- 1X TBE: Add 50 mL of 10X TBE in a 500 mL volumetric flask. Add D.D. water until the final volume is 500 mL.
- 0.5X TBE: Add 25 mL of 10X TBE in a 500 mL volumetric flask Add D.D. water until the final volume is 500 mL.
- 1X TAEMg: Add 50 mL of 10X TAEMg in a 500 mL volumetric flask. Add D.D. water until the final volume is 500 mL.
- 10X TAE: Dissolve 28.6 g of magnesium acetate with 1 L of a 10X TAE mixed together on a stir plate for 10 – 20 minutes.
- The gel cassette is assembled with two glass plates and two clamps.
- Be sure that the gel cassette is firmly held into place on the gel-holding apparatus.
- Prepare the gel with the 20ml of mixture of 20% gel stock and 0% gel stock.
- Prepare the ammonium persulfate (APS - 10% w/v water) and TEMED solution. For APS, the estimated volume should be 150 – 180 uL. For TEMED, the estimated #volume should be 15 – 18 uL.
- Add the gel stock, APS, and TEMED into a 50 mL tube.
- Pour the gel using the large plastic pipet and insert the comb between the plates. Be sure to carry this out quickly because the gel mixture polymerizes quite quickly. Pouring is often most efficient when the pipet is kept at about 30 degrees below vertical, perpendicular to the plane of the plates.
- Let the entire assembly sit for one hour to ensure complete polymerization.
- Place 1.2 g of agarose in a clean 600 mL beaker on a scale.
- Prepare the 0.5X TBE buffer as described.
- Add 30 mL of D.D. water.
- Microwave for 2 minutes.
- If the solution is not clear, swirl and microwave for another 15 seconds.
- Cool the beaker in an ice water bath. Keep swirling the beaker to prevent polymerization. Add the thermometer and keep checking the temperature.
- When the solution reaches 60°C, remove it from the ice water bath.
- Add 1 mL of 1.2 M magnesium chloride.
- Pour the contents of the beaker into the gel cassette and insert the comb.
- Use a glass Pasteur pipet to push any bubbles to the corners of the gel.
- Leave the cassette covered and let the assembly sit for thirty minutes to complete ensure complete polymerization.
- Set to nucleic acid measurement mode.
- Clean the metal ball on the pedestal with a kim wipe.
- Pipet 1.5 uL of D.D. water onto the small metal ball, and blank.
- Clean the metal ball on the pedestal with a kim wipe.
- Pipet 1.5 uL of the sample onto the small metal ball, and measure.
- Read the absorbance at 260 nm.
- This is done in triplicate and the average is obtained.
Capsid Buffer Exchange
- From the capsid stock solution, pipette an amount sufficient for the entire experiment into a 50 kDa centrifugal filter tube. (Note: #Make sure to return the capsid stock solution to the fridge.)
- Fill the centrifugal filter tube up to 500 uL with 1X TAE-Mg and close the lid tightly.
- Place the centrifugal filter tube into the centrifuge and spin at an RCF of 5 for 6 minutes.
- Dispose the waste solution at the bottom of the centrifugal filter tube.
- Repeat steps 3–5 twice more.
- Spin for additional time as needed to reduce the final volume.
Capsid/Origami Binding Anneal
- Exchange capsid buffer to 1X TAEMg.
- Pipet origami and capsid into PCR tube in the desire ratio.
- Add the nonstandard components as needed for a particular experiment.
- Add buffer to bring anneal to the desired concentration for a particular experiment.
- Use a microcentrifuge to collect he sample at the bottom of the PCR tube.
- Place the PCR tubes for the samples in the PCR machine and run the anneal protocol. (Follow the Annealing protocols for the PCR machine.)
Origami Sample Preparation
There are three phases to the preparation of origami samples, which can be performed at different times if the solutions are stored frozen in between steps. These phases are (1) staple stock solution preparation, (2) preparation of the complete origami solution, and (3) the anneal.
Preparation of Staple Stocks
- Take the origami staple strand plates from the freezer and allow it to defrost to room temperature. This generally takes 30 minutes. (Note: The concentration of these solutions are each 100 uM.)
- If necessary, spin the plates in the salad spinner to form all the strand solution to the bottom of the wells.
