Biomod/2012/Harvard/BioDesign/protocols: Difference between revisions

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{{Template:Biomod/2012/Harvard/BioDesign}}
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<font size="5">Protocols</font>


----
<br>


Clarity: Is the project description well-written and easy to understand? Does it include the background and motivation of the project, '''methods''', results, and discussion? '''Are the figures easy to understand?''' (10 points)
=How to Analyze DNA Structures=


Transparency: Are all of the raw experimental data and source files easily accessible? '''Would it be straightforward to attempt to reproduce the team's results?''' (5 points)
==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 6&mu;L 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 5&mu;L of sample to 1&mu;L of loading dye and add to each well
*Add 1&mu;L 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


=Small Canvas SST=
==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, 100&mu;m 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


=DNA Origami=
==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


==Designing and Ordering Strands==
==Preparing Mica for AFM==
# Download [http://cadnano.org Cadnano2] and follow installation directions
*Put 5-minute epoxy into small weighboat
# Watch the [http://www.youtube.com/watch?v=cwj-4Wj6PMc tutorials] on YouTube
*Use large pipette to mix epoxy together
# Download and manipulate [[Image: DNA_Origami.json]] and generate DNA sequences. Edit and add sequences as needed.
*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 x&mu;L (see [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#Large_Canvas_AFM_Specific_Notes Large Canvas AFM Protocol] and [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#Small_Canvas_AFM_Specific_Notes Small Canvas AFM Protocol]) of sample
*Add y&mu;L (see [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#Large_Canvas_AFM_Specific_Notes Large Canvas AFM Protocol] and [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#Small_Canvas_AFM_Specific_Notes Small Canvas AFM Protocol]) of 1xTE Buffer
*Add z&mu;L (see [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#Large_Canvas_AFM_Specific_Notes Large Canvas AFM Protocol] and [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#Small_Canvas_AFM_Specific_Notes Small Canvas AFM Protocol]) of nickel solution if necessary


=Large Canvas SST=
*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


=How to Make L-DNA Layer=




==Dynamic Workbench: DD==


==DD==
*Login
http://107.22.192.99:3000/
*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


#Login
==NuPack==
#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
http://nupack.org/partition/new
*Set concentration: 1 &#956;M
 
Settings:
*Compute melt
*Concentration: 1 &#956;M


==Oligo Analyzer==
==Oligo Analyzer==
http://www.idtdna.com/analyzer/applications/oligoanalyzer/


Settings:
Settings:
Line 50: Line 112:
*Mg++ Conc: 10mM
*Mg++ Conc: 10mM
*dNTPs Conc: 0 mM
*dNTPs Conc: 0 mM
 
*Use Analyze and Self-Dimer to optimize
Use Analyze and Self-Dimer to optimize


==Sequence Massager==
==Sequence Massager==
http://www.attotron.com/cybertory/analysis/seqMassager.htm
Click Reverse and Complement
==Mfold==
http://mfold.rna.albany.edu/?q=mfold/DNA-Folding-Form
Settings:
Na+: 0 mM


Mg++: 10 mM
*Click Reverse and Complement as needed


Folding temperature: 25&#176;C
==M-Fold==


=Orders=
*Na+: 0 mM
*Mg++: 10 mM
*Folding temperature: 25&#176;C


==Experiment 1 Order==
==Annealing onto Template==
===Column Order===
'''40°C Down Anneal'''
'''10 nmol'''
Loading Scheme: Columns (A1, B1, C1...)
Amount Per Well Type: Full Yield
Purification: Standard Desalting
Amount Per Well: 0 nm
Synthesis Scale: 10 nmole DNA
Plate Oligo Concentration: 100 µM
Shipping Option: Wet
Final Volume: 0 µL Ship
Remainder: No
CE Service: No
Documentation: Email
Plate Type: V-Bottom
Dilutant: RNase-Free Water


