Biomod/2012/Harvard/BioDesign/protocols

From OpenWetWare
Revision as of 19:09, 27 October 2012 by Wesley Chen (talk | contribs) (→‎TEM)
Jump to navigationJump to search
The printable version is no longer supported and may have rendering errors. Please update your browser bookmarks and please use the default browser print function instead.

<html>

<head>

<link href='http://fonts.googleapis.com/css?family=Open+Sans' rel='stylesheet' type='text/css'>

</head>



<style>

/*BUT ACTUALLY*/ body { 

 font-family: 'Open Sans', sans-serif;
 overflow-y: scroll;

}

.container { 

 background-color: #ffffff;
 margin-top:0px

} .OWWNBcpCurrentDateFilled { display: none; }

h5 { 

 font-family: 'Open Sans', sans-serif;
 font-size: 11px;
 font-style: normal;
 text-align: center;
 margin:0px;
 padding:0px;

}

  1. column-content

{ 

 width: 0px; 
 float: left; 
 margin: 0 0 0 0;
 padding: 0;

} .firstHeading { 

 display:none;
 width:0px;

}

  1. column-one

{ 

 display:none; 
 width:0px;
 background-color: #ffffff;

}

  1. globalWrapper

{ 

 width: 900px;
 background-color: #ffffff;
 margin-left: auto;
 margin-right: auto

}

  1. content

{ 

 margin: 0 0 0 0;
 align: center;
 padding: 12px 12px 12px 12px;
 width: 876px;
 background-color: #ffffff; border: 0;

}

  1. bodyContent

{ 

 width: 850px;
 align: center;
 background-color: #fffffff;

}

  1. column-content

{ 

 width: 900px;
 background-color: #ffffff;

}

  1. footer

{ 

 position: center;
 width: 900px;

} @media screen { 

   body { background: #000000 0 0 no-repeat;  /* changed default background */ }

}

  1. menu

{ 

 position: fixed;
 float: left;
 width: 180px;
 padding: 10px;
 background-color: #FFFFFF;

}

  1. pagecontent

{ 

 float: right;
 width: 620px;
 margin-left: 300px;
 min-height: 400px

}

  1. toc { display: none; }

/*Expanding list*/ ul { list-style: none; }

  1. exp li ul { display: none; }
  2. exp li:hover ul { display: block; }
  3. exp li a:active ul { display: block; }

a:link {color:#FF6060;} a:visited {color:#FF6060;} /* visited link */ a:hover {color:#B24343;} /* mouse over link */ a:active {color:#B24343; } /* selected link */

</style> </html>

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

  • 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

Retrieved from "https://openwetware.org/mediawiki/index.php?title=Biomod/2012/Harvard/BioDesign/protocols&oldid=647556"