- Find an un-used lane on an empty 96-well plate. Label it accordingly to indicate that it is in use.
- Prepare three boxes of long-reach 10 uL pipet tips.
- Depending on the type of origami, remove the tips from each of these three boxes and store or discard. Refer to the oligo order forms for which tips to remove – claw will remove tips 5B and 8B, while triangles will remove tips 5A, 5B, 5C, 5E, and 5G, where A – H represent rows and 1 – 12 represent columns.
- Set an 8-well pipetter to 8 uL.
- Remove the top from one of the 96-well plate and carefully place it aside. Place the plate and one of the prepare tip boxes next to each other such that the columns line up.
- For each column of tips, carefully pipet from the corresponding column of the 96-well plate to the previously marked destination column of your empty plate and then discard those tips into the sharps bin. Watch carefully to ensure that each tip actually withdraws and deposits fluid. (Note: Some wells are supposed to be empty; consult with the order forms to determine which.)
- When finished pipetting the 96-well plate, replace the cover and place to the side.
- Repeat the last three steps for the remaining strand plates.
- Replace the cover on the destination plate.
- Using the salad spinner, centrifuge the destination plate to move all solution to the bottom of the well.
- Using a 100 uL pipet transfer all the solution from the destination column to a 2 mL centrifuge tube. Carefully check the bottom of each well to ensure that most of the solution is extracted. When finished, mark the lane of the plate as used.
- Fill the 2 mL centrifuge tube to the 2 mL with (0.22 microfiltered) deionized water. Label the tube with the type of origami (CLAW or TRIANGLE), the date, and “General Staples: 04 uM.”
- Using a 10 uL pipet (and a different tip for each well), pipet 8 uL of each of the wells skipped over in (step 5) into another 2 mL centrifuge tube. Fill this tube to the 2 mLmark with filtered D.D. water.
- Label the tube with the type of origami (CLAW or TRIANGLE), the date, and “Blunt Staples: 0.4 uM.”
- Using a 10 uL pipet (and a different tip for each tube), pipet 8 uL of each of the stick strands into another 2 mL centrifuge tube. Fill this tube to the 2 mL mark with filtered D.D. water.
- Label this tube with the type of origami (CLAW or TRIANGLE), the date, and “Sticky Staples: 0.4 uM.”
- Clean up and return strand plates and sticky strand tubes to the freezer.
Preparation of Origami Solution
Produces solution at 50:1 Staple to Plasmid Ratio, 4 nM Concentration of Plasmid, 1X TAEMg, and total 250 uL of solution per picomole (Note: This protocol assumes 2 pmol of origami are being prepared, and gives volumes to produce that quantity.)
- Take the staple stocks and one or two tubes of plasmid stock from the freezer and allow it to defrost at room temperature. (Note: The NEB stocks are at 0.1 uM.
- While the stocks are defrosting, prepare a 2 mL tube for each type of origami being made and label with the type and date.
- Pipet 20 uL [or 10X] NEB plasmid into each destination tube.
- Pipet 250 [or 125X] uL of 0.4 uM “General Staples” for the appropriate origami design into each destination tube.
- Pipet 250 [or 125X[ of 0.4 uM “Sticky Staples” for the appropriate origami design into each destination tube of STICKY origami.
- Pipet 250 [or 125X] of 0.4 uM “Blunt Staples” for the appropriate origami design into each destination tube of BLUNT origami.
- Pipet 50 uL [or 25X] of (0.22 micro filtered) 10X TAEMg into each destination tube.
- Pipet 50 uL [or 25X] of deionized water into each destination tube.
- Return the remaining staple and plasmid stocks to the freezer.
There are two different methods we use to anneal origami samples; either produces acceptable results for our purposes.
Slow anneals using a hot water bath
- Fill a 2L beaker with tap water. (Using hot water would save time.)
- Place the beaker on the hot plate, turn the intensity to 10, and insert a thermometer.
- Wait for the thermometer to reach 90°C.
- While waiting, inset each reaction tube of interest into a 50 mL tube. To be efficient, double up and insert two reactions tubes into a single 50 mL tube.
- Insert each of these 50 mL tubes into a blue flotation device.