===Row Order===
Temperature Control Mode: Calculated
'''10 nmol'''
Loading Scheme: Rows (A1, A2, A3...)
Amount Per Well Type: Full Yield
Purification: Standard Desalting
Amount Per Well: 0 nm
Synthesis Scale: 10 nmole DNA
Plate Oligo Concentration: 100 µM
Shipping Option: Wet
Final Volume: 0 µL Ship
Remainder: No
CE Service: No
Documentation: Email
Plate Type: V-Bottom
Dilutant: RNase-Free Water


=Folding for 6 x 11 D-DNA SST Rectangle=
Lid Control Mode: Tracking at 5°C above
Making 1µM D-DNA Strand Solution
#Stock: 66 unique 100 µM strands
#Add 5 µL of each D-DNA strand into a PCR tube
##Nothing left in Well F3
##Nothing left in Well C4
#Add 170 µL of DD H2O
#End: 500 µL of 66 unique 1 µM D-DNA strands


Making 160mM Mg Buffer
*Incubate at 40.0°C for 20 minutes
#Stock: 1M MgCl2
*Decrease by 1.0°C every cycle
#Dilute to 160 mM MgCl2 -1.6 mL 1 M MgCl2 and 8.4 mL H2O
*Cycle to step 1 for 15 more times
#End: 10 mL of 160mM Mg Buffer
*Incubate at 4.0°C forever


Setting Up SST Reaction
=How to Make Small SST Canvas=
#Add 20 µL of D-DNA strand solution to 20 µL of our diluted buffer
==Strand Mixture==
#Add 160 µL of H2O
*Mix 1 &mu;M solution of D-DNA SSTs from even rows (30 SSTs)
#End: 200 µL solution of 100 nM of each D-DNA strand and 16 mM of MgCl2 buffer
*Mix 1 &mu;M solution of D-DNA SSTs from protector strands (12 strands)
*Mix 1 &mu;M solution of D-DNA SSTs from rows 3, 5, 7, and 9 for each handle variation


PCR 
==Adding the L-DNA==
#Turn dial to big tube
*Add L-DNA strand A to canvas solution in a 1:7 ratio
#Files -> New
*Add L-DNA strand B to purified solution in a 1:7 ratio
#Lid temperature set at 105 degrees
#Wait
#Select 44 degrees
#Set at 30 minutes
#Hold at 20 degrees
#Exit
#Save as “Hold44”
#Run
#Approximately 12:20PM-12:50PM
#It will read “Hold at 20 degrees” when done.


=Folding for 24H x 29T D-DNA SST Rectangle=
==Annealing Template==
20&mu;L solution
'''PCR Machine: 17 Hour Anneal'''
10&mu;L 200 nM DNA solution
2&mu;L 125 mM MgCl2
8&mu;L ddH2O


For 200 nM DNA solution
Temperature Control Mode: Calculated
1000&mu;L solution
2&mu;L&times;362 strands of 100 &mu;M strands
276&mu;L H2O


=Folding for 10H x 10T D-DNA SST Canvases (for purification and AFM imaging in Experiment 1)=
Lid Control Mode: Tracking at 5°C above
*2.5uL per pre-combined tube 1 of D-DNA SSTs
*2.5uL per pre-combined tube 2 of D-DNA SSTs
*2.5uL per pre-combined tube 3 of D-DNA SSTs
*2.5uL for 10x folding buffer with MgCl2
*15uL ddH2O


=Folding for D-DNA SST ribbons=
*Incubate at 90.0°C for 10 minutes
2 &mu;L Ribbon A (100 &mu;M)
*Decrease by 1.0°C every cycle
2 &mu;L Ribbon B (100 &mu;M)
*Cycle to step 1 for 29 more times
20 &mu;L MgCl2 (125 mM)
*Incubate at 60.0°C for 20 minutes
176 &mu;L ddH20
*Decrease by 1.0°C every cycle
*Cycle to step 3 for 35 more times
*Incubate at 4.0°C forever


200 &mu;L final solution: 1 &mu;M DNA, 12.5 &mu;M MgCL2.
==Small Canvas AFM Specific Notes==
*Use 5&mu;L sample and 30&mu;L 1x TE Buffer with 20&mu;L nickel and follow [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#General_AFM_Protocol General AFM Protocol]


==Annealing process==
=How to Make DNA Origami=
20 &mu;L solution above in PCR machine.


