- 1X TBE: Add 50 mL of 10X TBE in a 500 mL volumetric flask. Add D.D. water until the final volume is 500 mL.
- 0.5X TBE: Add 25 mL of 10X TBE in a 500 mL volumetric flask Add D.D. water until the final volume is 500 mL.
- 1X TAEMg: Add 50 mL of 10X TAEMg in a 500 mL volumetric flask. Add D.D. water until the final volume is 500 mL.
- 10X TAE: Dissolve 28.6 g of magnesium acetate with 1 L of a 10X TAE mixed together on a stir plate for 10 – 20 minutes.
- The gel cassette is assembled with two glass plates and two clamps.
- Be sure that the gel cassette is firmly held into place on the gel-holding apparatus.
- Prepare the gel with the 20ml of mixture of 20% gel stock and 0% gel stock.
- Prepare the ammonium persulfate (APS - 10% w/v water) and TEMED solution. For APS, the estimated volume should be 150 – 180 uL. For TEMED, the estimated volume should be 15 – 18 uL.
- Add the gel stock, APS, and TEMED into a 50 mL tube.
- Pour the gel using the large plastic pipet and insert the comb between the plates. Be sure to carry this out quickly because the gel mixture polymerizes quite quickly. Pouring is often most efficient when the pipet is kept at about 30 degrees below vertical, perpendicular to the plane of the plates.
- Let the entire assembly sit for one hour to ensure complete polymerization.
- Place 1.2 g of agarose in a clean 600 mL beaker on a scale.
- Prepare the 0.5X TBE buffer as described.
- Add 30 mL of D.D. water.
- Microwave for 2 minutes.
- If the solution is not clear, swirl and microwave for another 15 seconds.
- Cool the beaker in an ice water bath. Keep swirling the beaker to prevent polymerization. Add the thermometer and keep checking the temperature.
- When the solution reaches 60°C, remove it from the ice water bath.
- Add 1 mL of 1.2 M magnesium chloride.
- Pour the contents of the beaker into the gel cassette and insert the comb.
- Use a glass Pasteur pipet to push any bubbles to the corners of the gel.
- Leave the cassette covered and let the assembly sit for thirty minutes to complete ensure complete polymerization.
- Set to nucleic acid measurement mode.
- Clean the metal ball on the pedestal with a kim wipe.
- Pipet 1.5 uL of D.D. water onto the small metal ball, and blank.
- Clean the metal ball on the pedestal with a kim wipe.
- Pipet 1.5 uL of the sample onto the small metal ball, and measure.
- Read the absorbance at 260 nm.
- This is done in triplicate and the average is obtained.
VLP Buffer Exchange
- From the VLP stock solution, pipet an amount sufficient for the entire experiment into a 50 kDa centrifugal filter tube.
- Note: Make sure to return the VLP stock solution to the fridge.
- Fill the centrifugal filter tube up to 500 uL with 1X TAE-Mg and close the lid tightly.
- Place the centrifugal filter tube into the centrifuge and spin at an RCF of 5 for 6 minutes.
- Dispose the waste solution at the bottom of the centrifugal filter tube.
- Repeat steps 3–5 twice more.
- Spin for additional time as needed to reduce the final volume.
VLP/Origami Binding Anneal
- Exchange VLP buffer to 1X TAEMg.
- Pipet origami and VLP into PCR tube in the desire ratio.
- Add the nonstandard components as needed for a particular experiment.
- Add buffer to bring anneal to the desired concentration for a particular experiment.
- Use a microcentrifuge to collect he sample at the bottom of the PCR tube.
- Place the PCR tubes for the samples in the PCR machine and run the anneal protocol. (The standard anneal for binding was from 37 to 4 deg C, at 1 deg C per minute.)
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
- 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 solutions are each 100 uM.)
- If necessary, spin the plates in the salad spinner to form all the strand solution to the bottom of the wells.
- Find an un-used lane on an empty 96-well plate. Label it accordingly to indicate that it is in use.
- Prepare three boxes of long-reach 10 uL pipet tips.
- Depending on the type of origami, remove the tips from each of these three boxes and store or discard. Refer to the oligo order forms for which tips to remove – claw will remove tips 5B and 8B, while triangles will remove tips 5A, 5B, 5C, 5E, and 5G, where A – H represent rows and 1 – 12 represent columns.
- Set an 8-well pipetter to 8 uL.
- 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.