- When the water reaches 90°C, stir the contents with a thermometer and test the upper middle part of the water. (Note: The beaker is typically hotter at the base.)
- When the upper middle part of the water is between 90°C – 94°C, turn off the hot plate, remove the beaker, insert the flotation device with the 50 mL tubes, and weigh it down with an empty glass tray. Wait for the water to reach room temperature. (This will take about four hours, and can be left overnight if needed.)
Rapid anneals using a thermocycler
- Split the origami solution into small PCR tubes. (Note: Put no more than 80 uL in a single tube. Also, use a color-coding system to keep track of the different samples to avoid having to label all of these tubes.)
- Insert the tubes into a PCR machine. The OpenPCR machines will fit 16 tubes at a time.
- Run the appropriate program to begin annealing (eg "CLAW-53deg-15min).
Gels of Triangle Tiles and Extenders
- DNA was obtained in the purified form. (Sets A – A1, A2, A3, A4)
- Stock solutions of 5 uM were made for each strand.
- Dissolve the DNA in 150 uL of D.D. water.
- Heat to 90οC.
- Vortex for 20 seconds and spin down with the benchtop centrifuge for 20 seconds.
- Follow the protocol for the nanodrop to read the concentration.
- Each stock solution was diluted with D.D. water until the concentration reached 5 uM.
- After using them, store the stocks in the freezer.
Formation of DNA Complexes
- The DNA complexes are to be made at concentrations of 1.05 uM.
- Defrost the DNA strand stocks for 15 – 20 minutes at room temperature.
- Take a new 1.5 mL reaction tube and the following and add the following:
Strands Stock Solution 10X TAEMg
- D. Water
A1 + A2 + A3 +A4
- 5 uL of each strand
35 uL 21 uL
Note: The stock concentrations of each strand should be 5 uM and They should be annealed for four hours at 92οC.
- Annealed the sample for four hours.
Formation of DNA Duplexes
- Follow the DNA Preparation Protocol
- Take a new 1.5 mL reaction tube and the following and add the following:
Strand Stock Solution 10X TAEMg DD Water Ah1 + Ah2 60 uL + 60 uL 15 uL 15 uL Av1 + Av2 60 uL + 60 uL 15 uL 15 uL
Note: The stock concentrations of each strand should be 5 uM and The concentration of each duplex should be 2.5 uM.
- Annealed the sample for four hours.
Formation of DNA Complexes – Tiles
- Defrost the Tile A stocks for 15 – 20 minutes at room temperature.
- Take a new 1.5 mL reaction tube and the following and add the following:
Tile A + Ah1/Ah2 5 uL of tile A + 12 uL of Ah1/Ah2 (duplex) Tile A + Av1/Av2 5 uL of Tile A + 12 uL of Av1/Av2 (duplex) Tile + both duplexes 8 uL of tile A + 19.2 uL of each duplex
Note: The concentration of Tile + duplex is 0.31 uM. The concentration Of Tile + duplex is 0.18 uM.
AFM Imaging on the DNA Complex with Both Extenders
Made stocks of complexes: Made up of Tile A (Made up of Brookhaven Strands A1, A2, A3, A4) and its extenders (Av1/Av2 and Ah1/Ah2). Took quantities from stock of Tile A and quantities from stock of paired extenders (Av1/Av2 and Ah1/Ah2). Took 25 uL of Tile A (Previous annealed at 1.05 uM/26.25 pmol)
- Added 60 uL of Ah1/Ah2 (Previously annealed at 2.5 uM/150 pmol)
- Repeat the step for the Ava/Av2.
- The goal is to create an excess of strand quantity compared to complex quantity.
The sample was annealed following annealing protocol. Sample was prepared (Instructions.) The sample was diluted by a factor of 11.
- The sample was not centrifuged.
- The sharp tip (lower frequency is required) was used to image.
Deposited sample for 30 seconds on the mica. (Instructions.) Begin imaging using the AFM.
Dynamic Light Scattering (DLS)
- Follow the protocol for making DNA origami.
- Dilute the Dorigami sample up to 25 nM DNA with TAEMg buffer up to 1X TAEMg.
- Virus Capsid: Dilute up to 300 uM with 0.8 M Sodium Phosphate Buffer at pH = 8.
