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RNA extraction-PDF

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This is a protocol for the acid-phenol extraction of RNA from E. coli, used in the Endy Lab. Contact: Heather Keller or Caitlin Conboy

Contents

Set-up

  1. Set sand bath to 100°C.
  2. Set H2O bath to 67°C.

Prepare RNase-Diminished bench

  1. Wipe bench and pipettes with EtOH.
  2. Wipe with Rnase Zap wipes.
  3. Wipe with wet (H20) paper towels (2x).

Culture growth and sample collection

  1. Samples from continuous culture in chemostat:
    1. See Chemostat protocol
    2. At desired time, measure OD600 of chemostat culture
    3. Pull 2mL sample from chemostat through bubbler line. Immediately proceed to cell collection.
  2. Samples from batch culture:
    1. Inoculate 5mL standard media (see protocol, M9_recipe) from fresh plate (<2 weeks). Grow to saturation at 37° C, approx 20 hr.
    2. Measure OD600 of saturated cultures. All should be ~2.5.
    3. Dilute all saturated cultures to OD600=0.0025 (dilution ~1:1000) into 5mL fresh media. (5 uL of culture if OD600=2.5).
    4. Grow to mid-log, OD600 ~ 0.4
    5. Pull 2mL sample and proceed immediately to cell collection.

Cell collection

  1. Immediately combine 0.8mL cold stop solution with each 1mL sample in a pre-chilled centrifuge tube. (Stop solution= 5% H2O saturated phenol in ethanol).
  2. Store on ice while plating dilutions of culture for CFU count. Plate three replicates of a 1:10^4 and a 1:10^5 dilution.
  3. Centrifuge samples at 4°C in Sauer Lab tabletop centrifuge. 13,000rpm, 10 minutes.
  4. Decant the supernatants from the pellets and discard.
  5. (optional) At this point, the pellets can be frozen at -80°C until ready to proceed with step 4. Thaw on ice before lysis.

Lysis

  1. Resuspend each pellet in 0.5 ml of Lysis buffer (2% SDS and 4 mM EDTA).
  2. Incubate the cells in a 100°C sand bath for 3 minutes to lyse the cells.
  3. Add 15 ul of 3M NaOAc to each tube and transfer to ice.
  4. (optional) spike in control mRNA to evaluate the efficiency of extraction

Phenol Extraction with heat

  1. Add an equal volume (0.5 mL) of water-saturated phenol to each tube.
  2. Invert several times to mix.
  3. Transfer to a 67°C water bath for 6 minutes and invert every 40 seconds.
  4. Immediately transfer to ice.
  5. Centrifuge in an Eppendorf 5414 table top microcentrifuge (15,000 RPM) at 4°C for 10 minutes.
  6. Transfer as much of each aqueous layer as possible to new tubes. The aqueous layer is the upper layer and is distinct from the organic phase which is tinted yellow by 8-Quinolinol in the phenol

Phenol/Chloroform Extraction

  1. Add an equal volume (0.5 mL) of water-saturated phenol:chloroform:isolamyl alcohol (25:24:1) to each tube.
  2. Invert several times to mix.
  3. Centrifuge in an Eppendorf 5414 table top microcentrifuge (15,000 RPM) at 4°C for 5 minutes.
  4. Transfer as much of each aqueous layer as possible to new tubes.
  5. Repeat phenol/chloroform extraction once.

Ethanol Precipitation

  1. To each tube, add: 1/10 volume (50 ul) 3M NaOAc, 1/10 volume (50 ul) 1 mM EDTA, and 2-2.5 volumes (1 mL) of cold, 100% ethanol.
  2. Invert to mix and then incubate at -80°C for 20 minutes.
  3. Centrifuge in an Eppendorf 5414 table top microcentrifuge (15,000 RPM) at 4°C for 25 minutes.
  4. Decant the ethanol from the pellets and discard.
  5. Wash each pellet in 1 ml of cold 80% ethanol.
  6. Centrifuge in an Eppendorf 5414 table top microcentrifuge (15,000 RPM) at 4°C for 5 minutes.
  7. Decant the ethanol from the pellets and discard.
  8. Repeat wash with cold 80% EtOH twice, for a total of three washes.
  9. Dry the pellets in a 37°C heat block after the final wash is removed, ~15 min
  10. Resuspend each pellet in 50 ul of Buffer EB.

Absorbance reading

  • Nanodrop (make sure to set for RNA). Record concentration and A260/A280 ratio. An ideal purity has an A260/A280 between 1.8 and 2.2.

Storage

  • If the sample is to be treated immediately with DNase, proceed to the next section. Otherwise, dilute and/or aliquot the samples if necessary and store at -20°C until needed.

DNase treatment

  1. Combine:
    • 50 uL of 10x DNase I Buffer
    • 50 uL of RNA Extraction product
    • 1 uL of Superase Inhibitor
    • 2 uL DNase I (~1 ug enzyme)
    • 397 uL of H2O (RNase-free) to total volume of 500 uL
  2. Incubate 10 min at 37°C.
  3. Optional: If using NEB DNase I, heat inactivate 10 min at 75°C.

Phenol/Chloroform Extraction

  1. Add an equal volume (0.5 mL) of water-saturated phenol:chloroform:isolamyl alcohol (25:24:1) to each tube.
  2. Invert several times to mix.
  3. Centrifuge in an Eppendorf 5414 table top microcentrifuge (15,000 RPM) at 4°C for 5 minutes.
  4. Transfer as much of each aqueous layer as possible to new tubes.

Ethanol Precipitation

  1. To each tube, add: 1/10 volume (50 ul) 3M NaOAc, 1/10 volume (50 ul) 1 mM EDTA, and 2-2.5 volumes (1 mL) of cold, 100% ethanol.
  2. Invert to mix and then incubate at -80°C for 20 minutes.
  3. Centrifuge in an Eppendorf 5414 table top microcentrifuge (15,000 RPM) at 4°C for 25 minutes.
  4. Decant the ethanol from the pellets and discard.
  5. Wash each pellet in 1 ml of cold 80% ethanol.
  6. Centrifuge in an Eppendorf 5414 table top microcentrifuge (15,000 RPM) at 4°C for 5 minutes.
  7. Decant the ethanol from the pellets and discard.
  8. Repeat wash with cold 80% EtOH twice, for a total of three washes.
  9. Dry the pellets in a 37°C heat block after the final wash is removed, ~15 min.
  10. Resuspend each pellet in 30 ul of nuclease free water if to be analyzed by RPA or RT-PCR or in 10 uL Buffer EB if to be used in Northern Blot.

