Biomod/2012/Harvard/BioDesign/protocols

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=Analyzing the DNA Structures=
=Analyzing the DNA Structures=

Revision as of 02:14, 28 October 2012

Protocols


Contents

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

  • 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

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

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

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