Lukeman Lab
Presented by Team Toehold Conga Nanny
USING DNA ORIGAMI FOR
A NANOSCALE ELECTROCHEMICAL POSITIONER MODEL
Presented by Team Toehold Conga Nanny
Buffer Preparation
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10xHEPES buffer (200 mM MgCl2, 50 mM NaCl, 200 mM HEPES): add 166.67 mL of 1.2 M MgCl2, 50 mL of 1 M NaCl, 200 mL of 1 M HEPES into a 1000 mL volumetric flask and fill with double distilled water.
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10xHEPES buffer (150 mM MgCl2, 50 mM NaCl, 200 mM HEPES): add 125 mL of 1.2 M MgCl2, 50 mL of 1 M NaCl, 200 mL of 1 M HEPES into a 1000 mL volumetric flask and fill with double distilled water.
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10xHEPES buffer (100 mM MgCl2, 50 mM NaCl, 200 mM HEPES): add 81.33 mL of 1.2 M MgCl2, 50 mL of 1 M NaCl, 200 mL of 1 M HEPES into a 1000 mL volumetric flask and fill with double distilled water.
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10xHEPES buffer (50 mM MgCl2, 50 mM NaCl, 200 mM HEPES): add 41.66 mL of 1.2 M MgCl2, 50 mL of 1 M NaCl, 200 mL of 1 M HEPES into a 1000 mL volumetric flask and fill with double distilled water.
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10xHEPES buffer (0 mM MgCl2, 50 mM NaCl, 200 mM HEPES): add 0 mL of 1.2 M MgCl2, 50 mL of 1 M NaCl, 200 mL of 1 M HEPES into a 1000 mL volumetric flask and fill with double distilled water.
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1xHEPES buffer (20 mM MgCl2, 5 mM NaCl, 20 mM HEPES): pour 100 mL of 10xHEPES buffer (200 mM MgCl2, 50 mM NaCl, 200 mM HEPES) into 1000 mL volumetric flask and fill it up with double distilled water.
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1xHEPES buffer (15 mM MgCl2, 5 mM NaCl, 20 mM HEPES): pour 100 mL of 10xHEPES buffer (150 mM MgCl2, 50 mM NaCl, 200 mM HEPES) into 1000 mL volumetric flask and fill it up with double distilled water.
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1xHEPES buffer (10 mM MgCl2, 5 mM NaCl, 20 mM HEPES): pour 100 mL of 10xHEPES buffer (100 mM MgCl2, 50 mM NaCl, 200 mM HEPES) into 1000 mL volumetric flask and fill it up with double distilled water.
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1xHEPES buffer (5 mM MgCl2, 5 mM NaCl, 20 mM HEPES): pour 100 mL of 10xHEPES buffer (50 mM MgCl2, 50 mM NaCl, 200 mM HEPES) into 1000 mL volumetric flask and fill it up with double distilled water.
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1xHEPES buffer (0 mM MgCl2, 5 mM NaCl, 20 mM HEPES): pour 100 mL of 10xHEPES buffer (0 mM MgCl2, 50 mM NaCl, 200 mM HEPES) into 1000 mL volumetric flask and fill it up with double distilled water.
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1X TBE buffer: pour 100 mL of 10X TBE buffer into 1000 mL volumetric flask and fill with double distilled water.
*Note that pH of all 1x buffers was in the range 6.86-7.02.
Gel Pouring
Agarose Gel
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Place a clean 600 mL beaker on a scale and tare it.
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Weigh out 1.5 g of agarose powder into the beaker.
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Add 1xHEPES (5 mM MgCl2, 5 mM NaCl, 20 mM HEPES) buffer until the balance shows 150 g. That creates 1% agarose solution.
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Mix the solution and microwave for 2 minutes and 45 seconds.
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Cool the beaker in an ice water bath. Keep swirling the beaker to prevent polymerization. Add the thermometer and keep checking the temperature.
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When the temperature reaches 45 °C, take the beaker out of the ice bath and pour the contents of the beaker into the gel cassette and insert the clean comb.
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Use a glass Pasteur pipet to push any bubbles to the corners of the gel.
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Leave the cassette covered and let the assembly sit for at least sixty minutes to complete ensure complete polymerization.
