LB Agar Plates
1. 1L deionized H2O
2. 20g LB Powder
3. 20g Agar Powder
4. (Optional) Antibiotic
1. 1L deionized H2O
2. 20g LB Powder
3. (Optional) Antibiotic
Murashige and Skoog Plates
1. Obtain a 2L flask and drop a large stir bar inside.
2. Measure 900 mL of ddH2O in a graduated cylinder and dissolve 2.15g of Murashige and Skoog powder.
3. Adjust pH to 5.7 with the gradual addition of KOH and HCl.
4. Add ddH2O to adjust volume to a total of 1000 mL.
5. Add 8g of agar powder.
6. Sterilize the medium by autoclaving on the Liquid 20 setting.
7. Cool the medium to ~60C prior to the addition of antibiotics (as required).
8. Add 1mL of hygromycin antibiotic stock solution (25 mg/mL, filter sterilized) to obtain a final concentration of (25 ug/mL).
9. Mix the medium well for 10 minutes.
10. Carefully pour the plates in a sterile environment by filling ¾ of a plate and swirling to ensure equal distribution of medium.
11. Wrap the plates in a petri plate sleeve and distribute.
Floral Dip Infiltration Media
1. Obtain a 2L flask and drop a large stir bar inside (slowly to not break the glass).
2. Measure 900 mL of ddH2O in a graduated cylinder and dissolve 2.15g of Murashige and Skoog powder.
3. Add 50g of sucrose
4. Adjust pH to 5.7 with the gradual addition of KOH.
5. Add ddH2O to adjust volume to a total of 1000 mL.
6. Add 10 uL of 1mg/mL 6-Benzylaminopurine.
7. Add 200 uL of Silwet L-77.
8. Mix well by using the stir bar for 10-20 minutes.
1. Retrieve overnight bacterial cultures from the shaking incubator in the warm room within 3202.
2. In a microcentrifuge 1.5mL tube, add 1 mL of the overnight culture. Centrifuge the tube at 8000rpm for 2min. Check for a pellet at the bottom and carefully discard the supernatant by pouring it out without disturbing the pellet.
3. While this spins, place 50uL of elution buffer per culture to warm at 65C (same tube).
4. Repeat Step 2 until there is no more culture in your overnight tube. Reserve 2mL if planning to make glycerol stock or do PCR verification. There should be a visible pellet at the bottom with minimal amounts of supernatant. Carefully remove excess supernatant with pipette
5. Resuspend the pellet by adding 250uL of Resuspension solution (Only reagent stored in the fridge). Gently pipette the pelleted cells to be resuspended in the solution. Do not vortex.
6. Add 250uL of the lysis solution to the microcentrifuge tube and mix gently by inverting the tube 4-6 times. Let sit for 2:30 minutes. Do not allow this to sit for more than 5min.
7. Add 350uL of the neutralization solution and immediately invert the tube 4-6 times. The neutralized bacterial lysate should be cloudy. If it’s not cloudy, the kit may be expired.
8. Centrifuge for 5min at 12000 rpm on a tabletop microcentrifuge. Cell debris will be pelleted down the side of the microcentrifuge tube.
9. Transfer the supernatant to the supplied GeneJET spin column by decanting or pipetting. Avoid disturbing or transferring the white precipitate. Let sit for 2 minutes.
10. Centrifuge the column for 1 min at 12000 rpm. Discard the flow through. 11. Add 500uL of wash solution and centrifuge for 1 min at 12 000rpm. Discard flowthrough. Repeat
12. Discard the flow-through and centrifuge for an additional 1 min to remove residual Wash Solution. This step is essential to avoid residual ethanol in plasmid preps.
13. Transfer the GeneJETspin column into a fresh 1.5 mL microcentrifuge tube (not included). Add 50 μL of the Elution Buffer to the center of GeneJETspin column membrane to elute the plasmid DNA. Take care not to contact the membrane with the pipette tip. Incubate for 2 min at room temperature and centrifuge 12000 rpm for 2 min.
14. Pipette the elution back into the spin column and centrifuge again for 2 min at 12000rpm. The flowthrough is the final product of the plasmid miniprep.