- 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.)
- When finished pipetting the 96-well plate, replace the cover and place to the side.
- Repeat the last three steps for the remaining strand plates.
- Replace the cover on the destination plate.
- Using the salad spinner, centrifuge the destination plate to move all solution to the bottom of the well.
- Using a 100 uL pipet transfer all the solution from the destination column to a 2 mL centrifuge tube. Carefully check the bottom of each well to ensure that most of the solution is extracted. When finished, mark the lane of the plate as used.
- Fill the 2 mL centrifuge tube to the 2 mL with (0.22 microfiltered) deionized water. Label the tube with the type of origami (CLAW or TRIANGLE), the date, and “General Staples: 04 uM.”
- Using a 10 uL pipet (and a different tip for each well), pipet 8 uL of each of the wells skipped over in (step 5) into another 2 mL centrifuge tube. Fill this tube to the 2 mLmark with filtered D.D. water.
- Label the tube with the type of origami (CLAW or TRIANGLE), the date, and “Blunt Staples: 0.4 uM.”
- Using a 10 uL pipet (and a different tip for each tube), pipet 8 uL of each of the stick strands into another 2 mL centrifuge tube. Fill this tube to the 2 mL mark with filtered D.D. water.
- Label this tube with the type of origami (CLAW or TRIANGLE), the date, and “Sticky Staples: 0.4 uM.”
- Clean up and return strand plates and sticky strand tubes to the freezer.
Preparation of Origami Solution
Produces solution at 50:1 Staple to Plasmid Ratio, 4 nM Concentration of Plasmid, 1X TAEMg, and total 250 uL of solution per picomole (Note: This protocol assumes 2 pmol of origami are being prepared, and gives volumes to produce that quantity.)
- Take the staple stocks and one or two tubes of plasmid stock from the freezer and allow it to defrost at room temperature. (Note: The NEB stocks are at 0.1 uM.
- While the stocks are defrosting, prepare a 2 mL tube for each type of origami being made and label with the type and date.
- Pipet 20 uL [or 10X] NEB plasmid into each destination tube.
- Pipet 250 [or 125X] uL of 0.4 uM “General Staples” for the appropriate origami design into each destination tube.
- Pipet 250 [or 125X[ of 0.4 uM “Sticky Staples” for the appropriate origami design into each destination tube of STICKY origami.
- Pipet 250 [or 125X] of 0.4 uM “Blunt Staples” for the appropriate origami design into each destination tube of BLUNT origami.
- Pipet 50 uL [or 25X] of (0.22 micro filtered) 10X TAEMg into each destination tube.
- Pipet 50 uL [or 25X] of deionized water into each destination tube.
- Return the remaining staple and plasmid stocks to the freezer.
There are two different methods we use to anneal origami samples; either produces acceptable results for our purposes.
Slow anneals using a hot water bath
- Fill a 2L beaker with tap water. (Using hot water would save time.)
- Place the beaker on the hot plate, turn the intensity to 10, and insert a thermometer.
- Wait for the thermometer to reach 90°C.
- While waiting, inset each reaction tube of interest into a 50 mL tube. To be efficient, double up and insert two reactions tubes into a single 50 mL tube.
- Insert each of these 50 mL tubes into a blue flotation device.
- 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.)
- When the upper middle part of the water is between 90°C – 94°C, turn off the hot plate, remove the beaker, insert the flotation device with the 50 mL tubes, and weigh it down with an empty glass tray. Wait for the water to reach room temperature. (This will take about four hours, and can be left overnight if needed.)
Rapid anneals using a thermocycler
- Split the origami solution into small PCR tubes. (Note: Put no more than 80 uL in a single tube. Also, use a color-coding system to keep track of the different samples to avoid having to label all of these tubes.)
- Insert the tubes into a PCR machine. The OpenPCR machines will fit 16 tubes at a time.
- Run the appropriate program to begin annealing (eg "CLAW-53deg-15min).
De-stapling Annealed Origami/Origami Buffer Exchange
- Transfer annealed origami solution to 500 uL 50kDa centrifugal filters.
- Top the solution up to 500 uL with 1X TAEMg.
- Spin at 5,000 g’s for 10 min.
- Remove and collect filtrate in separate tube.
- Fill the filter cup back up with more 1X TAEMg.
- Repeat steps 3-5 for a total of 5 buffer exchanges.
- After the 5th exchange, invert filter cup into a new tube (included in the box of centrifugal filters).