Absorbance reading

  • Nanodrop (make sure to set for RNA). Record concentration and A260/A280 ratio. An ideal purity has an A260/A280 between 1.8 and 2.2.

Storage

  • Store at -20°C until needed.

Knight:RNA electrophoresis-PDF

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in progress! has not been tested

Contents

Overview

Electrophoresis permits assessment of RNA by size and amount. In general, electrophoresis of RNA is done as a step prior to Northern analysis. However, this protocol is for visualizing the RNA in the gel. Since this is a denaturing protocol, the RNA can be assessed both for quality and size.

Materials

Reagents

  • RNAse free water (make lots for rinsing glassware and electrophoresis chambers)
    • Add DEPC to final concentration of 0.1%.
    • Incubate 1hr at 37°C.
    • Autoclave for 15 mins at 15 psi.
  • 10X BPTE electrophoresis buffer (The final pH of this 10x buffer is ~6.5.)
    • 100 mM PIPES
    • 300 mM Bis-Tris
    • 10 mM EDTA
  • HPLC grade or better DMSO
  • Glyoxal
    • Commercially available stock solutions of glyoxal contain both hydrated forms of glyoxal and oxidation products that can degrade RNA. These must be removed.
  • Glyoxal reaction mixture (divide into small aliquots and store at -70°C)
    • 6mL DMSO
    • 2mL deionized glyoxal
    • 1.2mL of 10X BPTE electrophoresis buffer
    • 0.6mL of 80% glycerol
  • RNA size marker
  • RNA gel loading buffer
    • 95% deionized formamide
      • Purchase a distilled deionized preparation of formamide and store in small aliquots under nitrogen at -20°C.
    • 0.025% (w/v) bromophenol blue
    • 0.025% (w/v) xylene cyanol FF
    • 5 mM EDTA (pH 8.0)
    • 0.025% (w/v) SDS

Equipment

  • 37 °C incubator
  • 55 °C water bath
  • Ice water bath
  • Electrophoresis apparatus

Procedure

Prepare RNase free water

  1. Add DEPC to final concentration of 0.1% to H2O
  2. Incubate 1hr at 37°C.
  3. Autoclave for 15 mins at 15 psi.

Prepare BPTE electrophoresis buffer

  1. Prepare 10X buffer by adding the following to 90 ml of distilled H2O
    • 3 g of PIPES (free acid)
    • 6 g of Bis-Tris (free base)
    • 2 ml of 0.5 M EDTA
  2. Treat the solution with final concentration of 0.1% DEPC for 1 hour at 37°C
  3. Autoclave.

Dilute 10X buffer 10-fold with RNase free H2O.

Prepare glyoxal reaction mixture

  1. 6mL DMSO
  2. 2mL deionized glyoxal
  3. 1.2mL of 10X BPTE electrophoresis buffer
  4. 0.6mL of 80% glycerol

Divide into small aliquots and store at -70°C.

Prepare RNA gel loading buffer

  1. Mix the following
    • 95% deionized formamide
      • Purchase a distilled deionized preparation of formamide and store in small aliquots under nitrogen at -20°C.
    • 0.025% (w/v) bromophenol blue
    • 0.025% (w/v) xylene cyanol FF
    • 5 mM EDTA (pH 8.0)
    • 0.025% (w/v) SDS

Denature RNA samples

  1. Mix 10 μL glyoxal reaction mixture with 1-2 μL RNA (up to 10 μg).
  2. Also mix 10 μL glyoxal reaction mixture with RNA size marker.
  3. Incubate RNA samples at 55°C for 1 hr.
  4. Chill RNA samples for 10 mins in ice water.
  5. Centrifuge 5 seconds to collect liquid at the bottom of the tubes.

Cast gel

Do this step during 1 hr denaturation of samples.

  1. Cast 1.5% agarose gel in 1X BPTE electrophoresis buffer.
  2. Use a comb with at least 4 extra lanes for size markers and running dyes.
  3. Place gel in chamber.
  4. Cover with 1X BPTE electrophoresis buffer to cover gel to a depth of 1mm.

Run gel

  1. Add 1-2μL RNA gel loading buffer to glyoxylated RNA samples
  2. Immediately load samples. Leave two outside lanes on each side empty.
  3. Load RNA size markers on outside lanes of gel.
  4. Electrophorese at 5V/cm.

Stain gel

  1. Prepare fresh 1:10,000 dilution in RNase free water of SYBR gold.
  2. Ensure pH is 7.0-8.5.
  3. Pour into staining tray.
  4. Place gel in plastic staining container.
  5. Shield from light.
  6. Agitate gently for 10-40 mins at room temperature.
  7. Image with Knight:Gel imager. (Place a clear ruler next to gel to more accurately assess length.)

Notes

References

  1. Molecular Cloning: Separation of RNA According to Size: Electrophoresis of Glyoxylated RNA through Agarose Gels

    [MolecularCloning]

Transforming chemically competent cells-PDF

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Transforming chemically competent cells-PDF

Method

  1. Thaw TSS cells on ice.
  2. Add DNA, and pipette gently to mix (1μl of prepped plasmid is more than enough).
    • Note: If you are adding small volumes (~1μl), be careful to mix the culture well. Diluting the plasmid back into a larger volume can also help.
  3. Let sit for 30 minutes on ice.
    • Note: If you are in a rush, you can shorten this incubation time to 5-10 min.
  4. Incubate cells for 30 seconds at [math]\displaystyle{ 42^o }[/math]C.
    • Note: According to the original TSS paper and qualitative experience (JM), this step is completely optional and may reduce transformation efficiency.
    • I tested this with DH5a Z1 and pUC19 and found that heat shock at 42C for 30 sec improved transformation efficiency 10-fold (Paul Jaschke)
  5. Incubate cells on ice for 2 min.
  6. Add 1 mL SOC (2XYT and LB are also suitable, original paper suggests LB + 20mM glucose) at room temp.
  7. Incubate for 1 hour at [math]\displaystyle{ 37^o }[/math]C on shaker.
    • Note: Can also save some time here by reducing incubation to ~45 min.
  8. Spread 100-300 μl onto a plate made with appropriate antibiotic.
  9. Grow overnight at 37 °C.
  10. Save the rest of the transformants in liquid culture at 4 °C. If nothing appears on your plate, you can spin this down, resuspend it in enough medium to spread on one plate, and plate it all. This way you will find even small numbers of transformants.
  • hugh kingston 09:46, 4 February 2008 (CST): I tried a few variations on this protocol, and found
  • using SOC instead of LB + 20mM glucose increased efficiency 3 fold
  • heat shock of 42°C 45s increased efficiency 15-20 fold compared to no heat shock

BioCoder version

Following is the Transforming chemically competent cells protocol in BioCoder, a high-level programming language for expressing biology protocols. What you see here is the auto-generated text output of the protocol that was coded up in BioCoder (see Source code). More information about BioCoder can be found on my home page. Feel free to mail me your comments/ suggestions. Vaishnavi

 

Miniprep/Qiagen kit-PDF

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Miniprep/Qiagen kit-PDF

Materials

For purifying plasmid DNA from Escherichia coli cells, the Qiagen Spin Miniprep Kit produces quite reliable results.