15. Test the purity and concentration of the plasmid DNA using the nanodrop and use elution buffer as the blank. Record A260/A280 value and concentration on side of tube and Benchling. 16. Store the purified plasmid DNA at -20°C. Label the concentration on the side of the tube.
Polymerase Chain Reaction
1. Prepare 75ul of colony water, vortex to mix.
2. Obtain the tabled reagents required for your PCR and let them thaw at room temperature and place them back into the freezer once finished.
3. Calculate how much total of each reagent you need and prepare a Master Mix (MM) by combining all reagents into one tube (except for colony water). Make enough MM for 2-3 extra. You may need to make a separate master mix if you are also doing agrobacterium
4. Mix the PCR tube by pipetting up and down and do a spin down for 3-5 seconds to ensure all the liquid is pooled at the bottom.
5. Use the same thermocycler settings as the previous protocol
Glycerol Stock Preparation
1. Create a sterile environment by wiping the benchtop with 70% ethanol and paper towel and lighting a Bunsen burner once the ethanol has completely dried out.
2. Within the sterile region, obtain a 2mL cryovial (do NOT use an Eppendorf tube as they will explode when thawing them) and dispense 950µL of liquid culture (from a 3mL overnight culture) into the vial for preservation.
3. Then add 950µL of the 40% glycerol solution and invert for 2 minutes.
4. Store the newly created (20%) glycerol stock in a -80˚C freezer
1. Measure out 100 mL of 1x TAE into a graduated cylinder and pour into a 250 mL Erlenmeyer flask.
2. Weigh out 1g of agarose in a weigh boat and pour into the 250 mL Erlenmeyer flask.
3. Place a couple of folded up Kimtech wipes to cover the top of the flask and reduce the chance of the liquid to boil over.
4. Microwave the flask for 1 minute and swirl for 5 seconds.
5. Repeat step 4 until no particles remain and carefully monitor the solution to not boil over. You can microwave for less than 45 seconds if you do not think you need to.
6. Let the flask cool until warm to the touch but not for too long because the gel will harden! If gel hardens, you may microwave it again UNLESS you have already added RedSafe. RedSafe CANNOT go into the microwave.
7. When warm, add 5.0 uL of RedSafe to allow intercalating of the DNA for visibility under UV. Swirl vigorously.
8. Obtain a cassette and tape the open ends of the cassette using masking tape. Make sure that there is a good seal to prevent spillage.
9. Place a comb in the cassette for the number of required wells and pour the gel into the cassette.
10. Wait 30 min to allow the gel to cast. Once completed, remove the masking tape.
11. Take up the dyed-PCR product and load 10-12µL of it in each well, starting from the second well. Be careful not to puncture the gel by getting too close but ensure that the pipette tip is inside the well.
12. Pipette 10µL of PCR reaction directly into the gel wells (one gel per reaction). Put the lid on the apparatus and double check that it’s: “Running to red”. Turn on the power supply and clock the voltage to ~115V. Tiny bubbles should be forming on each end of the apparatus in the TAE buffer. The gel will take ~40 mins of running until completion. The dye should be 2/3 of the way down the gel when it is ready to be imaged
13. Turn off the power supply and unplug lid.
14. Take the gel to a gel reader and interpret the results.
A. tumefaciens Competent Cell Preparation
1. Grow fresh colonies on a LB-gentamycin plate at 28C for 24h.
2. In the morning, pick a colony from Day 1 to make a 5 ml liquid starter culture using LB + gentamicin medium. Allow 24hrs for growth shaking at 28C.
3. Make a large 200 mL liquid culture, add 2 mL of starting culture into LB medium. Place the culture back into the 28C incubator.
4. Watch the OD value change. The OD will reach 0.4-0.5 usually after 3-4 hours (need to check a few times, especially for the first time). Do NOT let the OD value go over 0.5.
