Biomod/2012/Harvard/BioDesign/protocols
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Protocols
Analyzing the DNA Structures
Making a Gel
- Create gel mixture
- 120mL 0.5x TBE buffer +
- 2.4g agarose (powder) +
- ~10mL extra H2O for correction of volume during evaporation
- Heat up in microwave for 2 min at full power until agarose melts and solution boils
- Wait ~5 min for the solution to cool down, swirl in water briefly to aid
- Add 1mL of 1.2M MgCl2
- Add 6μL of 10,000x SybrSafe stock solution
- Pour into snug gel tray in gel box, and put in combs, let cool for >15 min
- Use comb to scoop out bubbles in agarose as needed
Running a Gel
- Turn gel sideways in gel box
- Add .5xTBE with 10mM Mg buffer to the gel box until at fill line
- Mix 5μL of sample to 1μL of loading dye and add to each well
- Add 1μL of ladder to lanes on both ends
- Add top onto tray so that red terminal is pointed towards you, black terminal is pointed away
- Set at 90V
- Set max current to 400 mA
- Set max power 100 Watts
- Add ice water to outside of gel tray if necessary
- Run for 1.5 to 3 hours
Imaging a Gel
- Take gel out and place on grid of scanner
- Open Typhoon FLA 9000 icon, Fluorescence setting
- Type in a filename and select destination folder
- Choose: SYBR Safe Mode, 400V PMT, 100μm resolution, preset values for correction
- Set pre-scan area
- Adjust final scan area with pre-scan data
- Scan
- Adjust brightness
- Remove gel
- Clean machine with ethanol and water
Gel Purification
- Transport gel to darkroom
- Place gel on viewing surface
- Wear UV protection glasses and view gel under UV
- Cut out the glowing band and remove the piece
- Lay it on its side and trim the band
- Place gel band in labelled tube
- Dispose of excess gel in waste container labeled specifically for gel waste
- Use pestle to crush sample inside the tubes using the pointed end.
- Spin crushed gel at 400 rcf for 30 seconds to get gel to bottom
- Cut off tip and invert inside freeze and squeeze tube
- Use tube cutter to cut off tip of tube with gel contained within.
- Spin down gel at 400-1000 rcf for 4-5 minutes
Preparing Mica for AFM
- Put 5-minute epoxy into small weighboat
- Use large pipette to mix epoxy together
- Add a small dot of epoxy to disk center
- Place mica on disk
- Evenly distribute epoxy below surface of mica by pushing down on it with a pipette
General AFM Protocol
- Run NanoScope 8.1 software
- Select "Tapping Mode" in Fluid” and load settings
- Use tape to remove top layer of mica disk
- Add xμL (see Large Canvas AFM Protocol and Small Canvas AFM Protocol) of sample
- Add yμL (see Large Canvas AFM Protocol and Small Canvas AFM Protocol) of 1xTE Buffer
- Add zμL (see Large Canvas AFM Protocol and Small Canvas AFM Protocol) of nickel solution if necessary
- Place mica in AFM
- Place tip on cleaned fluid cell and secure with spring clip
- Secure fluid cell
- Move the mica up so that the tip is close to the surface
- Algin laser with the tip, adjust to increase sum through the two laser knobs and mirror
- Set vertical and horizontal offset closer to zero
- Auto-tune, can adjust Q
- Check 5k sweep frequency for clean peaks
- Engage
- Set scan size to 1nm, check amplitude setpoint, set offsets to 0, integral gain to 3 and 6
- Change scan size to desired image size
- Select capture directory and capture
- Withdraw when done
- Remove and clean fluid cell
- Remove and clean mica
About the L-DNA Layer
Dynamic Workbench: DD
- Login
- Click Tools -> DD
- Add sequences, and fix base positions - capital letters remain constant, lower case letters mutate (double click on sequence to edit)
- Select desired nucleotides to include in mutations (double click on composition and choose from scroll down menu)
- Hit mutate - the lower the score, the better
NuPack
- Select compute melt
- Set concentration: 1 μM
Oligo Analyzer
Settings:
- Target type: DNA
- Oligo Conc: 1 μM
- Na+ Conc: 0mM
- Mg++ Conc: 10mM
- dNTPs Conc: 0 mM
- Use Analyze and Self-Dimer to optimize
Sequence Massager
- Click Reverse and Complement as needed
M-Fold
- Na+: 0 mM
- Mg++: 10 mM
- Folding temperature: 25°C
Annealing onto Template
40°C Down Anneal
Temperature Control Mode: Calculated
Lid Control Mode: Tracking at 5°C above
- Incubate at 40.0°C for 20 minutes
- Decrease by 1.0°C every cycle
- Cycle to step 1 for 15 more times
- Incubate at 4.0°C forever
Small Canvas SST Specifics
Strand Mixture
- Mix 1 μM solution of D-DNA SSTs from even rows (30 SSTs)
- Mix 1 μM solution of D-DNA SSTs from protector strands (12 strands)
- Mix 1 μM solution of D-DNA SSTs from rows 3, 5, 7, and 9 for each handle variation
Adding the L-DNA
- Add L-DNA strand A to canvas solution in a 1:7 ratio
- Add L-DNA strand B to purified solution in a 1:7 ratio
Annealing Template
PCR Machine: 17 Hour Anneal
Temperature Control Mode: Calculated