For Qiagen buffer compositions, please see the Qiagen Buffers page. Note: Qiagen buffer also works for Epoch Life Science’s spin columns which are sold in bulk at a much lower price. Yield and purity are quite close.

Protocol

See here or here for the handbook for the Qiagen Spin Miniprep Kit. If you have never done this protocol before, read the background information in the handbook (like the Important Notes section). It contains useful information. The following has been reproduced from the handbook and annotated based on experience with the kit.

Protocol: QIAprep Spin Miniprep Kit Using a Microcentrifuge

This protocol is designed for the purification of up to 20 μg of high-copy plasmid DNA from 1–5 ml overnight cultures of E. coli in LB (Luria-Bertani) medium. For purification of low-copy plasmids and cosmids, large plasmids (>10 kb), and DNA prepared using other methods, refer to the recommendations on page 37. Please read “Important Notes” on pages 19–21 before starting. Note: All protocol steps should be carried out at room temperature.

Procedure

  1. Resuspend pelleted bacterial cells in 250 µl Buffer P1 (kept at 4 °C) and transfer to a microcentrifuge tube.
    Ensure that RNase A has been added to Buffer P1. No cell clumps should be visible after resuspension of the pellet.
  2. Add 250 μl Buffer P2 and gently invert the tube 4–6 times to mix.
    Mix gently by inverting the tube. Do not vortex, as this will result in the shearing of genomic DNA. If necessary, continue inverting the tube until the solution becomes viscous and slightly clear. Do not allow the lysis reaction to proceed for more than 5 min.
  3. Add 350 μl Buffer N3 and invert the tube immediately but gently 4–6 times.
    To avoid localized precipitation, mix the solution gently but thoroughly, immediately after the addition of Buffer N3. The solution should become cloudy.
  4. Centrifuge for 10 minutes at 13,000 rpm (~17,900 x g) in a table-top microcentrifuge.
    A compact white pellet will form.
  5. Apply the supernatants from step 4 to the QIAprep spin column by decanting or pipetting.
  6. Centrifuge for 30–60 s. Discard the flow-through.
    Spinning for 60 seconds produces good results.
  7. (Optional): Wash the QIAprep spin column by adding 0.5 ml Buffer PB and centrifuging for 30–60 s. Discard the flow-through.
    This step is necessary to remove trace nuclease activity when using endA+ strains such as the JM series, HB101 and its derivatives, or any wild-type strain, which have high levels of nuclease activity or high carbohydrate content. Host strains such as XL-1 Blue and DH5α™ do not require this additional wash step.
    Although they call this step optional, it does not hurt your yield and you may think you are working with an endA- strain when in reality you are not. Again for this step, spinning for 60 seconds produces good results.
  8. Wash the QIAprep spin column by adding 0.75 ml Buffer PE and centrifuging for 30–60 s.
    Spinning for 60 seconds produces good results.
  9. Discard the flow-through, and centrifuge for an additional 1 min to remove residual wash buffer.
    IMPORTANT: Residual wash buffer will not be completely removed unless the flow-through is discarded before this additional centrifugation. Residual ethanol from Buffer PE may inhibit subsequent enzymatic reactions. They are right about this.
  10. Place the QIAprep column in a clean 1.5 ml microcentrifuge tube. To elute DNA, add 50 μl Buffer EB (10 mM Tris·Cl, pH 8.5) or water to the center of each QIAprep spin column, let stand for 1 min, and centrifuge for 1 min.
    If you are concerned about the concentration of the DNA, you can alternatively add 30 μL water to the center of the column, incubate at room temperature on the bench for 5 minutes, and then centrifuge for 1 minute. This will increase the concentration of DNA in your final sample which can be useful in some cases. See notes below for why you should elute in water rather than the Buffer EB they recommend if you plan to sequence your sample. Even if you are not sequencing, it may be beneficial to elute in water. For instance, if you elute in buffer EB and you are using this DNA in a restriction digest, then the additional salts in your sample can affect the salt content of your digest. This may matter with some finicky enzymes.

Notes

  • If you are doing more than ~10 mini preps simultaneously, it can save time to switch to the vacuum manifold version of this protocol since you eliminate having to load and unload samples into the centrifuge.
  • The sequencing center has begun using new machines and as a result, you may want to consider eluting in water rather than EB. See note from sequencing center.
    The elution is dependent on pH, however, measuring the pH of unbuffered water is difficult. However, anecdotally we have been able to get good yields using the water from the stock room. Eluting in deionized water from the Knight lab has also produced good results.
  • I use the “mini-fuge” for the binding and washing steps. You still have to do the drying step after the PE wash in a “real” microfuge though.
  • Passing the lysate over the column twice increases yield by about 20%.
  • Contaminating salt from the initial lysate or the PB will ruin a sequencing reaction more frequently than eluting in the EB (10 mM Tris as a small component of the total sequencing reaction is negligible). I always elute with EB and my reaction sequence is just dandy. There are two major sources of salt contamination: the inside upper edge of the spin column and the residual PB mixing with the PE wash. When you add the initial PE, it mixes with the leftover junk in the column. Spinning this through can only lower the salt to a level that was present after mixing. To get around these problems, I do two PE washes of about 300-500 μL. For the first, I dispense the liquid from the pipette tip along the inner ledge of the spin column in a circular motion to wash off the residue there. I follow the first PE wash with a second to further de-salt the sample before the drying spin. Yes, it adds a step, but the time spent here is far less than waiting three days only to find out your sequencing didn’t work.
  • Heating the elution buffer to 55°C before loading on the column can slightly increase yields.
  • Similarly, doing the elution in two steps (first a 30 μL elution and then a 20 μL dilution) can also slightly increase yields.

 

DNA ligation using NEB Quick Ligation Kit-PDF

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Materials

  • We use the Quick Ligation Kit from NEB
  • Deionized, sterile H2O
  • Purified, linearized vector (in H2O)
  • Purified, linearized insert (in H2O)

Ligation Mix

This is the same protocol that NEB recommends.