5. Chill the flask on ice for 10-15 min. (From this step, always keep the cells on ice at 4C, IMPORTANT!)
6. Transfer to a 50ml tube, and spin at 4500 rpm for 15 min at 4C, discard supernatant and leave on ice.
7. Add 5ml pre-chilled 20 mM CaCl2 into each tube, resuspend gently using a pipette tip, do NOT vortex.
8. Centrifuge again, at 4500 rpm for 10 min at 4C, discard supernatant via pipetting, and add 1ml 20 mM CaCl2 containing 15% glycerol (850 ul CaCl2 + 150 ul glycerol) into each tube. There will be 4 mL of solution in four tubes. Resuspend well.
9. Dispense 100ul aliquot to 1.5 ml Eppendorf tube (pre-cold on ice). Try to avoid touching the bottom of the tube with your hand and keep the cells cold.
10. Drop all 1.5ml tubes containing cells into liquid nitrogen to quickly freeze and store in a - 80C freezer.
E. coli TSS Chemically Competent Cell Preparation
1. Grow a 5 mL seed culture of cells in LB medium to saturation overnight. Dilute this culture back into 25–50 mL fresh LB in a 250 mL Erlenmeyer flask. You should aim to dilute the seed culture by at least 1/100.
2. Grow the diluted culture to an OD600 = 0.2–0.5. (You will get a very small pellet if you grow 25 mL to OD600 = 0.2-3. Put microfuge tubes on ice now so that they are cold when the cells are aliquoted into them later. If your culture is X ml, you will need X tubes. At this point you should also make sure that your TSS is being chilled (it should be stored at 4°C but if you have just made it fresh, then put it in an ice bath).
3. Split the culture into two 50 mL falcon tubes and incubate on ice for 10 min. All subsequent steps should be carried out at 4°C and the cells should be kept on ice wherever possible
4. Centrifuge for 10 min at 3000 rpm and 4°C. 2. Remove supernatant. The cell pellets should be sufficiently solid that you can just pour off the supernatant if you are careful. Pipette out any remaining media.
5. Resuspend cells in chilled TSS buffer. The volume of TSS to use is 10% of the culture volume that you spun down. Keep an eye out for small cell aggregates even after the pellet is completely off the wall. Pipette up and down gently to dissolve all of the cells. Chill on ice for 15 minutes.
6. Add 50-200 μL aliquots to your chilled microfuge tubes and flash freeze with either ethanol-dry ice or liquid nitrogen. Store at -80°C.
E. coli Transformation
1. If you are doing a Golden Gate reaction, you must add 1µL of Bsa1 into each reaction tube and incubate in the thermocycler at 37ºC for 1 hour and 85ºC for 20 minutes.
2. Obtain and label an appropriate number of sterile 1.5 mL tube. You can use the tube that the cells are currently in as a reaction tube because each frozen cell tube has 100µL and you only need 50µL per reaction. So, the number of tubes you take out should be half of the number of DNA samples you are running. Pre-chill the tubes on ice.
3. Take competent DH5a cells on ice taken from the -80ºC freezer (100µL total) and immediately place on ice (bring the ice bucket with you to the -80ºC freezer). They take 10-15min to thaw. Only thaw on ice and NOT with your hands, these cells are sensitive.
4. Monitor the thawing progress regularly. IMMEDIATELY after the cells have thawed, Pipette 50 µL of competent cells into the sterile chilled 1.5 mL tubes so each tube has 50µL of competent cells, and QUICKLY Pipette 10-50ng of DNA into the competent cells. If this is DNA from a Golden Gate reaction, pipette all of the reaction volume (varies between 10-15µL, so make sure you adjust you pipette as you go to get all of it). The cells can lose their competency if they sit on the ice for too long, so it is essential you get the DNA into the tubes as fast as possible.
5. Mix cells gently by pipetting up and down. DO NOT VORTEX.
6. Incubate cells and DNA in 1.5 mL tubes on ice for 30 minutes. While this is happening, warm a 50mL conical tube of LB at 37ºC.