Acrylamide Gel
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Prepare a gel cassette using two glass plates and two clamps.
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Firmly place the gel cassette into place on the gel-holding apparatus.
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Prepare the gel by mixing 22.5 mL of 20% gel stock and 2.5 mL of 0% gel stock.
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Prepare the ammonium persulfate (APS – 10% w/v water) in a 10 mL tube and TEMED solution.
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Add the gel stock, 160 uL of 10% APS, and 16 uL of TEMED into a 50 mL tube.
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Pour the gel using the large plastic pipet and insert a 20-lane comb between the plates. Be sure to carry this out quickly because the gel mixture polymerizes quickly. Pouring the gel is most efficient when the pipet is held at about 30 degrees below the vertical, perpendicular to the plane of the plates.
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Let the entire assembly sit for at least sixty minutes to ensure complete polymerization.
Origami Sample Preparation
There are three phases to the preparation of origami samples, which can be performed at different times if the solutions are stored frozen in-between steps. These phases are (1) staple stock solution preparation, (2) preparation of the complete origami solution, and (3) the anneal.
Preparation of Staple Stocks
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Take the origami staple strand plates from the freezer and allow it to defrost to room temperature. This generally takes 30 minutes. (Note: The concentration of these strands are each 100 uM.)
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If necessary, spin the plates in the salad spinner to form all the strand solution to the bottom of the wells.
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Prepare three boxes of long-reach 10 uL pipet tips.
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Refer to the oligo order forms to know which wells to avoid while pipetting strands out.
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Set a volume on the pipet, depending on how much volume you want to be there in stock solution.
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Remove the top from one of the 96-well plate and carefully place it aside. Place the plate and one of the prepare tip boxes next to each other such that the columns line up.
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For each column of tips, carefully pipet from the corresponding column of the 96-well plate to the previously marked destination column of your empty plate and then discard those tips into the sharps bin. Watch carefully to ensure that each tip actually withdraws and deposits fluid. (Note: Some wells are supposed to be empty; consult with the order forms to determine which.)
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When finished pipetting the 96-well plate, replace the cover and place to the side.
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Repeat the last three steps for the remaining strand plates.
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Replace the cover on the destination plate or close the lid of the tube (depending on where you have pipetted your strands).
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If you’re using a destination plate, transfer all the solutions from the destination column to a 2 mL centrifuge by using a 100 uL pipet.
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Fill the 2 mL centrifuge tube to the desired volume with (0.22 microfiltered) deionized water. Label the tube with the type of staple stock, the date, and concentration.
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By using the steps above, finish making the staple stocks based on the order forms.
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Clean up and return strand plates and staples stocks tubes to the freezer.
Preparation of Origami Solutions
Produces solution at 25:5:1 Dye (methylene blue and/or anthraquinone) to Staple to Plasmid Ratio.
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Take the staple stocks and one or two tubes of plasmid stock from the freezer and allow it to defrost at room temperature.
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While the stocks are defrosting, prepare a 2 mL tube for each type of origami being made and label with the type and date. Assume 5 pmol of 20 nM positioner is made:
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12.5 uL of 0.4 uM plasmid is added to a 2 mL tube for each type of origami.
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43.9 uL of 0.57 uM core staples, 2 uL of 12.5 uM rep arm staples, 2 uL of 12.5 uM thiol anchor staples, 2 uL of 12.5 uM needed adjuster staples, 1.3 uL of 20 uM rep base staples, 5 uL of 25 uM all dyes stock staples are added into tubes for each type of origami.
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Since total volume needs to be 250 uL, we fill up each tube with 25 uL of 10xHepes (150 mM MgCl2) and 156.3 uL of 1xHepes (15 mM MgCl2).
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After completion of the above steps, close the tubes, vortex, spin them down and start the anneal.
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Place stock tubes back to the freezer.
Annealing Origami
Slow anneals using a hot water bath
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Fill a 2L beaker with tap water. (Using hot water would save time.)
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Place the beaker on the hot plate, turn the intensity to 10, and insert a thermometer.
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Wait for the thermometer to reach 90°C.
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While waiting, insert each reaction tube of interest into a 50 mL tube. To be efficient, insert up to three reactions tubes into a single 50 mL tube.