  • X μL vector (equivalent to 50 ng)
  • Y μL insert (3-fold molar excess, see below)
  • 10 μL 2X Ligase Buffer
  • (10 – X – Y) μL deionized H2O
  • 1 μL Quick Ligase

Calculating Insert Amount

[math]\displaystyle{ \rm{Insert\ Mass\ in\ ng} = 3\times\left[\frac{\rm{Insert\ Length\ in\ bp}}{\rm{Vector\ Length\ in\ bp}}\right]\times \rm{Vector\ Mass\ in\ ng} }[/math]

Procedure

  1. Add appropriate amount of deionized H2O to sterile 0.6 mL tube
  2. Add 10 μL ligation buffer to the tube.
    Vortex buffer before pipetting to ensure that it is well-mixed.
    Remember that the buffer contains ATP so repeated freeze, thaw cycles can degrade the ATP thereby decreasing the efficiency of ligation. It is recommended that you aliquot the Ligation Buffer into smaller quantities.
  3. Add appropriate amount of insert to the tube.
  4. Add appropriate amount of vector to the tube.
  5. Add 1 μL ligase.
    Vortex ligase before pipetting to ensure that it is well-mixed.
    Also, the ligase, like most enzymes, is in some percentage of glycerol which tends to stick to the sides of your tip. To ensure you add only 1 μL, just touch your tip to the surface of the liquid when pipetting.
  6. Incubate 5 mins on the benchtop.
  7. Place on ice until transformation.
  8. Generally 1 μL of ligation mix is sufficient for either chemical transformation or electroporation. The amount of salt in 1 μL ligation mix should not cause arcing.
  9. Optional Heat-inactivate by incubating at 65°C for 20 mins. Then do a purification step to remove PEG. (See notes on DNA ligation.

DNA Quantification-S2-PDF

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DNA Quantification-S2-PDF

Overview

This protocol uses a spectrophotometer to quantify the amount (μg/mL or ng/μL) of DNA and then uses a simple equation to convert this mass concentration into a molar concentration. The molar concentration is much more useful for most enzymatic processes.

  • Example: digesting 500ng of a 2KB plasmid is twice as much “enzymatic work” as digesting 500ng of a 4KB plasmid with the same multiple cloning site.

Procedure

1. Get DNA by any means necessary.
2. Run the DNA quantification (260/280) test on a spectrophotometer.

  • Be sure blank a sample first.
  • You can use only 1μL of sample if you use a NanoDrop
  • Or you can use 90μL of diluted sample using a UV cuvette
1. Dilute the DNA sample 30X by combining the following in a cuvette:

  • 87µl water
  • 3µl DNA prep
2. Run on the spectrophotometer with a dilution of 30.
3. Make sure that the A260 measurement is between 0.1 and 1.

  • If is too low then repeat the measurement using 15X dilution.
  • 84µl water
  • 6µl DNA prep

3. Calculate the molar concentration of DNA using the following equation:

  • Picomoles/µl = DNA Concentration(µg/ml) / [0.66*DNA Size(bp)]

4. Or calculate the μL of dna to add to obtain a desired molar amount of DNA.

  • μL = Picomoles*[0.66*DNA Size(bp)]/DNA Concentration(µg/ml)

Notes

  • One mole of single base pairs weighs 660 grams.
    • One picomole of 1000bp weighs 660ng.
  • 1ug = 1000ng
  • 0.001ug = 1ng

 

Agarose gel electrophoresis-S2-PDF

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Casting agarose gels

We precast our gels.

Materials

  • 1X TAE
  • SYBR safe (10,000X stock)
  • Agarose
  • Microwave
  • Stir plate and stir bar (optional)

Procedure

  1. Add 300mL 1X TAE to a 500 mL bottle.
  2. Measure out sufficient agarose to cast either a 1% (3 g) or 1.5% (4.5 g) gel.
  3. Add the agarose to the TAE buffer in the 500 mL bottle.
  4. Swirl to mix.
  5. Microwave bottle with loosened cap on high until the gel starts to bubble and is transparent.
    This generally takes just over two minutes for 300 mL. If you microwave too long, the gel will bubble over causing a big mess and you will need to start over.
  6. Remove from microwave and let cool by either sitting on bench top or adding stir bar and placing on stir plate.
    The advantage of the stir plate is that, if you forget about your gel for a while, it is less likely to solidify accidentally.
    If you are in a hurry, you can place the bottle in a beaker of room temperature water on the stir plate to speed the cooling process significantly.
  7. While gel is cooling, assemble casting trays and gel combs and verify that the trays are level.
  8. Once gel is cooled so that it can be touched comfortably with your gloved hand, add 30 μL SYBR Safe (10,000X concentrate).
  9. Pour gel into casting trays.
    The height of the gel will depend on how much you wish to load. Diagnostic gels can be reasonably shallow since typically 10 μL volumes are loaded. For gel purifications, the gel should be deeper to enable loading of large sample volumes.
  10. Let gels sit until they are solidified.
    Gels are solid when they are cloudy in appearance and firm to the touch.
  11. Gels may be used immediately. Alternatively, gels may be individually sealed in 6 x 10 inch polyethylene bags, labelled with initials, date and percentage and stored at 4 °C.
    It is a judgement call as to whether a gel is too old to be used. If it takes on a shrivelled appearance, don’t use it. If there is lots of condensation on the bag, only use it if your intended experiment isn’t critical.

Running agarose gels

Materials

  • Prepared DNA ladder
  • Precast gel with the appropriate percentage and well size/numbers for your samples (see above)
  • 1X TAE
  • Loading dye

Procedure

  1. Take a gel from the 4°C fridge.
    If the number of gels is getting low, cast more gels as described above.
  2. Place your gel in gel box.
  3. Add 1X TAE buffer to gel box such that buffer just covers the top of the gel.
  4. Remove comb.
  5. Load 12 μL prepared ladder
    Typically load ladder in left-most lane and sometimes right-most lane as well depending on whether you have the space.
  6. Use 2 μL loading dye per 10 μL of sample.
  7. Load samples left to right.
    The capacity of the 8 well, 1.5mm wide well is approximately 45 μL. The capacity of the 15 well, 1.5mm well is approximately 15 μL.
  8. Place gel box cover on gel box such that your samples will run towards the positive, red electrode.
  9. Run your gel at ~85 volts for 1 hr 20 mins. Use the timer to enable automatic shutoff of your gel.
    If you are in a hurry the gel can be run faster at ~95 volts for less than an hour.
  10. Verify that bubbles are rising from the electrodes once you start your gel to ensure your gel is running properly.