7. Heat shock cells at 42ºC in a hot plate or water bath for 45 seconds. NOTE: Make sure the time is exactly 45 seconds.
8. Immediately let the cells incubate ice for 5 minutes
9. Add 250 µL of SOC (ideal) or LB media (not so ideal) into each transformation tube.
10. Incubate cells at 37ºC for 1 hour and shake at 200 rpm.
11. Warm up LB kanamycin plates in the 37ºC incubator to reduce condensation and avoid temperature shocking the cells too much. Ensure plates are agar-side up
12. Pipette (do not pour) all of the reaction mixture onto a single plate.
13. Spread the culture using a sterile glass spreader.
14. Leave plates cracked open beside a flame until dry if needed, or place inside the biosafety hood if you have been shown how. This may take more than 30 minutes. You can come back to the plates every so often to spread around the remaining liquid by tilting the plates (you do not need the glass spreader), which will increase the surface area of the liquid and help it dry faster.
15. Parafilm the plates and incubate transformations overnight at 37ºC agar-side up to avoid condensation.
A. tumefaciens Transformation
1. Obtain the isolated plasmid from the -20ºC freezer to be taken up by the competent Agrobacterium and let thaw on ice. Calculate how many mL of plasmid you will need to obtain 1µg per reaction before taking out the competent cells to thaw.
2. Obtain a 1.5mL tube containing competent Agrobacterium cells (~100µL cells per tube) per reaction from the -80ºC freezer. When thawed, split it into two tubes such that they have ~50µL of competent cells each
3. Dispense 1 ug of plasmid into each tube of competent cells.
Note: To find the volume of the pure plasmid to dispense, use c=n/v. Generally you will always be using more than 3-5µL of plasmid
4. Mix the combined solution via flicking and let sit of ice for 15 min.
5. Cold shock the Agrobacterium by placing the chilled tube in the -80ºC freezer for 2min. You can look for some piles of snow in the freezer and use those as the snow will facilitated cooling much faster
Note: If liquid nitrogen is accessible, submerse the tube in liquid nitrogen for 1 min for slightly better results.
6. Incubate the mixture on a 37ºC hot plate for 3 min.
7. Add 300 uL of nutrient rich SOC broth to the mixture and incubate at 28ºC on a shaker for 2 hours at 200rpm.
Note: If SOC broth is unavailable LB broth should work with reduced efficiency.
8. Dispense ALL of the transformed Agrobacterium mixture onto a single LB plate containing Gentamicin and Kanamycin antibiotics as selective markers (Confirm in advance the resistance that the transformed bacterium should exhibit based on the experiment design).
9. Spread the plate using a sterile glass hockey stick to evenly distribute the liquid culture.
10. Seal the plates with parafilm and incubate at 28ºC for 48 hours.
Golden Gate Cloning
1. Grab a bucket of ice and put in the following reagents EXCEPT for the NEB GG Mix BsaI. This tubes contains both BsaI and ligase. You must add in the reagents in the order you see below (e.g water first, plasmid second, etc). We only take out enzymes right when we are about to use them as they quickly denature at room temperature. All of your tubes should be kept on ice for the entirety of this protocol, so get a wide bucket to fit your tube rack.
2. Once you have added in all of your reagents (other than the enzyme mix), grab NEB GG mix BsaI from the freezer and quick add it to your reactions. Be sure not to touch the bottom of the tube where the enzyme sits.
3. Once finished, place the NEB GG Mix Bsa-I quickly back in the freezer.
Thermocycler settings for BOTH protocols:
2 min @ 37 degrees
5 min @ 16 degrees
Repeat steps 1 & 2 (99 times)
15 min @ 65 degrees
20 min @ 80 degrees
Infinite time for 4 degrees
4. The reaction is performed for approx. 13 hrs
5. 1µL of Bsa1 must be added to each reaction mixture when the entire run is complete. This is to cut up any non-cloned plasmid. The person performing the transformation will be doing this the following day.
1. Pick up a single colony on “day 1” to create a 5 ml overnight culture to be incubated at 28 degrees. On “day 2”, make a large volume of overnight culture (300 mL) using 1-5 mL of starting culture. On “day 3”, check the culture’s OD value, which should be between 1.2-1.6 when using a spectrophotometer.
2. Centrifuge at 4500-5000rpm for 20 min at room temperature, discard supernatant, and then suspend with infiltration medium to a final OD = 0.8. Be sure to label your centrifuge bottles with the strain. You should be able to calculate how much of the culture you will need and how much infiltration media to make by knowing that OD is linearly proportional to concentration.