Preparation of cuvettes
- Pipet 15 uL of each sample into a cuvette.
- Insert the cuvette into the Delsa Nano C Particle Analyzer.
- Set up the software.
- Beckman Coulter v2.21
- Measured Parameters
- Size measurements
- Dust Limit 5
- Upper Dust Limit 10
- Lower Dust Limit 100
- Minimum Intensity 3000
- Pinhole 50 μM
- Analysis Parameters
- Analysis Methold CONTIN
- Fitting Range G2(T)
- G2(T)max 2
- G2(T)min 1.003
- Noise cut level 0.3
- Cell Parameters
- Cell Name Size Cell (Micro)
- Cell Type Size Cell (Micro)
- Correlator Type Log
- Accumulations 100
- Diluent Properties
- Diluent Water
- Refractive Index 1.33
- Viscosity 0.89
- Dielectric Constant 78.3
- Run the instrument.
Forester Resonance Energy Transfer (FRET)
- Pour a 2% agarose gel following the protocol.
- Prepare 11 mM MgCl2
- Prepare 0.5 TBE with 11 mM MgCl2
- Add sufficient MgCl2 to agarose gel mixture up to 11 mM concentration
- Prepare sufficient agarose gel to make a 2% gel with at least 1cm thickness
- Follow standard gel pouring protocol
Running the Instrument
- Using the TYPHOON FLA 9500 scanner, input the settings.
- These settings are for use with ATTO 647N and ATTO 550 dyes
- FRET Fluorescence
- use LPFR (long pass filter) and scan with 532nm laser (green)
- DONOR Fluorescence
- use BPFG (band pass filter green) and scan with 532nm laser
- ACCEPTOR Fluorescence
- use LPFR (long pass filter) and scan with 635nm laser
- Run scans, save images as .gel format.
The highlighted color of the image during scanning is irrelevant as it is false color. The channels will be saved as different layers and shown as overlapped. Select each individual layer to export. Export images to bmp in ImageQuant with 600 dpi for sufficient resolution.
- Prepare two columns by pipetting 250μl neutravidin agarose resin into each centrifuge column.
- Label one the control, which will not be functionalized with A prime strand, and one experimental, which will be functionalized with A prime strand。
- Twist the bottom off columns and spin at 5000xg.
- Wash three times by loading 250μl 1X TAEMg onto column and spinning again at 5000xg at two minutes per wash. Discard all washings.
- Functionalize experimental column with 40 picomoles of A prime strand in 80μl water. Let it sit for ten minutes, then wash six times with 1X TAEMg at two minutes per wash. Discard all washings.
- Apply 1000 picomoles of T15 strand in 80μl water to both control and experimental columns. Let it sit for ten minutes, wash six times with 1X TAEMg. Discard all washings.
- Prepare two sticky claw/blunt triangle mixtures (abbreviated SCBT) by combining 0.5 picomoles sticky claw and 1.5 picomoles blunt triangle in two reaction tubes.
- Apply one mixture to each column, and let it sit for ten minutes.
- Spin down and wash three times with 70 uL 5% formamide 1X TAEMg per wash, two minutes per wash. Keep washings as nonspecific wash. (Note: There should be a volume of 210μl per column.
- Apply 70 ul 1X TAEMg to each column and expose to UV light for fifteen minutes. Spin down for two minutes and apply two additional washes of 70 ul for a total volume of 210μl per column. Keep washings as UV wash.
- In four centrifugal filter units, pipet your washings: control nonspecific wash, control UV wash, experimental nonspecific wash, and experimental UV wash. Spin at 5000xg for ten minutes right-side up.
- Obtain clean outer centrifuge tubes, label each, and place each filter unit upside down in the appropriate tube. Spin for ten minutes at 5000xg.
- Run an agarose gel with a 1kb marker lane, a lane for a fresh and untouched SCBT mixture, and four lanes for resin wash control, resin wash experimental, resin cleave control, and resin cleave experimental.
Making Fab Fragments
Preparation of Buffer
- Preparation of Digestion Buffer: 43.9 mg of cysteine-HCl was dissolved in 10 mL of the supplied IgG-1 Digestion Buffer. After adding the cysteine-HCl, the pH should be around 5.6.