Annealing program:
==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


[[Experiment 2a#Experiment 2a-1: Annealing at 90 degrees C|Experiment 2a-1:]] Brian Wei's 17H program
==Adding the L-DNA==
 
*Add 2.5 uL of 10uM first ribbon of L-DNA
 
*Anneal Ribbon A - See [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#Annealing_onto_Template Annealing onto Template]
=Suspending L-DNA=
*Purify - [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#Gel_Purification Gel Purification]
*Calculate volume of 1xTE (Tris EDTA) buffer needed to make 100uM solution based on nmoles of solid provided
*Anneal Ribbon B - [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#Annealing_onto_Template Annealing onto Template]
*Add 1xTE buffer to solid L-DNA
*Vortex and centrifuge
 
==Folding for L-DNA ribbons==
See Folding for D-DNA ribbons
 
=Gel Preparation=
*Clean gel plate with ddH2O
*120mL 0.5x TBE buffer
*2.4g agarose (powder)
*~10mL extra H2O for correction of volume during evaporation
*(heat up in microwave for 2min in full power until agarose melts and solution boils)
*(wait ~5min for the solution to cool down a bit, or cool by swirling around in bucket of water several times until beaker is cool enough to hold in hands indefinitely)
*1mL 1.2M MgCl2
*6uL 10,000x SybrSafe stock solution (NOTE: SybrSafe Gold is extremely toxic. Be very careful when handling this solution)
*(pour into sealed gel box, and put in combs, let cool for >15 min)
*Use comb to scoop out bubbles as needed
*Note on combs: narrow-thick wells for 5uL sample + 1uL dye loading.
*Turn gel sideways after it is formed
*Put .5 TBE + 10mM Magnesium buffer until at fill line
*Add 5 microliters of DNA solution and 1 microliter of loading dye (spin down as needed)
 
=Gel Electrophoresis=
*Add top onto tray so that red terminal is pointed towards you, black terminal is pointed away
*90V
*400 mA (set as maximum limit)
*100 Watts (set as maximum limit)
*Timer set in unit of hours, 1 hour 30 min.
 
=Gel Imaging=
*Take gel out and place on grid
*Select the Typhoon FLA 9000 icon
*Click on the Fluorescence setting
*Type in a file name
*Save to Data -> Biomod
*Method: SYBR Safe Mode, 400V PMT, 100&mu;m resolution, preset values for correction
*Set scan area
*Click on Pre-scan
*On the bottom border of the scanning dialog make sure it says "Gel + TIFF", if not change at Options > Preferences.
*Adjust brightness, then hit return
*Adjust scan areas, then hit scan
*Remove gel
*Spray ethanol (isopropyl if ethanol not available) on glass area in contact with gel. Wipe down with Kimwipes.
*Spray water on glass area in contact with gel. Wipe down with Kimwipes.


=Gel Purification=
==Annealing==
*Place gel back on original gel electrophoresis tray
'''PCR Machine: 72 Hour Anneal'''
*Transport gel to darkroom
*Place gel on viewing surface
*Place orange screen over gel
*Wear orange glasses
*Turn on UV lights
*Cut vertically down the sides of the desired gel band
*Cut horizontally above and below the desired gel band
*Use blade and leverage to carefully dig out gel band
*Place gel band inside SEPARATE tubes (NOT Freeze-and-Squeeze tubes)
*Dispose of excess gel in waste container labeled specifically for gel waste (next to the entrance of the Yin lab)
*Use pestle to crush sample inside the tubes using the POINTY end.
*Place tubes inside centrifuge and DON'T FORGET metal lid
*Spin down gel at 4,000 rcf for approximately 20 seconds.
*Use tube cutter to cut off tip of tube with gel contained within.
*Invert tube tip into the "Freeze-and-Squeeze" tube
*Place tube (balance using another "Freeze-and-Squeeze" tube) inside centrifuge and DON'T FORGET metal lid
*Spin down gel at 400-1000 rcf for 3-4 minutes
*Remove filter from "Freeze-and-Squeeze" tube
*Store purified DNA sample in fridge if temporary. Store in freezer if to be stored for 1 month or longer.