Visualizing agarose gels

Note that this procedure is under development

Materials

  • FluorChem 8800 gel imager

Procedure

  1. Remove gel from gel box shaking gently to allow residual buffer to fall back into gel box.
  2. Place in middle of UV box inside gel imager (you can leave the gel in gel tray).
  3. Make sure that filter wheel is set to position 4 (SYBR gold).
  4. Close the door and turn on reflective white light button.
  5. In gel imager software, click the “Acquire” button such that gel displays on screen.
  6. Adjust gel position on UV box so that the entire gel is within the frame.
  7. Close the door, turn off reflective white light and turn on transilluminating UV light.
  8. In gel imager software, click “Acquire image” button to capture gel image to the screen.
    It is occasionally necessary to adjust exposure time to improved image.
  9. Increase filtering if bands are difficult to see.
  10. Annotate gel as necessary.
  11. Save a copy of gel picture in your user folder.
  12. Print.
  13. Remove gel and throw in trash.
  14. Wipe down UV box if necessary.

Interpreting results

  • If you are getting unexpected bands on your gel you may want to look at the common agarose gel issues.

Notes

  • We have replaced Ethidium Bromide with SYBR Safe DNA gel stain. According to Reet Mand at Alpha Innotech Corp Technical Support: For SybrSafe dye, we recommend Fluorescein filter which is centered at 535nm. The part number is FLSC-500 and you can order it from us. Tom says this is the same as the SYBR-Gold filter installed in position 4 of the filter wheel.

Electrophoretic mobility shift assay-PDF

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Overview

An assay to check for protein-DNA binding.

Materials

Protein-DNA binding

  • See Knight:Protein DNA binding

Electrophoresis

  • Loading solution
    • Light sensitive
    • Stored at -20°C
    • Comes in EMSA kit from Invitrogen (manual, catalog number – E-33075)
    • Alternatively, Tom thinks we could get away with using our typical gel loading buffer.
    • Is this compatible with the DNA retardation gels gels?
  • Novex DNA Retardation Gels (manual, catalog number – EC6365BOX (10 well) or EC63652BOX (12 well))
  • 0.5X TBE running buffer
    • 44.5 mM Tris base
    • 44.5 mM Boric acid
    • 1 mM EDTA (free acid)
    • pH ? (do not use acid or base to adjust the pH of the solution)
    • Can make yourself or buy from Invitrogen (catalog number LC6675)
  • DNA ladder
    • Use standard 12μL of 2-log ladder with orange G dye? Seems to bleed a bit into acrylamide. Drop the loading volume down to 2 μL. Then the ladder will give an intensity more comparable to the 50ng probe but it still bleeds a bit.

Staining

  • SYBR Green EMSA nucleic acid gel stain
    • 10,000X concentrate in dimethylsulfoxide
    • Light sensitive
    • Stored at -20°C
    • Comes in EMSA kit from Invitrogen (manual, catalog number – E-33075)
    • Supposedly different from “normal” SYBR Green nucleic acid stain but not sure this is true.
    • “Normal” SYBR green nucleic acid stain seems to work just as well.
  • SYPRO Ruby EMSA protein gel stain
    • Light sensitive.
    • Store at room temperature.
    • Comes in EMSA kit from Invitrogen (manual, catalog number – E-33075)
    • Supposedly different from “normal” SYPRO Ruby protein stain but not sure this is true.
    • When mixed with TCA, it is stable for 6 months.
  • Trichloroacetic acid (TCA)

Procedure

Protein-DNA binding

See Knight:Protein DNA binding.

Add 2μL gel loading solution to each 10μL sample.

Electrophoresis

  1. Wear nitrile gloves.
  2. Prepare 1000mL of 0.5X TBE running buffer from 5X stock solution.
  3. Remove the NuPAGE gel from the pouch.
  4. Rinse the gel cassette with deionized water.
  5. Peel the tape from the bottom of the cassette.
  6. Gently pull the comb from the cassette in one smooth motion.
    • If you don’t do it smoothly, you can rip the wells.
  7. Rinse the sample wells with 0.5X TBE running buffer.
    • Use a pipetman and pipet to squirt in running buffer.
  8. Invert and shake to remove buffer.
  9. Repeat rinse two more times.
  10. Orient the two gels in the Mini-Cell such that the notched “well” side of the cassette faces inward towards the buffer core.
  11. Seat the gels on the bottom of the Mini-Cell and lock into place with the gel tension wedge.
    • Use the plastic buffer dam if you are only running one gel.
  12. Fill the upper buffer chamber with a small amount of 0.5X TBE running buffer to check tightness of seal.
    • If there is a leak, discard buffer, reseal chamber and try again.
  13. Fill upper buffer chamber. Buffer level should exceed level of the wells. Requires about 200mL
  14. Load 3μL 2-log DNA ladder.
  15. Load samples.
  16. Fill lower buffer chamber at the gap near locking mechanism with 600mL 0.5X TBE running buffer.
    • Should the buffer be chilled?
  17. Run at 100V for 90 minutes.
    • Gel showed some bowing at 100V when run for 65 mins. Drop the voltage?
    • When the Orange G dye front reaches the bottom, the 100bp DNA band is just over halfway down the gel.
  18. Shut off the power.