- Spin at 1,000 g’s for 2 min.
- Add 25 uL of 1X TAEMg into the filter cup and re-invert cup into the same tube that now holds the concentrated origami solution.
- Spin again at 1,000 g’s for 2 min.
- Transfer final origami solution to appropriate tube for use/storage.
Cryo-protecting VLP’s with Trehalose
- Separate the VLP solutions into the desired aliquots.
- Make a 1 M solution of trehalose.
- To each of the desired aliquots, add 500 mol equivalents of trehalose.
- In a vacuum flask, add enough dry ice to acetone until the dry ice no longer bubbles/fizzes (i.e. until it is at a constant temperature. Caution: Solution is at an extremely low temperature.
- Hold the tube with the aliquoted VLPs and added trehalose in the dry-ice-acetone bath for about 1 min to quick-freeze the solution.
- Store tube(s) in -20 °C freezer for long term storage until needed.
Removing Trehalose from Cryo-protected VLPs
Follow previously mentioned buffer exchange protocols with these updates
- Use 500 uL 3kDa centrifugal filters.
- Use 50 mM sodium phosphate 150 mM NaCl (pH 6.38) as the buffer used to the the buffer exchange.
- Buffer exchange spin parameters: 10,000 g’s for 5 min
- Spin for at least 4 full exchanges of buffers (may need more than 1 spin to get to a low enough volume to count as a full exchange).
- Invert filter cup spin parameters: 1,000 g’s for 2 min
- After trehalose has been removed via buffer exchange, store VLP solutions in 4 °C fridge for use/short term storage.
Modifying Staple Strands to Fluor-tagged Staple Strands via Click Chemistry (CuACC)
The Baseclick Oligo-Click-M Kit was used to modify certain staple strands
- The following samples were prepared as per the kit instructions:
- 0.1 - 1 mM solution of the oligo-alkyne strand
- 10 mM solution of the Atto550-azide or Atto647N-azide
- Transfer 5 uL of the kit’s activator into the green vial with the kit’s Reactor M catalyst.
- Pipette the appropriate volume of the oligo-alkyne strand solution to the vial.*
- Pipette the appropriate volume of the Atto550-azide or Atto647N-azide solution to the vial.*
- Gently vortex the vial for 10 s.
- Put in thermomixer, incubator, or other appropriate machines to heat at 45 °C for 1-4 hr.
- After heating, transfer liquid phase into a new empty tube.
- Wash the solid catalyst in green vial with 60 uL of 3M NaOAc.
- Transfer liquid phase to tube from step 7.
- Follow your preferred DNA precipitation protocol to precipitate the modified strands.
Cross-Linking to Increase Structural Integrity
Cross-Linkers Increase Thermal Stability of DNA Origami
A crosslinker is able to go in between DNA bases and a covalent reaction occurs to permanently attach. The specific crosslinker is called 8-MOP, which is “activated” by UV radiation
1. Prepare quantities of 8-MOP, Triangle origami in buffer solution, as well as fluorescent DNA strand. Making your batch.
2. Vortex components together and quickly add back on ice
3. Separate chilled solution into PCR tubes in small quantity less than 8ul.
4. Transfer samples to ice bath under UV lamp.
5. Let samples sit in ice for 8 hours ensuring that they stay cool under the UV lamp.
6. After samples have crosslinked run an acrylamide gel to samples without fluorescent DNA strands to expose origami to denaturing conditions.
7. Samples labeled with Flour (from same batch) should be annealed before running agarose gel for imaging.
Anneal samples labeled with flour from 65 degrees until room temperature Run an agarose gel on samples with fluorescent DNA.
Forster Resonance Energy Transfer (FRET)
- Pour a 2% agarose gel following the protocol.
- Prepare 11 mM MgCl2
- Prepare 0.5 TBE with 11 mM MgCl2
- Add sufficient MgCl2 to agarose gel mixture up to 11 mM concentration
- Prepare sufficient agarose gel to make a 2% gel with at least 1cm thickness
- Follow standard gel pouring protocol
Running the Instrument
- Using the TYPHOON FLA 9500 scanner, input the settings.
- These settings are for use with ATTO 647N and ATTO 550 dyes
- FRET Fluorescence
- Use LPFR (long pass filter) and scan with 532nm laser (green)
- DONOR Fluorescence
- Use BPFG (band pass filter green) and scan with 532nm laser
- ACCEPTOR Fluorescence
- Use LPFR (long pass filter) and scan with 635nm laser
- Run scans, save images as .gel format.