3. Get the plants from the phytotron. Ensure they are properly prepared and the appropriate structures have been cut (see “Preparation of Plants” up top).
4. Let the plants sit upside down and immerse all the stem and flower buds into the medium. Do NOT immerse the plants for more than 30 seconds. You can use the same infiltration medium for multiple plants.
5. Prepare a growth tray, put a bench paper on the bottom to let the paper absorb the residual medium. After 30 seconds, take the plants out, and lie down on some paper towel, cover with a black tray on the top, and leave them in a dark location (No direct sunlight) for 24 hours. Go to the phytotron to get the blue plant markers and label each of your plants
6. Move the plants to a growth chamber set to normal conditions (28C).
1. Turn on the UV lamp in the biosafety cabinet for 5 minutes.
2. While the biosafety cabinet is sterilizing, obtain the following reagent and instruments:
MS media petri plates, 20% Bleach, Aliquoted Sterile MilliQ water, P1000 and P200 pipettes and pipette tips, 0.1% agar solution, Tube rack
Ensure that the listed materials are all sterilised with 70% ethanol prior to use (excluding the petri plates)
3. Ensure that the microfuge tube contains seeds at the 0.1 mL line (one microfuge tube filled to 0.1 mL is enough for a single MS/Hygromycin plate). If you use a 15 mL conical fill the tube up to the 1mL line (but this means all the seeds in the 15 mL tube will need to be split into 10 plates because 0.1mL of seeds is the max number of seeds that should go onto one plate).
4. Add 75% ethanol to the Eppendorf to fill it up halfway, then flick to ensure that seeds are evenly distributed in the ethanol.
5. Lay the microfuge tube sideways inside the cabinet for 2 minutes
6. Centrifuge the seeds outside of the cabinet for 5 seconds using the short or pulse button. Seeds should be pelleted at the bottom.
7. Return to the cabinet and wipe hands and tube with ethanol.
8. Remove the remaining ethanol via pipetting. Avoid disturbing the seeds.
9. Pipette 750 uL of 20% bleach into the microfuge tube and flick to distribute seeds in the bleach. Lay the tube in the cabinet sideways for 10 minutes while flicking every minute.
10. Centrifuge the seeds outside of the cabinet for 5 seconds using the short or pulse button. Seeds should be pelleted at the bottom.
11. Return to the cabinet and wipe hands and tube with ethanol.
12. Remove the bleach via pipetting. Avoid disturbing the seeds but it is likely that some will be taken up your pipette.
13. Add 750 uL of water to the tube, mix by flicking and centrifuge for 5 seconds. Wipe hands and microfuge tube with ethanol and repeat 4 additional times.
14. Remove the water via pipetting. Avoid disturbing the seeds.
15. Add 1 mL of 0.1% agar solution per 0.1 mL of seeds and mix by gently pipetting up and down.
16. Plate droplets on the MS plate, ensure that all the seed-agar suspension is used.
17. Take a pipette tip or sterile toothpick to spread the seeds evenly over the surface of the MS media. Be careful not to puncture the agar.
18. Parafilm the plate and place it on shelf 4.3 in the tissue culture room in the phytotron. If seeds were just harvested the same day as sterilization, they need to be placed in the 4C fridge for 3-4 days before being moved in order to stratify the seeds.
1. Make a little boat out of a sheet of paper (fold up each side, doesn’t need to be perfect).
2. Remove siliques from the plant while working above/on the boat to catch as many seeds as possible.
3. Remove siliques from the plant either one at a time or by running pinched fingers up the stem of the plant. The siliques should break open easily if they were dried out properly. Half of the silique may stay attached to the stem so ensure all seeds are collected.
4. If some siliques did not open, bunch them up in your fingers and shuffle them around to break open.
5. Shuffle the boat back and forth to separate siliques and other plant matter away from seeds. Remove siliques and other plant matter using fingers or tweezers.