Staining the gel

  1. Before opening, warm the SYBR green EMSA gel stain concentrate to room temperature.
  2. Vortex and centrifuge tube.
  3. Dilute 5μL of 10,000X SYBR green EMSA gel stain concentrate into 50 mL 0.5X TBE buffer and pour into gel staining tray.
    • Exact amount depends on size of gel staining tray.
  4. Disconnect electrodes.
  5. Remove gels.
  6. Insert a knife in between the two plates and pry the plates apart.
    • You should hear a cracking noise as you break the bond between the two plates.
  7. Gently separate the two plates attempting to leave the gel on the bottom slotted plate.
  8. Cut to separate gel from bottom lip.
  9. Flip over and transfer gel to clean staining tray.
    • Use the lid of a 1000μL pipette tip box.
  10. Incubate ~20 mins on orbital shaker set at 50 rpm, protected from light.
    • Don’t use a glass container (glass adsorbs the dye).
    • Don’t reuse staining solution.
    • Staining time may vary with gel.
    • Store unused staining solution for 7 days in plastic container at 4°C
  11. Wash the gel in 150mL dH2O for ~10 secs.
  12. Repeat the wash step again.
  13. Wipe transilluminator with soft cloth and dH2O.
  14. Take a gel picture using 300nm tranilluminator. Set the filter wheel to SYBR green.
    • Use a piece of foil to help transfer the gel from the UV box back into the staining tray.
  15. When doing the protocol for the first time …
    1. Pour ~100mL of the SYPRO Ruby EMSA protein gel stain into the bottle of TCA.
    2. Wait ~5mins for the TCA to dissolve.
    3. Pour the TCA solution back into the bottle containing the rest of the SYPRO Ruby EMSA protein gel stain.
    4. Replace the cap securely.
    5. Mix by inverting at least 10 times.
    6. Check the box on the bottle indicating TCA
    7. Store at room temperature protected from light.
  16. Place the gel in a clean staining tray.
  17. Add 100mL SYPRO Ruby EMSA protein gel stain with TCA.
    • Exact amount depends on size of gel staining tray.
  18. Incubate ~3 hours on orbital shaker set at 50 rpm, protected from light.
    • Don’t use a glass container (glass adsorbs the dye). The lid of a 1000μL pipette tip box seems to be about the right size and work well.
    • You can leave the gel in stain overnight.
    • Do not dilute the stain.
    • Do not reuse the staining solution.
  19. Wash the gel in 150mL dH2O for ~10 secs.
  20. Repeat the wash step again.
  21. Destain the gel in 10% methanol, 7% acetic acid for 60 mins.
    • A gel stained overnight may need longer destaining.
  22. Wash the gel in 150mL dH2O for ~10 secs.
  23. Repeat the wash step again.
  24. Wipe transilluminator with soft cloth and dH2O.
  25. Take a gel picture using 300nm tranilluminator. Set the filter wheel to SYPRO Red.
  26. False color and superimpose images.

References

  1. EMSA kit from Invitrogen
  2. Jing D, Beechem JM, and Patton WF. The utility of a two-color fluorescence electrophoretic mobility shift assay procedure for the analysis of DNA replication complexes. Electrophoresis. 2004 Aug;25(15):2439-46. DOI:10.1002/elps.200405994 | PubMed ID:15300760 | HubMed [Jing-Electrophoresis-2004]
  3. Jing D, Agnew J, Patton WF, Hendrickson J, and Beechem JM. A sensitive two-color electrophoretic mobility shift assay for detecting both nucleic acids and protein in gels. Proteomics. 2003 Jul;3(7):1172-80. DOI:10.1002/pmic.200300438 | PubMed ID:12872218 | HubMed [Jing-Proteomics-2003]

All Medline abstracts: PubMed | HubMed

Notes

  • This kit is not sensitive enough for a complex mixture of protein and RNA. This kit has most optimum results when used with a more purified sample. The analysis of a complex solution with low concentrations of the actual target molecule requires more sensitivity than fluorescence can provide. From Molecular Probes technical assistance.
  • Promega has a useful FAQ.
  • Don’t bother running a protein standard. It doesn’t come out very well (either faint or blotchy depending on amount).
  • To date, the best staining tray I’ve found is the lid of a 1000μL pipette tip box. Then use a piece of mesh that just fits inside the lide to either keep the gel in place while changing solutions or to move the gel to and from the light box. This method requires smaller volumes of stain than the staining tray from Invitrogen.

Safety

  • Use nitrile gloves while handling acrylamide gels.
  • TCA is highly corrosive and hazardous. Use proper personal equipment protection.
  • SYBR green and SYPRO red stains must be disposed of as hazardous waste in separate waste streams. The SYPRO red is mixed with trichloroacetic acid and thus should be marked as ignitable and corrosive and the SYBR green is mixed with DMSO and should be marked toxic. (Kathy Gilbert, EHS)

Cloning Protocol-PDF

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PCR

There are two types of PCR polymerases used in the lab, TAQ and Phusion. Phusion is more accurate, having fewer mutations or errors in PCR. TAQ will almost always give a good product and is used when accuracy is not an issue. TAQ has an error rate of about 1 bp change per Kb. The protocols for these are very similar

Phusion

50 ul reaction

1. ddH20 – 35.5 ul

2. 5X Phusion buffer— 10 ul

3. 10 mM dNTP’s— 1 ul

4. Primers (50 uM stock)—1 ul each for 2 ul total

5. Template DNA—1 ul

6. Phusion Polymerase— .5ul

====a.==== The polymerase is VERY expensive so make sure that you only get .5 uL per reaction.

====b.==== Make sure you just get out the polymerase after the other stuff is already mixed in to ensure the proper pH and to make sure that the polymerase stays cold. As with all enzymes, ALWAYS use the blue freezer boxes to keep it cold at your bench.

TAQ

50 ul reaction

1. ddH20— 41 ul 2. Standard 10X reaction Buffer— 5ul 3. 10 mM dNTP—1 ul 4. Primers—1ul each for 2 ul total (1ul forward and 1ul reverse) 5. DNA template—1ul of liquid or 1 colony 6. 1x TAQ DNA polymerase—1 ul

====d.==== After completing the reaction mix you will need to place it into the PCR machine. Sign into your profile and then select the program that correlates to the PCR reaction you will be running. Press start and then select tubes, or plates. It will usually take anywhere from 2-3 hours for shorter products. For the extension time, TAQ requires 1 min per kb of product and Phusion requires 2 min per kb of product. RECORD the PCR conditions, the primers, template and program used in your notebook.

DNA Electrophoresis

After completing the PCR you will need to run it out on gel. The purpose is to determine if the product is correct by visualizing the size of your PCR product (it should be the size you wanted for your gene of interest).

a. To make the gel you will need 75 ml of TAE and .75 grams of agarose (regular agarose, not low melt).

b. Put it into the microwave for about 90 seconds or until the agarose is completely dissolved.

c. Add 7.5 ul of ethidium bromide, make sure you wear gloves because it is a carcinogen.

d. After it is has cooled briefly, pour the liquid onto the gel bed and let it cool. It should only take about 30 minutes for it to cool.

e. You will need to have a ladder in one lane and then place your PCR products in the others. You will need a ladder in each row to be able to determine the size of your PCR products. We have 6 ul ladder which means you will just need 6 ul of the ladder. We have 10X dye which you will add to the PCR product to make it heavy enough to lad into the gel (the dye contains glycerol) . We typically run 10 uL of PCR reaction to confirm there is a product . Do this by adding 5 uL of 10X dye to 50 ul of PCR , then load only 10 uL of this mix.

f. Move the gel into the proper orientation in the gel box, cover your gel with 1X TAE buffer and load all your products into the gel.

g. Put the lid on and make sure that the negatives and positives are in the correct places (Red to red, black to black. DNA is negatively charged so the DNA will run toward the red probe) . Turn on the power supply and to run around 150-175 volts. It will take about 30-45 minutes to complete.

h. Turn it off and then the check it on the alphaimager. Print off the results and log in your notebook.