The highlighted color of the image during scanning is irrelevant as it is false color. The channels will be saved as different layers and shown as overlapped. Select each individual layer to export. Export images to bmp in ImageQuant with 600 dpi for sufficient resolution.
- Prepare two columns by pipetting 250μl neutravidin agarose resin into each centrifuge column.
- Label one the control, which will not be functionalized with A prime strand, and one experimental, which will be functionalized with A prime strand。
- Twist the bottom off columns and spin at 5000xg.
- Wash three times by loading 250μl 1X TAEMg onto column and spinning again at 5000xg at two minutes per wash. Discard all washings.
- Functionalize experimental column with 40 picomoles of A prime strand in 80μl water. Let it sit for ten minutes, then wash six times with 1X TAEMg at two minutes per wash. Discard all washings.
- Apply 1000 picomoles of T15 strand in 80μl water to both control and experimental columns. Let it sit for ten minutes, wash six times with 1X TAEMg. Discard all washings.
- Prepare two sticky claw/blunt triangle mixtures (abbreviated SCBT) by combining 0.5 picomoles sticky claw and 1.5 picomoles blunt triangle in two reaction tubes.
- Apply one mixture to each column, and let it sit for ten minutes.
- Spin down and wash three times with 70 uL 5% formamide 1X TAEMg per wash, two minutes per wash. Keep washings as nonspecific wash. (Note: There should be a volume of 210μl per column.
- Apply 70 ul 1X TAEMg to each column and expose to UV light for fifteen minutes. Spin down for two minutes and apply two additional washes of 70 ul for a total volume of 210μl per column. Keep washings as UV wash.
- In four centrifugal filter units, pipet your washings: control nonspecific wash, control UV wash, experimental nonspecific wash, and experimental UV wash. Spin at 5000xg for ten minutes right-side up.
- Obtain clean outer centrifuge tubes, label each, and place each filter unit upside down in the appropriate tube. Spin for ten minutes at 5000xg.
- Run an agarose gel with a 1kb marker lane, a lane for a fresh and untouched SCBT mixture, and four lanes for resin wash control, resin wash experimental, resin cleave control, and resin cleave experimental.
Aptamer(or T21)/VLP Binding
Two protocols were experimented with:
- The same binding protocol as with origami/VLP. (see above in VLP/Origami Binding Anneal Step 6.)
- Let aptamers and VLP sit at room temperature for 1 hr. (adapted from literature9)
An excess of aptamers/T21 with respect to NoV VLP was used for binding, for example, a 1:5 ratio of aptamer/T21:VLP. This ratio was experimented with but it was always an excess of aptamers.
Concentrations of Aptamers/T21 Used:
- AG3: 0.002 uM
- T21: 0.002 uM
Buffer Used: General Sensing Buffer, GSB (50 mM NaCl, 20 mM Tris-HCl, 3 mM MgCl2, 5 mM KCl, pH 7.4)
General Protocol for Using the Fluorometer
Taking Emission Spectra
- Turn on fluorometer, allow 20 minutes for warmup.
- Using micropipette, place 15-20 uL of sample into your microcuvette.
- Microcuvette used for our experiements: Quartz Fluormeter Cell from Starna Cells, Inc. Fits 12 uL.
- Cleaning protocol for fluorometer use: Follow instructions on cuvette cleaner provided with cuvette (several drops, rinse copiously)
- Open fluorometer software, according to model. Model used for our experiments: PerkinElmer LS 55 Fluorescence spectrometer
- Parameters for emission spectra of sample
- If excitation wavelength not known, use pre-scan function [200-700 nm excitation range]
- AG3 fluorescent aptamer: Ex Wave - 490 nm, Em Wave – 520 nm
- For parameters used for standard dyes, see literature.6
- Standard slit width used: 10 nm excitation slit; 10 nm emission slit
- Kept standard scan speed at 300 nm/min for all experiments
- Note: This software has an upper limit for the detection of intensity, thus, lower concentrations were used for both sensitivity and efficiency
- Start emission spectra of sample
- For a full wavelength range, this should take approximately three minutes
- Turn on fluorometer, allow 20 minutes for warmup.