=Making a Glycerol Gradient=
Temperature Control Mode: Calculated


*Centrifuge tube that holds 800 uL
Lid Control Mode: Tracking at 10°C above
**Box says 5 x 41 mm, Company: Beckman
*Glycerol solution preparation:
**5mM Tris pH~8.5, 1mM EDTA, 10mM MgCl2, anywhere to 45% to 15% glycerol by volume
**7 of these
*Pipette 80 uL of each percentage of glycerol solution into tube
** From 45% down to 15% in increments of 5 (45% at the bottom, 560 uL total)
**Hold pipette tip in solution until glycerol stops moving up
**When adding the next layer, add it slowly on top of the last layer
*Leave overnight in the cold room


=Running the Gradient=
*Incubate at 80.0°C for 5 minutes
*Take gradient from cold room, being careful not to tilt to disturb
*Decrease by 1.0°C every cycle
**Larger tubes get disturbed more easily
*Cycle to step 1 for 15 more times
*Take 15uL of sample and mix with 40% glycerol (to form 20uL of 10% glycerol layer)
*Incubate at 64.0°C for 1 hour 45 minutes
*Pipette to top without forming drops (which will disturb gradient)
*Decrease by 1.0°C every cycle
*Take Beckman centrifuge rack and insert gradient
*Cycle to step 3 for 40 more times
**Remember to follow the numbering and to add the small tube adapter
*Incubate at 4.0°C forever
*Make sure all tubes are balanced
*Take to ultracentrifuge room in Ingber lab (520)
*Spin for 1-3 hours at 48-50k RPM


=NanoDrop 2000=
==TEM==
#Open Nanodrop 2000/2000c. Click Nucleic Acid
'''2% aqueous uranyl formate stain solution (3 mL H20, 0.06 g uranyl formate):'''
#Start a new workbook or open an existing workbook
**Uranyl formate (EMS, powder, store with parafilm over lid because seems to degrade in air)
#Clean nanodrop well (top and bottom) with kimwipes and ethanol
**Weigh out 0.06 g uranyl formate in 10 mL beaker, using the balance in the hood
#Run routine verifcation (make sure arm is down).
**Put in magnetic stir bar
#Open arm. Blank with 1.2 to 1.5 &mu;L of 1x TE Mg buffer (a bead of sample should form). Close arm and click blank. (If bubble is not deformed, it's a sign of a clean measurement)
**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


Repeat steps below for all samples
'''Glow discharging the grids:'''
#Clean well with kimwipes
**Formvar/Carbon coated grids (SPI # 3440C)
#Input Sample ID
**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
#Load 1.2 - 1.5 &mu;L of sample and click measure twice for each sample
**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".


Note:
'''Putting sample on grid and staining:'''
*Want a 260/280 ratio of around 1.8 for DNA. Deviations from this ratio indicate possible contamination
**Materials: 15 mL falcon tube (wrap in foil), filter (Acrodisk, 0.2 um) that mounts on 5 mL syringe tip, 5 mL syringe (BD)
*Want a 260/230 ratio of around 2.0-2.2. In general this ratio should be greater than the 260/280 ratio. A low ratio may indicate contamination whereas a high ratio may indicate a dirty blank
**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