- Preparation of PBS Buffer: A stock solution of 0.1 M PBS Buffer Saline was made. A dilution was carried out to prepare a working solution of 0.01 M PBS Buffer Saline.
- In this experiment, 100 ug of IgG was used to generate Fab fragments.
- 100 uL of this sample which contains 100 ug of IgG was pipeted into an ependurf tube.
- 25 uL of PBS buffer was added to the sample to obtain a final volume of 125 uL.
The columns were prepared following the protocol listed in Pierce Fab Micro Preparation Kit purchased from Thermo Fisher Scientific (Product # 44680).
Generating the Fab Fragments
- Add the 125 uL of sample to the spin column that has the immobilized ficin.
- Incubate the reaction for 10 – 16 hours.
- Remove the bottom cap and place it into a reaction tube. Spin at 5000xg for 1 minute to separate the digest from the immobilized ficin.
- 10 ug of the digest was saved to run on the SDS gel.
- The volume of this is about 12.5 uL.
- Wash the resin with 125 uL of Protein A Binding Buffer. Place the spin column into an Eppendorf tube. Centrifuge the column at 5000 x g for 1 minute. Repeat for a total of 3 washes. Add the wash fractions to the original digest.
Purification of Fab Fragments
- Prepare the Protein A column following the protocol given in the kit. 1.
- Apply the sample to the column and place it into a new 2 mL tube.
- Tighten the yellow cap and invert the column. Incubate at room temperature with mixing for 10 minutes.
- Loosen the top cap and remove the bottom cap. Place the column in a new 2 mL collection tube and centrifuge for 1 minute and collect the flow through. This flow through contains Fab fragments.
- Take 2 30 K spin filters and fill with 500 uL of tween (0.05% w/v in water) and spin at 5,000xg for 5 minutes. The collect for these were discarded and the excess Tween was pipeted out.
- For 40 uL of protein, ¼ is 10 uL.
- For the 30 K columns, 5 minutes yields a volume of 42 uL and a concentration factor of 12.
- Two reaction tubes were prepared with 10 uL of the Fab and 90 uL of PBS.
- The 100 uL samples were pipeted onto the spin columns.
- 400 uL of PBS was added.
- The tubes were spun at 5,000xg for 5 minutes. The collect for these were discarded.
- The spin columns were inverted into new tubes and spun at 1,000xg for 2 minutes.
- Pipet the 40 uL of sample into the 30 K spin filters and add 460 uL of PBS. Repeat the centrifugation steps three more times.
- Protein A
- Pipet 10 uL of each sample into new columns and add 90 uL of PBS.
- Prepare the Protein A colum. Follow the protocol above for Protein A Column
An SDS gel was run on the IgG, digest, protein A flow-through, and protein A wash for both tasks.
Two stocks solutions of crude strands were used, TCA and TCAT3. A fraction of those stocks were purified while the rest was placed back into the freezer. The concentrations of the pure strands were determined using the nanodrop. An estimate of the concentrations of the crude strands was determined using the nanodrop. Coumarin Reaction
- The following concentrations of each component were needed:
DNA Coumarin Catalyst Sodium Phosphate Bufer 10 uM 200 uM 1000 uM 75 mM
- Preparation of Sodium Phosphate Buffer: A solution of sodium phosphate buffer (pH = 7.4) was prepared by dissolving 2.5 g of monosodium phosphate monohydrate and 15.6 g of disodium phosphate heptahydrate.
- Preparation of Coumarin
- A stock solution of 2 mg of coumarin was dissolved in 1 mL of sodium phosphate. The exact concentration was calculated.
- Vortex the solution and shake vigorously to make sure all the coumarin reagent dissolves.
- A diluted solution was made from the stock. The concentration was 400 uM.
- Preparation of Catalyst
- A stock solution of 2 mg of catalyst was dissolved in 1 mL of sodium phosphate. The exact concentration was calculated.
- Vortex the solution to make sure all the catalyst dissolves.
- A diluted solution was made from the stock. The concentration was 2000 uM.
- Preparation of Sample
- A. 100 pmol of pure DNA was used for each experiment. The appropriate volume was pipeted into the reaction tube.
- The sample was evaporated to dryness.
- To the control, 10 uL of catalyst was added. The final concentration of DNA is 10 uM and the final concentration of the catalyst is 1000 uM.