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Insert each of these 50 mL tubes into a blue flotation device.
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When the water reaches 90°C, stir the contents with a thermometer and test the upper middle part of the water. (Note: The beaker is typically hotter at the base.)
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When the upper middle part of the water is between 90°C – 94°C, turn off the hot plate, remove the beaker and place it into a big Styrofoam box.
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Insert the flotation device with the 50 mL tubes and weigh it down with an empty glass tray. Cover the box with the Styrofoam lid. Wait for the water to reach room temperature (we leave it to anneal overnight).​
Destapling Annealed Origami
Concentrated Samples (Low Volume)
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To wash the filter, place 500 uL of appropriate for your sample buffer into 100 kDa centrifugal filter tube and spin at 5xg for 5 minutes.
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Then, place 450 uL of the sample into the filter tube and spin at 5xg for 2 minutes.
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To wash the sample, load 450 uL of the same buffer into the filter tube and spin at 5xg for 2 minutes. This step has to be done 3 times.
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After the last buffer wash, invert the filter into a new test tube and spin it at 1xg for 2 minutes.
Dilute Samples (Bigger Volume)
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To wash the filter, place 3500 uL of appropriate for your sample buffer into 100 kDa 4mL centrifugal filter tube and spin at 5k rcf for 5 minutes.
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Then, place 4000 uL of the sample into the filter tube and spin at 2.5k rcf for 1 minute.
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To wash the sample, load 3450 uL of the same buffer into the filter tube and spin 2.5k rcf for 1 minute. This step has to be done 3 times.
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After the last buffer wash, get the sample out of the filter by pipetting it out to a new test tube.
Gel Imaging
Agarose Gel
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After running the gel, carefully remove the plugs from the power supply.
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Prepare a staining solution using Sybr Gold stain by dissolving 20 uL of Sybr Gold in 200 mL of double distilled water. Mix the solution gently and leave it for 30 minutes prior to usage.
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Gently take the gel out of the gel tray and place it into the box with Sybr stain for an hour. For better efficiency, place the box into the Shake’ N’ Bake.
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To image the gel, carefully lift and place the gel into the box with double distilled water for safe transportation to the Typhoon instrument.
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Open Typhoon program and click fluorescence, then for method pick Atto647NAcceptor for a pre-stain gel and pick LukemanSybr1 for stained gel. Those methods have proper wavelengths for the methylene blue to appear on the images.
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The instrument generates .tiff and .gel files that we analyze later by using Acorn (used for color inversions and annotations) and Image J (used to find the intensities of the bands, yields, and concentrations of samples) programs.
Acrylamide Gel
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After running the gel, carefully remove the plugs from the power supply.
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Prepare a staining solution using SYBR Gold stain by dissolving 20 uL of SYBR Gold in 200 mL of double distilled water. Mix the solution gently and leave it for 30 minutes prior to usage.
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Gently take the gel out of the gel cassette and place into the box with SYBR Gold stain for 15 minutes.
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To image the gel, carefully lift and place the gel into the box with double distilled water for safe transportation to the Typhoon instrument.
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Open the Typhoon program and click fluorescence, then for the method, pick LukemanSybr1 for the stained gel. This method has the proper wavelengths for the strands to appear on the images.
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The instrument generates .tiff and .gel files that we analyze later by using Acorn (used for color inversions and annotations) and Image J (used to find the intensities of the bands, yields, and concentrations of samples) programs.