=AFM Imaging=
=How to Make Large SST Canvas=
#Run NanoScope 8.1 software
==Strand Mixture==
#Select "Tapping Mode"
*Solution A: Mix 5 &mu;L of exterior SSTs and interior SSTs from even rows (340 SSTs)
#Intro screen: choose "tapping mode in fluid" and save file in BioMod workspace
*Solution B: Mix 5 &mu;L of interior SSTs from odd rows (35 SSTs)
#Change interface to "Expanded Mode"
*For 200 nM reaction, combine:
#Use tape to remove top layer off mica disk
**68 &mu:M solution A
#Add 5 uL Mg buffer and make sure not to scratch surface. Solution spreads easily if on smooth surface. <span style="color:#d00">BETTER RESULTS FOUND USING 0 uL</span>
**7 &mu:M solution B
#Add 5 uL DNA solution (for 2-3 nM DNA) <span style="color:#d00">BETTER RESULTS FOUND USING 20 uL - also make sure to wait a bit for the sample to be diluted by buffer before adding nickel!</span>
**10 &mu:M 10x 12.5 mM folding buffer
#Put mica centered on microscope
**15 &mu:M ddH<sub>2</sub>O
#Add 30 uL of 10 mM nickel solution to mica disk <span style="color:#d00">BETTER RESULTS FOUND USING 20 uL</span>
#Use "up" switch on piezo to move mica down, leaving more space for the cantilever
#Put cantilever in fluid cell
#*'''Note:''' cantilver has 4 tips; make sure cantilever is oriented with the end with the desired tip pointing up (higher than the other end).
#*Use spring on back to hold cantilever in place
#Place fluid cell into AFM so fluid inlets are facing outwards
#Screw top onto fluid cell using knob in back. Do NOT tighten excessively.
#Add 30 uL 1X Folding Buffer (Magnesium Buffer) to the buffer inlet in fluid cell
#*Forms drop around cantilever to protect it
#Turn on microscope light (1/3 of the way) and center field of view on desired cantilever tip using two lower knobs at bottom of AFM
#Focus (using larger knob at top) on mica surface (since mica is clear, focus above the metal beneath it)
#Use "down" switch on piezo until cantilever comes into focus
#Change switch to middle position (AFM & LFM) to see laser
#2 step process to move laser to cantilever tip
##use 2 knobs on scanner head to center laser on tip -- move as close to tip as possible while maximizing sum (>5 is good)
##use other 2 knobs next to camera lens to change x and y offsets (of the light diode) as close to zero as possible
##*wait to see if the offsets drift; if they do, then repeat until stable
#If desired, use right click to dock or undock viewing screen.
#On computer, click "Tune" to find resonance frequency, then "Autotune"
#*finds resonance peak of cantilever
#*increase sweep width to ~5 (Hz?) to zoom out to see peak
#Click "Engage"
#*tip steps down slowly until touching sample
#*make sure scan size is set to 1 nm so the microscopic with effectively not scan when finished engaging
#Scanning
#*Increase scan size to 1.0 uM
#*Left graph = amplitude, right graph = error
#*Red line = first pass, blue line = second pass (reversed)
#**lines should be correlated
#*Click "Save capture directory" in upper right and choose directory (BioMod 2012) & filename
#*Click "Save"
#When done scanning:
##Click "Withdraw"
##Press "Up" switch on piezo to retract fluid cell and cantilever
##Unscrew top
##Remove fluid cell
##Dispose of cantilever down the drain
##Spray fluid cell with water and clean with soap/isopropyl. Be sure not to bend clip on the fluid cell!
##Dry fluid cell using Kimwipes if not to be used for a while. If fluid cell needed again right away, use pressurized air to dry.
##Remove mica with tweezers, wipe off and store in special container (because of nickel)


==Troubleshooting AFM==
==Adding L-DNA==
*If green bar turns red and the dialogue reads "Retracted" then change "Amplitude Setpoint" to a lower value until bar goes back to green and the moving black bar hits the middle.
*Add L-DNA strand A to canvas solution in a 1:7 ratio
*If sum value is too low, clean cell with ethanol then ddH2O and then dry with pressurized air.
*Add L-DNA strand B to purified solution in a 1:7 ratio
*Drive amplitude below 100Hz is good. If drive amplitude is something like 1000Hz, you need to reset sweep width while autotuning to 5kHz
*Changing z limit to 1 um increases image resolution
*Weird patterns or fuzzy edges may mean that the structures are not well-fixed to the surface. Therefore, add more nickel solution
*To clean the fluid cell
**Put soap all over and make sure to clean in direction of clip to avoid bending it
**Clean with tap water
**Then use ethanol or isopropanol
**Then use DI water
**Then dry a bit with compressed air (use pipette in air tube to filter particulates
**Remove excess water with kimwipes and make sure that the electrode (metal part) is COMPLETELY dry to avoid a short circuit
*Target amplitude should be 112 mV
*Peak offset tells you how offset the reading is from the tip of the highest peak
*Don't worry about target amplitude value
*Z-limit must go down to get more data points - not really necessary to touch for these samples
*If you get an error message and withdraw, you might want to readjust vertical and horizontal knobs
*Set scan size, x offset, y offset to zero
*Never use the same tip twice because of contamination