- Pure TCA + TCAT3 Coumarin Reaction
- 10 uL of coumarin was added to each sample.
- 10 uL of catalyst was added to each sample.
- The final concentration of DNA is 5 uM, the final concentration of coumarin is 200 uM, and the final concentration of the catalyst is 1000 uM.
- Deprotection Samples
- For the deprotection samples, the evaporated DNA was first redissolved in 80% acetic acid for 30 minutes.
- Then the sample was desalted.
- Desalting Protocol
- The desalting protocol was carried out using the G-25 spin filters (GE Care)
- First the centrifuge was set to (700xg)
- The top is opened a little to make sure that it is not too tight.
- Break the bottom piece. Take the complementary tube and put the column on it.
- Spin for 2 minutes.
- Make sure the solution all goes down. Throw out the bottom tube.
- Apply the sample to the column and spin at (700xg) for 1 minute.
- Evaporate the sample to dryness.
- Carry out the reaction by adding the coumarin and the catalyst as given above.
- Desalting Samples: Repeat steps 8. (a) to (i) but be sure to add 100 uL of water in order to have volume to place onto the column.
- Repeat the protocol for the crude samples in preparing the crude strands by itself, crude strands + coumarin reaction, crude strands + deprotection, and crude strands + desalting.
- Let the reaction run overnight at 25°C.
- A denaturing gel was run on all the samples. For each sample, 10 pmol was run on a 12% gel. See the above for gel pouring protocol.
Running a Precast SDS Gel
The model of the precast gels that we work with is the Mini-Protean Tetra Cell (Biorad).
Preparing the gel
- The gel is precast or pre-poured. (12% Tris-Glycine) They come in packaged plastic plates with wrapping.
- To prepare the gel, simply cut off the tape along the black line at the bottom of the plate with a razor blade. Note: Be sure not to rip off the entire plastic or the plastic will open up allowing the gel to fall out.
- Remove the comb at the top of the plastic plate gently.
- Wash the wells out with 1X SDS Buffer. (Diluted from the 10X SDS Buffer. Refer to the preparation of buffers section.)
Setting up the Gel Clamp Assembly
- The instrument comes with a plastic container, a gel clamp assembly, and a cover, which contains the plugs for the power supply. Rinse each component with D.D. water. Note: Be sure not to rinse any component with acetone.
- Carefully dry each component with paper towel.
- Lay the gel clamp on a flat surface. Let the arm fall to the sides. Place the plate that contain on one side of the gel clamp. Be sure that the gel is on the interior.
- Also, place another plate on the other side of the gel clamp.
- Gently push them down onto the rubber plastic holder and push them forward so that they rest on the green rubber gasket.
- Gently, clamp one side at a time while making sure that the plates are at the same height on both sides. Note: This step is particularly important and is the main cause of leaks.
- Once both plates are clasped, pour in 1X SDS buffer until it reaches the top of the plates.
- Let the assembly sit for five minutes and be sure that there are no leaks.
Loading the Samples and Running the Gel
- Once it is confirmed that there are no leaks, place lane divider on the assembly.
- Prepare a box of 10 mL long-reach tips and a 10 mL pipet.
- Fill the tip with the proper volume of interest and gently slide the tip down into the lane that is being loaded. Let the tip press against the plate and slowly release the sample into the well. If the sample runs to the bottom, change the angle and press against the plastic to find the lane. Note: The metal syringes should not be used in loading protein samples because proteins adhere to the metal surface.
- Once all the samples are loaded into the wells, carefully place the assembly into the plastic container and remove the lane divider. Be sure to line up the colors of the plugs with the colors on the side of the container.
- Fill the container up to the line for 1 gel with 1X SDS buffer.
- Gently place the cover onto the top of the container making sure that it lines up perfectly with the plugs atop the gel clamp assembly. Connect the plugs to the power supply and run the SDS gel.