Atomic Force Microscopy
AFM Set-up
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Items required:
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Freshly filtered (through a 0.2 um filter) water
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AFM samples
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Small round adhesive stickers
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Pucks
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Sheet of mica
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Hole Puncher
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1-10 uL pipettors and 100-1000 uL pipettor with their tips
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Can of dust remover spray
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AFM tips/ tip holder
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Multimode Nanoscope with software
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Scanner
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Laser head
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Put the Multimode Nanoscope on the microscope stage
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Fit and connect the appropriate scanner to the Multimode Nanoscope
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Properly place the laser head onto the scanner and connect it to the Multimode Nanoscope
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Secure the laser head by attaching the springs from the Nanoscope onto the sides of the laser head
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Attach the cable from external AFM controller to the AFM
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Turn on computer
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Turn on the external AFM controller
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Choose appropriate options according to AFM software that is being used
Calibrating AFM
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After mounting the tip onto the tip holder, place into the laser head
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Using the knob on the back (cantilever holder clamp), lower the clamps onto the tip holder until a slight resistance is felt
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Use camera attached and knobs on the stage to focus on the cantilever
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Focus the laser on the cantilever using the knobs on the laser head
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Maximize the signal on the photodiode while switch is on AFM/LFM mode
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Adjust the mirror on the back of the laser head
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Move the laser to reposition it on the cantilever
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Using the photosensor knobs, get the RMS and VERT values as close to zero on the Multimode Nanoscope
Sample Preparation
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Take a puck and apply an adhesive sticker to provide a sticky surface for the mica
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Hole punch a piece of mica and place it on the [sticky] puck
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Press the mica onto the puck by the edges and hold for 1 minute
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Nothing must touch the center of the mica
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Now that the mica is securely placed on the puck, hold the puck with tweezers
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Take a small piece of tape, place on the mica and peel off
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Lift up a piece of tape to the light to make sure that there are no cracks on the mica
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If there are cracks on the tape, use a new piece of tape on the mica
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Continue to place and peel tape on the pica until there are no noticeable cracks on the tape
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Pipet origami solution (about 3-5 uL) onto the center of the mica
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Allow solution time to set (30 seconds to 1 minute)
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Take 1 mL of filtered deionized water and gently squirt small portions onto the mica surface
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Take the edge of a Kimwipe and wick the edges of the mica to soak up excess water
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To dry the mica surface, grab a dust remover spray and gently blow on the mica for 1-2 minutes
Loading Sample
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Raise the tip up by using the right AFM switch and flipping it forward for about 10 seconds
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Remove the laser head carefully by removing springs
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Carefully put mica onto the stage
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Put laser head back on and secure it in place with springs
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Lower the tip back down by using the same AFM switch as in the first step and flipping it backward for about 10 seconds
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Make sure that all settings on the software correlate with the type of mode that is planning to be used
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Follow instructions of the Multimode Nanoscope that is being used to engage the tip to begin scanning the sample
AFM Imaging
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Once scanning begins, adjust both the integral and proportional gains. Raising both gains will allow for better resolution. Make sure gains are not too high (feedback). Proportional gain must always be higher than the integral gains
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If needed, lower the amplitude setpoint voltage to make the cantilever scan with a greater force so that lift-off does not happen
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Adjust scan angle and scan speed (usually 2Hz to start with) to maximize image resolution
Measuring Positioner Angles (in TEM images)
We made 9 different versions of the positioner, each with a different adjuster length between the base and the arm. The different lengths create different angles between the base and the arm than can measure. With increasing adjuster lengths, we hoped to see an increase in angle size for the positioner.
The first point to note when we measure angles is to know what positioner to choose. The criteria for choosing the correct positioner is
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The positioner has to be well-folded and oriented; no oligomers
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Both arms and the adjuster helices must be able to be seen
To measure the angle of positioners, the angle was drawn down the midline of the base and arm of the positioner
This is an example of a TEM image of our positioner
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Figure 1- The positioners numbered 2, 5, and 6 are good examples of positioners to take angles of based on the above criteria. For positioner 1, the adjuster helices are not clearly defined. Positioner 3 is misfolded. Positioners 4 and 8 are oligomerized. For positioner 7, both arms are not shown.
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Figure 2- This is how the angles for all of the positioners were drawn
Electrochemistry
Chip Cleaning
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Place freshly diced gold chips on inverted glass vials
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Squirt each gold chip with HPLC-grade acetone for 1 minute
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Squirt each gold chip with hplc-grade 2-propanol for 1 min
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Place gold chips on a clean paper towel and allow them to air dry for 10 minutes
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Insulate the gold surfaces from the copper tape by placing an adhesive-coated and perforated silicone gasket on top of the substrates.
