|MATLAB Analysis]]'' - *Your data should be visible in the "workspace" window with the name "data" or whatever you might have renamed it to. Right click on it -> "open selection" to view it. + - *The data is normalized such that the start of the reaction is at 1, and the completion level is at 0, by translating the completion level down, and then scaling the data. + - :time1 = data(1, start:finish); + - :time2 = data(3, start:finish); + - :time3 = data(5, start:finish); + - :time4 = data(7, start:finish); + - :data1 = data(2, start:finish); + - :data2 = data(4, start:finish); + - :data3 = data(6, start:finish); + - :data4 = data(8, start:finish); + - :time1 = time1 - time1(1); + - :time2 = time2 - time2(1); + - :time3 = time3 - time3(1); + - :time4 = time4 - time4(1); + - :ndata1 = data1 - data1(finish - start + 1); + - :ndata2 = data2 - data2(finish - start + 1); + - :ndata3 = data3 - data3(finish - start + 1); + - :ndata4 = data4 - data2(finish - start + 1); + - :ndata1 = ndata1/ndata1(1); + - :ndata2 = ndata2/ndata2(1); + - :ndata3 = ndata3/ndata3(1); + - :ndata4 = ndata4/ndata4(1); + - :plot(time1, ndata1, time2, ndata2, time3, ndata3, time4, ndata4); + - {{Template:DeoxyriboNucleicAwesomeFooter}} +

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Protocols

Oligonucleotides

Most of the sequences were from Lulu Qian’s sequences, designed as described in [1]. Modifications were added using a computer program NUPACK, and staple sequences for the rectangular origami were taken from Rothemund’s [7]. All oligonucleotides were ordered from Integrated DNA Technologies (IDT) Inc. Most of the DNA molecules were ordered standard desalted and unpurified, and modified molecules are ordered HPLC purified. Staple strands were ordered in wet plate form at 150μM in 1x TE buffer. Dried DNA was reconstituted in an appropriate amount of ddH2O, and quantified with the biophotometer.

More detailed article about quantification protocol : Quantitation

Buffer conditions

1x TE/Mg2+ (TE=Tris-EDTA, Tris 40 mM, EDTA 2 mM, and Magnesium acetate, 12.5 mM, pH 8.0) buffer was used for all experiments.

Gel Electrophoresis/ Gel Purification

5% polyacrylamide gels of 1 mm in thickness were used. Native gel was polymerized by mixing 3.75 ml 19:1 40% polyacrylamide, 1 mL 10x TAE/Mg++, 5.25 ml ddH2O, 60 μL 10% APS, 6 μL TEMED. Native gels were run at 150V at 25 ºC. 1x TAE/Mg2+ was used as a running buffer. Gels were stained with SybrGold for 15 min and then scanned with Quantity 1 FX scanner. Some complexes were annealed using PCR machine and gel purified using bigger gel. Same recipe was used to make native gel. Glycerol was used as a loading dye for the modified strands. Under UV light, the bands were cut and DNA was obtained by diffusion in 1x TE/Mg2+ buffer for 48 hours.

More detailed article about gel puriciation protocol: Gel Purification

DNA origami formation

Rectangular DNA origami consists of M13 viral DNA and 202 DNA staples. Staples at the side of the origami were not included in the 202 staples to avoid the stacking problem. Sometimes, a probe at the walker start site was intentionally left out, making a hole in the middle of the origami. M13 scaffold DNA and staple strands were mixed in 1x TE/Mg2+ buffer with molar ratio of 1:4 between the M13 and staple strands. For the speed of assembly, origami were annealed at 90nM, and diluted into different final concentration for the purpose of various experiments; 15 nM of M13 were used for fluorescence experiments, and 10 nM, 5 nM, or 1 nM for or AFM experiments. Origami was annealed by heating up to 90 ºC then cooling down to 20 ºC at 1 ºC/min using an Eppendorf PCR machine. After annealing, tracks were added to the annealed origami at 1:6 ratio of M13 to tracks. Walker goal and a walker start complex which is a preannealed complex “walker – walker inhibitor – track1 – probe for track 1” were also added to the origami at 1:1 ratio of M13 to strands. Mixed solution was incubated at 37 ºC overnight.

More detailed article about origami puriciation protocol: Origami Purification

Atomic force microscopy (AFM)

5 μL of sample was deposited onto a freshly cleaved mica, and 20 μL of 1x TE/Mg2+ buffer was added. 40+ μL of 1x TE/Mg2+ buffer was added to the fluid cell and the sample was scanned in a tapping mode with DNP tips. When streptavidin was used, 1 μL of streptavidin was added to the mica, and we waited 15 or more minutes before imaging to ensure the streptavidin and biotin could bind. The exception to this was when we used the Walker Lock-Down Protocol.

Walker Lock-Down Protocol

To see what this protocol was used for see the AFM experiments page. (When we tried this protocol, the concentration of origami was too high, so we tried using 2 μL of sample instead of 5 μL, but in the future we will dilute the sample before using it, and we will stick to using 5 μL of sample.) The sample, buffer, and streptavidin are mixed on mica as above, but after waiting 15 minutes, we do the following:

1. add 20 μL of 1x TE/Mg2+ buffer to mica, then remove as much liquid as possible (40+ μL) from the mica without touching the mica.

2. add 40 μL of 1x TE/Mg2+ buffer to mica, then remove as much liquid as possible. Repeat this 3 times. (In the future we may repeat this more times to remove as much excess streptavidin as possible.)

3. add 20 μL of 1x TE/Mg2+ buffer to mica.

4. add biotinylated† walkers at a 1-to-1 ratio to the tracks or higher (excess should not affect the results, and when implemented a 1-to-1 ratio would require a volume that is too small to be pipetted, so excess is likely necessary.)

5. wait for 15+ minutes before imaging sample.

†We have only seen the walkers when 5'-end biotinylated walkers were used, but 3'-end biotinylated walkers were only tried at a low concentration.

Based on our preliminary results using this protocol, streptavidin binds to mica, and if it is ever lifted off of the mica, it can bind to walkers on origami. Thus, we will try using the above protocol, but using 1x TE/Mg2+ with 100 μM Na+ instead to minimize the amount of streptavidin that binds to mica.

Spectrofluorometer (SPEX)

After cuvettes are cleaned with ddH2O and 70% ethanol, 10 nM - 1 uM sample was loaded in 1.5mL 1x TE/Mg2+ buffer into each cuvette for kinetics experiment in solution. For random walking experiment on origami, 200 ul of 15 nM samples were used. Fluorescent level of each cuvette was observed in real time.

More detailed article about SPEX protocol: SPEX

Normalizing SPEX Data in MATLAB

More detailed article: MATLAB Analysis