PCR Qiagen Purification Kit

a. This step will help to purify the DNA product away form the buffer and enzymes present in the PCR mix. You can follow the procedure in the Qiagen manual.

Restriction Digest

50 ul reaction

The purpose of this restriction digest is to cut the plasmid open and so that the gene of interest can be inserted into the place where the plasmid will be cut. You will need to have the proper restriction enzymes picked out so that they cut the plasmid in the proper place. There will be two different reactions. One will be with the Purified PCR product and the other will be the plasmid that the gene will be inserted into.

Purified PCR reaction

1. 5 ul NEB buffer (it depends on the R.E. used) 2. .5 ul 100X BSA 3. 40 ul PCR product 4. 1.5 ul Rectricion enzymes (total of 5 ul, 2.5 of each one)

Plasmid

1. 30 ul ddH20 2. 5 ul 10X NEB buffer 3. .5 ul 100X BSA 4. 10 ul of plasmid (3ug) 5. 1.5 ul Restriction enzymes (total of 3 ul, 1.5 ul of each one) ==b.== After all of these are mixed together they must be placed in the 37º bath for at least two hours or overnight.

Low-melt agarose gel

==a.== It is basically the same as the other gel except for 3 things, there is low-melt agarose gel powder that must be used, and it will only need to be in the microwave about 50 seconds or until completely dissolved. Second, it polymerizes slowly so put it in the fridge to solidify. Third, when running the gel, run at 70-90 Volts (do not go over this voltage or else it will melt). It will take about 1 to 1.5 hours to run.

==b.== After it is done running you will use the portable UV lamp and you will extract the products right out of the gel (get help if you do not have a lot of practice at this). Label each tube and place the products into a 1.5 ml eppendorf tube. Be sure to cut out as small of a slice as possible to avoid agarose which inhibits the ligation. It is okay to cut away some of the DNA, you just want to get the DNA to be as concentrated as possible. Also cut out a small slice from an area of the gel that has no DNA as a control for the insert.

Ligation

a.

The purpose of this is to combine the digested plasmid with the gene of interest and then ligate the gene of interest into the plasmid. You will need to set up at least two reations. The plasmid only with insert control and the plasmid with your gene of insert as the insert. To be able to use the DNA you will need to melt the gel at 65 degrees Celsius for about 5 minutes. While it is melting, make a master mix that has everything but the agarose slices and distribute into eppendorf tubes. Add the agarose slices quickly after they melt (before they resolidify). NOTE: you will need a tube for each insert, plus one for the plasmid only control.

Plasmid only

1. 3 ul of plasmid slice

2. 3 ul of insert control slice

3. 6.5 ul of ddH20

4. 1.5 ul 10X ligase buffer

5. 1 ul T4 DNA ligase

Plasmid plus insert

1. 3ul of Vector slice

2. 3ul of insert

3. 6.5 ul of ddH20

4. 1.5 ul 10X ligase buffer

5. 1ul T4 DNA ligase

b.

These reactions will need to incubate at room temperature for at least an hour.

Transformation

a.

Now, if all has gone well, the gene of interest will be in the plasmid and all that will be left to do is to put the plasmid into E. coli (or use other competent cells). The E. coli we use is DH5alpha.

1. Get a cooler and put ice into it (Make sure you have 42 degree water bath set up for a later step). You will also need to get the plates out with the proper selection marker.

2. Get the DH5alpha cells out of the -80 degree Celsius freezer and put them directly into the ice. You will need about 25-30 ul for every reaction you have.

3. They will need to thaw on the ice for a few minutes.

4. After they have cooled you will need to put 2 ul of the ligation mix and add it to about 25-30 ul of competent cells. Vortex them and put them back in the ice for 2-30 minutes

5. Heat shock them by putting them in the 42 degree water bath for 1 minute.

6. Put them back on ice for 2-5 minutes

7. Add .5 ml of LB to the competent cells for 30 minutes.

8. Put all of the mixture into onto the plates and then use the large beads to put the cells all across the plate.

9. Incubate them at 37 degrees Celsius

Checking the clones

a.

After the 24 hour incubation period, if the cloning worked then you should see individual colonies that hopefully contain the plasmid with the gene of interest in the plasmid. This is what you will need to check. (it is important that you do not incubate them over 24 hours because doing this creates “satellite colonies” which the surrounding cells have used up all of the Amp and thus do not have the plasmid in them but can survive because there is no Amp in that part of the plate).

b.

There are two ways to check your clones one is by doing a dirty plasmid prep and the other is to check it using PCR

Checking with PCR

Checking with PCR (you will need to have LB+selction plates out to plate each colony that you are testing. If one of them has the gene of insert in it then you will need to have something to go back to use it more.

1.

TAQ (on the PCR matching you will need to add a 95 degree Celsius for two minutes at the beginning, this will break open the cells)

=====a.===== For a 20 ul reaction

i. ddH20— 16.5 uL

ii. Standard 10X reaction buffer—2uL

iii. Primers—.5 uL each for for a total of 1 uL( .5ul Forward and .5 uL reverse)

iv. DNA template—1 colony (use colony from the transformation plate)

v. 1X TAQ DNA polymerase—.5 uL

Checking using a dirty plasmid Prep

1. After the transformation is done you will need to streak for single colonies. To do this you will need to get LB+ selection plates out. You will pick one and only colony off the transformation plate and then the plate and streak them onto the new plate. Incubate them for 24 hours at 37 degrees Celsius.

2. Now you will need to make overnights of a single colony off the plates you just streaked the day before. You will need to get test tubes out and put about 2mL of LB+selction in them. Label your tubes.

3. Once you have the tubes you ready, you will take a pipet tip in your hand and you will take one individual colony off and drop it into the test tube. For a control leave a test tube with just LB+selction (no colony). After all of the samples are ready you put them into the 37 degree shaker and place the plates into the 37 degree incubator for 24 hours.

4. After 24 hours you will take the test tubes out of 37 degree shaker. Get 1.5 mL eppendorf tubes out for each sample. You will pipet out about 1-1.5 mL from the test tube and put it into the eppendorf tube.

5. Spin down the eppendorf tubes at 13,000 rpm for 2 minutes. After the spin is done decant all of the LB out. It will be on the top. The cells will be on the bottom. You just want the pellet (cells).