Atomic Force Microscopy
- Items required:
- Freshly filtered (through a 0.2 um filter) water
- AFM samples
- Small round adhesive stickers
- Sheet of mica
- Hole Puncher
- 1-10 uL pipettors and 100-1000 uL pipettor with their tips
- Can of dust remover spray
- AFM tips/ tip holder
- Multimode Nanoscope with software
- Laser head
- Put the Multimode Nanoscope on the microscope stage
- Fit and connect the appropriate scanner to the Multimode Nanoscope
- Properly place the laser head onto the scanner and connect it to the Multimode Nanoscope
- Secure the laser head by attaching the springs from the Nanoscope onto the sides of the laser head
- Attach cable from external AFM controller to the AFM
- Turn on computer
- Turn on the external AFM controller
- Choose appropriate options according to AFM software that is being used
- After mounting the tip onto the tip holder, place into the laser head
- Using the knob in the back (cantilever holder clamp), lower the clamps onto the tip holder until a slight resistance is felt
- Use camera attached and knobs on the stage to focus on the cantilever
- Focus the laser on the cantilever using the knobs on the laser head
- Maximize the signal on the photo diode while switch is on AFM/LFM mode
- Adjust the mirror on the back of the laser head
- Move the laser to reposition it on the cantilever
- Using the photo sensor knobs, get the RMS and VERT values as close to zero on the Multimode Nanoscope
- Take a puck and apply an adhesive sticker to provide a sticky surface for the mica
- Hole punch a piece of mica and place it on the [sticky] puck
- Press the mica onto the puck by the edges and hold for 1 minute
- Nothing must touch the center of the mica
- Now that the mica is securely placed on the puck, hold the puck with tweezers
- Take a small piece of tape, place on the mica and peel off
- Lift up piece of tape to the light to make sure that there are no cracks on the mica
- If there are cracks on the tape, use a new piece of tape on the mica
- Continue to place and peel tape on the pica until there are no noticeable cracks on the tape
- Pipette origami solution (about 3-5 uL) onto the center of the mica
- Allow solution time to set (30 seconds to 1 minute)
- Take 1 mL of filtered deionized water and gently squirt small portions onto the mica surface
- Take the edge of a Kimwipe and wick the edges of the mica to soak up excess water
- To dry the mica surface, grab a dust remover spray and gently blow on the mica for 1-2 minutes
- Raise the tip up by using the right AFM switch and flipping it forwards for about 10 seconds
- Remove the laser head carefully by removing springs
- Carefully put mica onto stage
- Put laser head back on and secure it in place with springs
- Lower the tip back down by using the same AFM switch as in the first step and flipping it backwards for about 10 seconds
- Make sure that all settings on the software correlate with the type of mode that is planning to be used
- Follow instructions of the Multimode Nanoscope that is being used to engage the tip to begin scanning the sample
- Once scanning begins, adjust both the integral and proportional gains.
- Raising both gains will allow for better resolution
- Make sure gains are not too high (feedback)
- Proportional gain must always be higher than the integral gains
- If needed, lower the amplitude setpoint voltage to make the cantilever scan with a greater force so that lift off does not happen
- Adjust scan angle and scan speed (usually 2Hz to start with) to maximise image resolution
Gel Imaging for Agarose and Acrylamide
- After running the gel, carefully remove the plugs from the power supply.
- Prepare a staining solution using ethidium bromide (EtBr). Do this by adding 10 – 20 uL of 10mg/ml ethidium bromide into a glass tray. Then fill the tray up to 1 inch with D.D. water. Gently lift the gel from the plate and move it into the tray for staining. Cover the tray with aluminum foil, and let it stain for 10 – 15 minutes.
- To get more sensitivity, Sybr Gold can be used as a stain. The stain can be prepared with 20 uL of Sybr Gold 1000x concentrate and repeating the steps above for staining.
- To image the gel, carefully lift and place the gel on the pre-cleaned UV lightbox; turn the lightbox on
- Open the software for Carestream and capture the image maximizing the intensity without saturating the detector (use apeture/and exposure controls).
Gel Imaging for SDS-Gel
- The stain used for SDS gels is the Oriole stain (Biorad).
- Pour 50 mL of the stain into a plastic tray. Gently slide the gel into the tray for staining.
- Cover the tray with aluminum foil and let it shake gently for 90 minutes inside the incubator. Note: Oriole stain can damage the UV light box. Therefore, pour it out out and replace it with D.D. water, and rinse the gel.
- Image the gel in the same way using the UV lightbox.