6. Repeat shaking and removal of siliques until all non-seed plant matter is removed.
7. Pour seeds into a labelled Eppendorf tube and store at 4C for stratification.
8. Use a new sheet of paper for each different type of transformant.
RNA Extraction (Qiagen RNeasy Plant Mini Kit)
1. β-Mercaptoethanol (β-ME) must be added to Buffer RLT or Buffer RLC before use. Add 10 µl β-ME per 1 ml Buffer RLT or Buffer RLC. Dispense in a fume hood and wear appropriate protective clothing. Buffer RLT or Buffer RLC containing β-ME can be stored at room temperature for up to 1 month.
2. Buffer RPE is supplied as a concentrate. Before using for the first time, add 4 volumes of ethanol (96–100%) as indicated on the bottle to obtain a working solution.
3. Determine the amount of plant material. Do not use more than 100 mg. Weighing tissue is the most accurate way to determine the amount.
4. Immediately place the weighed tissue in liquid nitrogen, and grind thoroughly with a mortar and pestle. Decant tissue powder and liquid nitrogen into an RNAse-free,liquid-nitrogen–cooled, 2 ml microcentrifuge tube (not supplied). Allow the liquid nitrogen to evaporate, but do not allow the tissue to thaw. Proceed immediately to next step
5. RNA in plant tissues is not protected until the tissues are flash-frozen in liquid nitrogen. Frozen tissues should not be allowed to thaw during handling. The relevant procedures should be carried out as quickly as possible.
6. Add 450 µl Buffer RLT or Buffer RLC to a maximum of 100 mg tissue powder. Vortex vigorously. A short 1–3 min incubation at 56°C may help to disrupt the tissue. However, do not incubate samples with a high starch content at elevated temperatures, otherwise swelling of the sample will occur. Note: Ensure that β-ME is added to Buffer RLT or Buffer RLC before use.
7. Transfer the lysate to a QIAshredder spin column (lilac) placed in a 2 ml collection tube, and centrifuge for 2 min at full speed. Carefully transfer the supernatant of the flow-through to a new microcentrifuge tube (not supplied) without disturbing the cell-debris pellet in the collection tube. Use only this supernatant in subsequent steps. It may be necessary to cut off the end of the pipette tip to facilitate pipetting of the lysate into the QIAshredder spin column. Centrifugation through the QIAshredder spin column removes cell debris and simultaneously homogenizes the lysate. While most of the cell debris is retained on the QIAshredder spin column, a very small amount of cell debris will pass through and form a pellet in the collection tube. Be careful not to disturb this pellet when transferring the lysate to the new microcentrifuge tube.
8. Add 0.5 volume of ethanol (96–100%) to the cleared lysate, and mix immediately by pipetting. Do not centrifuge. Proceed immediately to the next step. Note: The volume of lysate may be less than 450 µl due to loss during homogenization. Note: Precipitates may be visible after addition of ethanol. This does not affect the procedure.
9. Transfer the sample (usually 650 µl), including any precipitate that may have formed, to an RNeasy spin column (pink) placed in a 2 ml collection tube (supplied). Close the lid gently, and centrifuge for 15 s at ≥8000 x g (≥10,000 rpm). Discard the flow-through. Reuse the collection tube in the next step. If the sample volume exceeds 700 µl, centrifuge successive aliquots in the same RNeasy spin column. Discard the flow-through after each centrifugation.
10. Add 700 µl Buffer RW1 to the RNeasy spin column. Close the lid gently, and centrifuge for 15 s at ≥8000 x g (≥10,000 rpm) to wash the spin column membrane. Discard the flow-through. Reuse the collection tube in the next step. Note: After centrifugation, carefully remove the RNeasy spin column from the collection tube so that the column does not contact the flow-through. Be sure to empty the collection tube completely.
11. Add 500 µl Buffer RPE to the RNeasy spin column. Close the lid gently, and centrifuge for 15 s at ≥8000 x g (≥10,000 rpm) to wash the spin column membrane. Discard the flow-through. Reuse the collection tube in the next step.