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Make an electrical contact to the gold chip using copper tape
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An electrochemical cell is assembled by inserting our modified working electrode between a poly(tetrafluoroethylene) cell top and a cell bottom which are screwed together. The top includes a hole that exposes just the gold surface, a built-in counter electrode, compartment to insert a reference electrode, and a well that can be filled with a liquid substance which electrochemical experiments will be carried out in (ex. buffers)
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The gold surface is then electrochemically cleaned
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1mL of 0.5M sulfuric acid added to the cell
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Substrates polarized to 1.9 V vs Ag/AgCl and swept back to 0.0 V. This is repeated for 30 cycles
*Before interrogating the chips by depositing positioner sample or monolayer solutions, the chips are gently rinsed 3 times with 1 mL of 5mM MgCl2 1x HEPES
Electrochemistry Deposition
* Refer to LukemanLab Echem spreadsheet*
Note: Positioner samples were either reduced with 10 mM TCEP before or after destapling the sample (refer to column P).
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Procedure 1
A clean chip was treated with ~10 uL of positioner sample (refer to column C) which was deposited for a variable amount of time (refer to column E). Carefully remove positioner sample and deposit ~50 uL of 5 mM MCH such that the entire surface area of the gold is saturated. Let the MCH deposition incubate on the gold surface for a variable amount of time (refer to column G). Carefully remove the MCH from surface of chip, avoiding contact between the chip and pipette. Carefully rinse chip 3 times with 1 mL of 5mM MgCl2 1x HEPES. All incubations done in a sealed container containing water to create a humid environment, placed in a refrigerator.
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Procedure 2
Deposit ~50 uL of 1 mM HDT to a clean chip and let the deposition incubate for a variable amount of time (refer to column G). Carefully remove HDT sample and deposit ~10 uL of positioner sample (refer to column C) such that the entire surface area of the gold is saturated. Let the positioner sample deposition incubate on the gold surface for a variable amount of time (refer to column E). Carefully remove the positioner sample from surface of chip, avoiding contact between the chip and pipette. Carefully rinse chip 3 times with 1 mL of 5mM MgCl2 1x HEPES. All incubations done in a sealed container containing water to create a humid environment, placed in a refrigerator.
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Procedure 3
Deposit ~10 uL of a mixture of MCH and positioner sample (refer to column C) and let deposition incubate for a variable amount of time (ref to column E). The mixture was prepared such that the final concentration of TCEP in the sample would be 1 mM. Carefully remove the mixture of positioner sample and MCH from surface of chip, avoiding contact between the chip and pipette. Carefully rinse chip 3 times with 1 mL of 5mM MgCl2 1x HEPES. All incubations done in a sealed container containing water to create a humid environment, placed in a refrigerator.
NHS Ester Modified Amino-Labeled Oligonucleotide Reaction
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Dilute commercially bought strands to 100 uM with double distilled water in 2 mL DNA LoBind tubes.
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Take a quarter of the total volume of each strand using a 100-1000 uL pipet and transfer to a separate, labeled 2 mL tube. Centrivap till a pellet forms in the tube.
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Prepare a 0.25 M NaHCO3/Na2CO3 buffer with a pH of ~8.99.
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Dissolve the sample strands in the 0.25 M NaHCO3/Na2CO3 buffer to 100 uM. The volume added is equivalent to the volume of sample taken out in step 2.
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Prepare 100 mM of NHS Ester dissolved in DMSO.
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Based on the 100 mM concentration of NHS Ester and the number of moles of strands used, calculate the volume of NHS Ester that would hold 200 times the picomoles of the strand.
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Add the individual volumes for each strand from step 6 into each strand tube.
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Vortex the tubes and allow them to sit at room temperature for 1 hour covered in foil or in a dark environment to react.
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After 1 hour, repeat step 7 and allow them to sit and react at room temperature covered in foil overnight.
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Store samples in the freezer the following day till use.
HPLC
In order to prepare our samples for our HPLC runs, we filtered our samples through 0.2 uM filters and took out 100 uL aliquots of each strand we wanted to test. To determine the best conditions for our samples’ runs on the HPLC, we tested four different conditions with our c18 column on one of our strands. The gradients we tested were 5% - 15%, 5% - 25%, 5% - 35%, and 5% - 45% Acetonitrile with buffer A as TEAA (0.1 M, pH 7) and buffer B as CH3CN, and, based off our 40-minute-long runs, the best gradient was the 5 - 45% Acetonitrile. After running these tests, we also shortened the run time down to 20 minutes with 10 minutes of column washing, as depicted on the HPLC spectrums.