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

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CaDNAno
DD
NUPACK
Oligo Analyzer
Sequence Massager
MFold


==DD==
Thermocycler
http://107.22.192.99:3000/
1-pot annealing
2-pot annealing


#Login
Gel Electrophoresis
#Click Tools -> DD
Gel Imaging
#Add sequences, and fix base positions - capital letters remain constant, lower case letters mutate (double click on sequence to edit)
Gel Purification
#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==
Centrifuge
http://nupack.org/partition/new


Settings:
AFM
*Compute melt
*Concentration: 1 μM
 
==Oligo Analyzer==
http://www.idtdna.com/analyzer/applications/oligoanalyzer/
 
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==
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
 
Folding temperature: 25°C
 
=Orders=
 
==Experiment 1 Order==
===Column Order===
'''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===
'''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=
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
#Stock: 1M MgCl2
#Dilute to 160 mM MgCl2 -1.6 mL 1 M MgCl2 and 8.4 mL H2O
#End: 10 mL of 160mM Mg Buffer
 
Setting Up SST Reaction
#Add 20 µL of D-DNA strand solution to 20 µL of our diluted buffer
#Add 160 µL of H2O
#End: 200 µL solution of 100 nM of each D-DNA strand and 16 mM of MgCl2 buffer
 
PCR 
#Turn dial to big tube
#Files -> New
#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=
20μL solution
10μL 200 nM DNA solution
2μL 125 mM MgCl2
8μL ddH2O
 
For 200 nM DNA solution
1000μL solution
2μL×362 strands of 100 μM strands
276μL H2O
 
=Folding for 10H x 10T D-DNA SST Canvases (for purification and AFM imaging in Experiment 1)=
*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=
2 μL Ribbon A (100 μM)
2 μL Ribbon B (100 μM)
20 μL MgCl2 (125 mM)
176 μL ddH20
 
200 μL final solution: 1 μM DNA, 12.5 μM MgCL2.
 
==Annealing process==
20 μL solution above in PCR machine.
 
Annealing program:
 
[[Experiment 2a#Experiment 2a-1: Annealing at 90 degrees C|Experiment 2a-1:]] Brian Wei's 17H program
 
 
=Suspending L-DNA=
*Calculate volume of 1xTE (Tris EDTA) buffer needed to make 100uM solution based on nmoles of solid provided
*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, 100μm 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=
*Place gel back on original gel electrophoresis tray
*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=
 
*Centrifuge tube that holds 800 uL
**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=
*Take gradient from cold room, being careful not to tilt to disturb
**Larger tubes get disturbed more easily
*Take 15uL of sample and mix with 40% glycerol (to form 20uL of 10% glycerol layer)
*Pipette to top without forming drops (which will disturb gradient)
*Take Beckman centrifuge rack and insert gradient
**Remember to follow the numbering and to add the small tube adapter
*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=
#Open Nanodrop 2000/2000c. Click Nucleic Acid
#Start a new workbook or open an existing workbook
#Clean nanodrop well (top and bottom) with kimwipes and ethanol
#Run routine verifcation (make sure arm is down).
#Open arm. Blank with 1.2 to 1.5 μL 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)
 
Repeat steps below for all samples
#Clean well with kimwipes
#Input Sample ID
#Load 1.2 - 1.5 μL of sample and click measure twice for each sample
 
Note:
*Want a 260/280 ratio of around 1.8 for DNA. Deviations from this ratio indicate possible contamination
*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
 
=AFM Imaging=
#Run NanoScope 8.1 software
#Select "Tapping Mode"
#Intro screen: choose "tapping mode in fluid" and save file in BioMod workspace
#Change interface to "Expanded Mode"
#Use tape to remove top layer off mica disk
#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>
#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>
#Put mica centered on microscope
#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==
*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.
*If sum value is too low, clean cell with ethanol then ddH2O and then dry with pressurized air.
*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=
*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
 
=Adding Streptavidin=
*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:'''
*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=
==Loading/Running==
*Mix 5 uL of sample with 1 uL 6x xylene cyanol (loading dye)
*Mark the bottom of each well with marker
*Lock pre-made gel into gel tray
*Remove well protectors
*Fill inner container above white rim with 0.5x TBE. Fill outer container about halfway with 0.5x TBE
*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==
*Mix 10 uL SYBR Safe in 0.5x TBE buffer on shaker
*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

Revision as of 17:36, 26 October 2012

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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)

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)

CaDNAno DD NUPACK Oligo Analyzer Sequence Massager MFold

Thermocycler 1-pot annealing 2-pot annealing

Gel Electrophoresis Gel Imaging Gel Purification

Centrifuge

AFM

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