=Preparing Mica=
==Annealing Template==
*Put 5-minute epoxy into small weighboat
'''PCR Machine: 17 Hour Anneal'''
*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


=Adding Streptavidin=
Temperature Control Mode: Calculated
*Concentration of streptavidin: 0.2mg/mL
*Amount of streptavidin 10&mu;L
*Amount of buffer: 30&mu;L
*Amount of sample: 5&mu;L
*Make sure to take fluid cell out before adding in the streptavidin directly to the mica surface
*Wait a few minutes for streptavidin to bind to the biotin


'''Observations when trying with Ralf:'''
Lid Control Mode: Tracking at 5°C above
*We first added streptavidin to the origami structures
*We hope to see the rectangle with six dots of streptavidin each
*After, we will add nickel to assess its effect on streptavidin binding. The nickel may result in nonspecific binding
*Withdraw
*Used Q-control to sharpen the peak
*Engage
*We may have used too much streptavidin
*In order to determine the difference between non-binding of streptavidin or perhaps just that the toehold is not sticking up, we need to use statistics to differentiate the scenarios
*Image is pretty good
*Streptavidin is not necessarily useful for determining whether or not the toeholds are sticking up because it will bind regardless
*Streptavidin can be seen as small white dots on the surface of the mica
*We now want to add nickel
*As expected, nickel induces streptavidin to completely coat the mica surface
*Also, Ralf notes that streptavidin is likely simply not useable for our purposes. It may be better to
**Add huge excess of streptavidin to the structure solution
**Purify using glycerol gradient
**Then use nickel
**Good thing is it doesn't have agarose, doesn't allow streptavidin binding to surface, we can image with glycerol on surface
**We may want to make an asymmetric structure to determine the orientation (face up or face down)


=PAGE Gel=
*Incubate at 90.0°C for 10 minutes
==Loading/Running==
*Decrease by 1.0°C every cycle
*Mix 5 uL of sample with 1 uL 6x xylene cyanol (loading dye)
*Cycle to step 1 for 29 more times
*Mark the bottom of each well with marker
*Incubate at 60.0°C for 20 minutes
*Lock pre-made gel into gel tray
*Decrease by 1.0°C every cycle
*Remove well protectors
*Cycle to step 3 for 35 more times
*Fill inner container above white rim with 0.5x TBE. Fill outer container about halfway with 0.5x TBE
*Incubate at 4.0°C forever
*Pipette up and down for each lane in TBU gel to get rid of urea/gunk in wells
*Load 3 uL of sample+dye in each lane
*Run at 120V, 400 mA, 100W, for 30 minutes


==Staining/Imaging==
==Large Canvas AFM Specific Notes==
*Mix 10 uL SYBR Safe in 0.5x TBE buffer on shaker
*Use 5&mu;L sample and 15&mu;L 1x TE Buffer with no nickel and follow [http://openwetware.org/wiki/Biomod/2012/Harvard/BioDesign/protocols#General_AFM_Protocol General AFM Protocol]
*Open gel frame with a chisel
*Place gel onto face of frame with "hole"
*Stain gel in buffer/SYBR Safe mixture, preserving orientation of gel, for 20 minutes
*Drain staining fluid
*Rinse with water
*Make a pool of water of gel tray
*Image gel

Latest revision as of 23:55, 27 October 2012

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Protocols


How to Analyze 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

How to Make 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

How to Make Small SST Canvas

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

How to Make DNA Origami

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

How to Make Large SST Canvas

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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