12. Add 500 µl Buffer RPE to the RNeasy spin column. Close the lid gently, and centrifuge for 2 min at ≥8000 x g (≥10,000 rpm) to wash the spin column membrane. The long centrifugation dries the spin column membrane, ensuring that no ethanol is carried over during RNA elution. Residual ethanol may interfere with downstream reactions. Note: After centrifugation, carefully remove the RNeasy spin column from the collection tube so that the column does not contact the flow-through. Otherwise, carryover of ethanol will occur.
13. Optional: Place the RNeasy spin column in a new 2 ml collection tube (supplied), and discard the old collection tube with the flow-through. Close the lid gently, and centrifuge at full speed for 1 min. Perform this step to eliminate any possible carryover of Buffer RPE, or if residual flow-through remains on the outside of the RNeasy spin column.
14. Place the RNeasy spin column in a new 1.5 ml collection tube (supplied). Add 30–50 µl RNAse-free water directly to the spin column membrane. Close the lid gently, and centrifuge for 1 min at ≥8000 x g (≥10,000 rpm) to elute the RNA. 12. If the expected RNA yield is >30 µg, repeat step 11 using another 30–50 µl RNAse-free water, or using the eluate from step 13 (if high RNA concentration is required). Reuse the collection tube from step 13. If using the eluate from step 13, the RNA yield will be 15–30% less than that obtained using a second volume of RNAse-free water, but the final RNA concentration will be higher.
DNase Digestion (Qiagen)
1. Mix the following in a microcentrifuge tube: ≤87.5 µl RNA solution (contaminated with genomic DNA), 10 µl Buffer RDD, 2.5 µl DNase I stock solution.
2. Make the volume up to 100 µl with RNAse-free water. The reaction volumes can be doubled if necessary (to 200 µl final volume).
3. Incubate on the benchtop (20–25°C) for 10 min.
4. Proceed with the RNA Cleanup protocol.
RNA Cleanup (Qiagen RNeasy Plant Mini Kit)
1. Add 350 µl Buffer RLT to sample from DNase Digestion and mix well.
2. Add 250 µl ethanol (96–100%) to the diluted RNA, and mix well by pipetting. Do not centrifuge. Proceed immediately to the next step.
3. Transfer the sample (700 µl) to an RNeasy Mini spin column placed in a 2 ml collection tube (supplied). Close the lid gently, and centrifuge for 15 s at ≥8000 x g (≥10,000 rpm). Discard the flow-through. Reuse the collection tube in the next step. Note: After centrifugation, carefully remove the RNeasy spin column from the collection tube so that the column does not contact the flow-through. Be sure to empty the collection tube completely.
4. Add 500 µl Buffer RPE to the RNeasy spin column. Close the lid gently, and centrifuge for 15 s at ≥8000 x g (≥10,000 rpm) to wash the spin column membrane. Discard the flow-through. Reuse the collection tube in the next step.
5. Add 500 µl Buffer RPE to the RNeasy spin column. Close the lid gently, and centrifuge for 2 min at ≥8000 x g (≥10,000 rpm) to wash the spin column membrane. The long centrifugation dries the spin column membrane, ensuring that no ethanol is carried over during RNA elution. Residual ethanol may interfere with downstream reactions. Note: After centrifugation, carefully remove the RNeasy spin column from the collection tube so that the column does not contact the flow-through. Otherwise, carryover of ethanol will occur.
6. Optional: Place the RNeasy spin column in a new 2 ml collection tube (supplied), and discard the old collection tube with the flow-through. Close the lid gently, and centrifuge at full speed for 1 min. Perform this step to eliminate any possible carryover of Buffer RPE, or if residual flow-through remains on the outside of the RNeasy spin column after step 5.
7. Place the RNeasy spin column in a new 1.5 ml collection tube (supplied). Add 30–50 µl RNAse-free water directly to the spin column membrane. Close the lid gently, and centrifuge for 1 min at ≥8000 x g (≥10,000 rpm) to elute the RNA.
8. If the expected RNA yield is >30 µg, repeat step 7 using another 30–50 µl RNAse-free water, or using the eluate from step 7 (if high RNA concentration is required). Reuse the collection tube from step 7. If using the eluate from step 7, the RNA yield will be 15–30% less than that obtained using a second volume of RNAse-free water, but the final RNA concentration will be higher.