- Using micropipette, place 15-20 uL of sample into your microcuvette
- For specifications on microcuvette, see above [How to emission spectra]
- When taking polarization measurements for binding experiments, follow specific binding protocols prior to fluorometer set-up
- Parameters for polarization of sample
- For AG3 aptamer: Ex Wave – 490 nm, Em Wave – 520 nm, Ex Slit – 10 nm, Em Slit – 10 nm, Integration time – 5 s
- Start polarization; about 20 measurements taken, repeat several times.
- Note: these conditions do not match those of the literature8 used as a basis for this data collection; we have found that these conditions produce more reproducible data.
Atomic Force Microscopy
- Items required:
- Freshly filtered (through a 0.2 um filter) water
- AFM samples
- Small round adhesive stickers
- Sheet of mica
- Hole Puncher
- 1-10 uL pipettors and 100-1000 uL pipettor with their tips
- Can of dust remover spray
- AFM tips/ tip holder
- Multimode Nanoscope with software
- Laser head
- Put the Multimode Nanoscope on the microscope stage
- Fit and connect the appropriate scanner to the Multimode Nanoscope
- Properly place the laser head onto the scanner and connect it to the Multimode Nanoscope
- Secure the laser head by attaching the springs from the Nanoscope onto the sides of the laser head
- Attach cable from external AFM controller to the AFM
- Turn on computer
- Turn on the external AFM controller
- Choose appropriate options according to AFM software that is being used
- After mounting the tip onto the tip holder, place into the laser head
- Using the knob in the back (cantilever holder clamp), lower the clamps onto the tip holder until a slight resistance is felt
- Use camera attached and knobs on the stage to focus on the cantilever
- Focus the laser on the cantilever using the knobs on the laser head
- Maximize the signal on the photo diode while switch is on AFM/LFM mode
- Adjust the mirror on the back of the laser head
- Move the laser to reposition it on the cantilever
- Using the photo sensor knobs, get the RMS and VERT values as close to zero on the Multimode Nanoscope
- Take a puck and apply an adhesive sticker to provide a sticky surface for the mica
- Hole punch a piece of mica and place it on the [sticky] puck
- Press the mica onto the puck by the edges and hold for 1 minute
- Nothing must touch the center of the mica
- Now that the mica is securely placed on the puck, hold the puck with tweezers
- Take a small piece of tape, place on the mica and peel off
- Lift up piece of tape to the light to make sure that there are no cracks on the mica
- If there are cracks on the tape, use a new piece of tape on the mica
- Continue to place and peel tape on the pica until there are no noticeable cracks on the tape
- Pipet origami solution (about 3-5 uL) onto the center of the mica
- Allow solution time to set (30 seconds to 1 minute)
- Take 1 mL of filtered deionized water and gently squirt small portions onto the mica surface
- Take the edge of a Kimwipe and wick the edges of the mica to soak up excess water
- To dry the mica surface, grab a dust remover spray and gently blow on the mica for 1-2 minutes
- Raise the tip up by using the right AFM switch and flipping it forwards for about 10 seconds
- Remove the laser head carefully by removing springs
- Carefully put mica onto stage
- Put laser head back on and secure it in place with springs
- Lower the tip back down by using the same AFM switch as in the first step and flipping it backwards for about 10 seconds
- Make sure that all settings on the software correlate with the type of mode that is planning to be used
- Follow instructions of the Multimode Nanoscope that is being used to engage the tip to begin scanning the sample
- 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
- If needed, lower the amplitude setpoint voltage to make the cantilever scan with a greater force so that lift off does not happen
- Adjust scan angle and scan speed (usually 2Hz to start with) to maximise image resolution
Gel Imaging for Agarose and Acrylamide
- After running the gel, carefully remove the plugs from the power supply.
- Prepare a staining solution using ethidium bromide (EtBr). Do this by adding 10 – 20 uL of 10mg/ml ethidium bromide into a glass tray. Then fill the tray up to 1 inch with D.D. water. Gently lift the gel from the plate and move it into the tray for staining. Cover the tray with aluminum foil, and let it stain for 10 – 15 minutes.
- To get more sensitivity, Sybr Gold can be used as a stain. The stain can be prepared with 20 uL of Sybr Gold 1000x concentrate and repeating the steps above for staining.
- To image the gel, carefully lift and place the gel on the pre-cleaned UV lightbox; turn the lightbox on
- Open the software for Carestream and capture the image maximizing the intensity without saturating the detector (use apeture/and exposure controls).