Lid Control Mode: Tracking at 5°C above
- Incubate at 90.0°C for 10 minutes
- Decrease by 1.0°C every cycle
- Cycle to step 1 for 29 more times
- Incubate at 60.0°C for 20 minutes
- Decrease by 1.0°C every cycle
- Cycle to step 3 for 35 more times
- Incubate at 4.0°C forever
Small Canvas AFM Specific Notes
- Use 5μL sample and 30μL 1x TE Buffer with 20μL nickel and follow General AFM Protocol
DNA Origami Specifics
Strand Mixture (50 uL)
- In a PCR tube, add 20 uL of 200 nM staples
- Add 12.5 uL of 200 nM p8064 scaffold
- Add 5 uL of 110 mM Mg++
- Add 7.5 uL ddH2O
Adding the L-DNA
- Add 2.5 uL of 10uM first ribbon of L-DNA
- Anneal Ribbon A - See Annealing onto Template
- Purify - Gel Purification
- Anneal Ribbon B - Annealing onto Template
Annealing
PCR Machine: 72 Hour Anneal
Temperature Control Mode: Calculated
Lid Control Mode: Tracking at 10°C above
- Incubate at 80.0°C for 5 minutes
- Decrease by 1.0°C every cycle
- Cycle to step 1 for 15 more times
- Incubate at 64.0°C for 1 hour 45 minutes
- Decrease by 1.0°C every cycle
- Cycle to step 3 for 40 more times
- Incubate at 4.0°C forever
TEM
2% aqueous uranyl formate stain solution (3 mL H20, 0.06 g uranyl formate):
- Uranyl formate (EMS, powder, store with parafilm over lid because seems to degrade in air)
- Weigh out 0.06 g uranyl formate in 10 mL beaker, using the balance in the hood
- Put in magnetic stir bar
- Put on magnetic stirrer under large inverted foil-topped beaker (don't stir it yet)
- Using a pipette, measure out 3 g H20 + 40 uL H20 = 3 mL H20 + 40 uL H20 (nuclease free, Ambion) in another 10 mL beaker
- Boil this water on a hot plate (NOT the same stirrer used for the uranyl formate) and shut off heat right when it boils
- Can carry over to the formate solution on the stirrer with gloves even though the water beaker is hot
- Pour hot water into formate beaker and turn on magnetic stirring; cover with large inverted foil-topped beaker
Glow discharging the grids:
- Formvar/Carbon coated grids (SPI # 3440C)
- Grab glass slide (e.g., VWR micro slides), gently wrap a piece of parafilm (e.g., 1.5 squares long, 1 sq wide) around a section of the glass slide
- Using tiny tweezers, grab grids from the edge and transfer to the parafilm wrapping on the slide
- Tweezers are Dumont N4AC from EMS. Buy your own tweezers; they break easily. Have one tweezer per grid if doing multiple grid preps in parallel.
- Put in EMS glow discharger. Settings: 25 mA, 45 s, 0.1 mBar, negative HT polarity. Press "start".
Putting sample on grid and staining:
- Materials: 15 mL falcon tube (wrap in foil), filter (Acrodisk, 0.2 um) that mounts on 5 mL syringe tip, 5 mL syringe (BD)
- Draw stain solution into the syringe, screw on filter, discharge through filter into fresh 15 mL tube
- Add 1 mL of uranyl formate solution into a fresh 2 mL eppendorf tube using a pipette
- Add 5 uL of 5N NaOH (JT Baker - this is DANGEROUS - DO NOT GET INTO EYES) into the 1 mL of stain solution in the eppendorf
- Vortex; the solution should get slightly darker
- Use a grid mat in a petri dish
- Line up tweezers, 1 tweezer per grid
- Grab grid edge and let rest on tweezers, suspended over air
- Pipette 3.5 uL of sample onto the grid for 4 min
- Take a piece of whatman paper, bend it in the middle
- Wick off sample by bringing whatman paper in contact with the grid edge from the side
- Immediately add 3.5 uL of stain solution onto the grid and let sit for 1 min
- Wick off the stain as before
- Leave for a minute or two to dry
- Transfer grid to mat at an angle: As grid nears mat surface, open tweezers slightly--the grid will still stick to the tweezers, and drag up to a ridge on the mat, at which point the grid will detach from the tweezers and rest up against the ridge on the mat
- It is crucial not to bend the grid or exert any forces on it
- Put stuff that contacted uranyl formate in the radioactive waste bin
Large Canvas SST Specifics
Strand Mixture
- Solution A: Mix 5 μL of exterior SSTs and interior SSTs from even rows (340 SSTs)
- Solution B: Mix 5 μL of interior SSTs from odd rows (35 SSTs)
- For 200 nM reaction, combine:
- 68 &mu:M solution A
- 7 &mu:M solution B
- 10 &mu:M 10x 12.5 mM folding buffer
- 15 &mu:M ddH2O
Adding L-DNA
- Add L-DNA strand A to canvas solution in a 1:7 ratio
- Add L-DNA strand B to purified solution in a 1:7 ratio
Annealing Template
PCR Machine: 17 Hour Anneal
Temperature Control Mode: Calculated
Lid Control Mode: Tracking at 5°C above
- Incubate at 90.0°C for 10 minutes
- Decrease by 1.0°C every cycle
- Cycle to step 1 for 29 more times
- Incubate at 60.0°C for 20 minutes
- Decrease by 1.0°C every cycle
- Cycle to step 3 for 35 more times
- Incubate at 4.0°C forever
Large Canvas AFM Specific Notes
- Use 5μL sample and 15μL 1x TE Buffer with no nickel and follow General AFM Protocol