6. Place them in the freezer for about an hour or more. This will help them to be able to break open more easily.

7. Remove them from the freezer and then make the tens solution. (Make sure that you use 10M NaOH to dilute it down to 1M NaOH. )

8. Here is the mix that you will need to make in order to make the plasmid prep per sample.

     a.	300 uL  TENS
     b.	3 uL NaOH 
     c.	.6 uL RNAaseA 
             i.	for example if you have 10 reactions to do then you will need to mix
                     1.	3000 uL of Tens
                     2.	30 uL of NaOH
                     3.	 6  uL of RNAaseA

9. Put 300 uL of TENS into each eppendorf tube with you pellet. You want your pellet mixed in well with TENS so it is easiest if you pipet up and down to mix it all together.

10. Pipet 150 uL of NaAc (sodium acetate) pH. 5.3 into each eppendorf tube. NO NOT VORTEX it will shatter the plasmid. Invert the tube up and down 4-6 times to mix it all together.

11. Place in centrifuge, 13,000 rpm for about 4 minutes.

12. The plasmid will not be in the supernatant, you will want to get new eppendorf tubes and pipet out the supernatant and put it into the new tubes. Throw away the debris.

13. Put 900 uL of cold 95% ETOH into each tube.

14. Centrifuge them for 2 minutes at 13,000 rpm.

15. Decant the ETOH

16. Add 1 mL of 70% ETOH and put them back in the centrifuge with the same settings as before.

17. Decant ETOH and repeat step 16.

18. After the second spin make sure you decant all of the ETOH. Let the eppendorf tubes air dry for about 5 minutes or until all the ETOH has evaporated.

19. You should be left with a white pellet.

20. Add 50 uL of EB buffer from the qiagen kit.

Restriction Digest

1. Mix plasmid in with the EB buffer.

2. Put 10 uL of plasmid into eppendorf tubes.

3. Make a master mix with your restrction enzymes. You will only use a 10 uL of mix per tube.

     a.	8  uL of ddH20
     b.	2 uL of  NEB buffer
     c.	.2 BSA
     d.	.2 Restrction enzyme A
     e.	.2 Restriction enzyme B

4. Put the mix of 10 uL of restriction enzyme mix, with 10uL of plasmid into the 37degree water bath for at least 1 hour.

5. Make a gel and run the 20 uL reaction on the gel. You should be able to see the size of the plasmid along with the insert.

6. If the insert looks like it is the right size. Follow the next protocol.

Sequencing gene of interest

1. Purify your plasmid by making a 5 mL overnight. Take a colony from the plate and place it in the test tube.

2. Spin the cells down and use the qiagen kit to purify your plasmid.

3. Dr. Grose will need to place an order for you on the DNA sequencing center website.

4. The mix you will need to make is

     a.	8 uL of ddH20 
     b.	2 uL of (1/20 dilution of primers) Talk to Dr. Grose about which primers you need to use.  They are not the same as the ones that you did your orginal PCR product with.

5. Take to room 690 in the widsoe for sequencing. Ask someone who has gone before for the exact details of what to do there.

Blunting After Restriction Digest

(Preheat a block incubator to 75C first) a. After the restriction digest perform a Qiagen PCR clean-up kit to each sample. b. Next make a 1:10 dilution of 10mM dNTPs c. Then to each sample add the following to your DNA/plasmids: 1. 1ul of 1:10 dilution of 10mM dNTPs 2. 10ul of 5X Phusion Buffer 3. 3ul of Phusion Polymerase d. Incubate the samples in a 37C water-bath for exactly 10 minutes, DO NOT GO

   LONGER!!!

e. Immediately place the samples in the 75C block incubator for 10 minutes (this

   inactivates Phusion Polymerase)

f. Perform another Qiagen PCR clean-up kit to each sample g. Perfrom CIP (Phosphotase) treatment to the plasmids only (do not do this to the

   DNA insert) to keep them from re-ligating together. This is accomplished by
   doing the following:

1. Add 5ul of NEBuffer 3 to each plasmid sample 2. Add 1ul of CIPhosphotase 3. Incubate all samples in 37C water-bath for 1 hour 4. Immediately place sample in 75C block incubator for 10 minutes (this inactivates CIP) h. Finally run your samples on low-melt agarose gel and isolate your fragments of

   interest and perform ligation as described in above methods.

QRT-PCR-PDF

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QRT-PCR-PDF

Overview

Quantitative reverse transcriptase PCR (QRT-PCR or qRT-PCR) is a PCR technique used to determine the amount of cDNA in a sample. It is the most commonly used form of quantitative PCR (qPCR). This technique is also called real-time reverse transcriptase PCR.

Comparison of normalization methods

There is an ongoing debate what is the best way to normalize qPCR data. Reference genes are the most common method, although single unverified reference genes invalidate the qPCR data generated. Total RNA, ribosomal RNA, and genomic DNA have been suggested as alternative methods.

Reference genes

The most common method. Frequently, a panel is used for normalization, e.g. [1] not just a single reference gene and including data on suitability as reference genes. Often housekeeping gene is used here instead of reference gene but the term is poorly defined and can be misleading. It is to be noted that panels are often composed of genes that are supposed to be stable based on their function. However, more than 100 peer-reviewed articles report problems related to genes chosen from a panel, because they were not suitable for a particular context. A recent approach is to select a reference gene based on its stability across microarrays done within one’s condition of interest. There is a public tool called RefGenes that searches a microarray database of more than 50,000 arrays to identify genes that are stable across subsets of conditions. It is available at the Genevestigator website [2].

RNA

Total rRNA [3] [4], or total RNA. Drawback: rapidly dividing cells will have more rRNA and different rRNA/mRNA ratio which will complicate comparison; difference in cDNA synthesis not taken into account.

Genomic DNA

Genomic DNA or cell number. Drawbacks: RNA degrades faster than RNA which can distort the data; sample cannot be DNase treated; efficiency of cDNA synthesis not taken into account.

Reference mRNAs

Main article: Choosing reference genes for qPCR normalisation

Picking reference genes will make or break your quantification via qPCR (real time PCR). If you pick only one reference gene and your pick is not constant across different conditions or samples, your results will be skewed. Choose several reference genes and check whether they satisfy the criteria for a good reference gene. Some commonly used reference genes, like 18S and GAPDH, are known to be problematic but continue to be used.

  • Ajeffs 06:55, 21 April 2007 (EDT): Screen a handful of ref genes, select the most stable using genorm, bestkeeper etc, use at least 2 reference genes for subsequent reactions and normalisation. Inlcude your genorm M values when publishing qPCR data.

Primer selection

Main article: Choosing primers for qPCR

Choosing suitable primers is an early crucial step in your qPCR experiment. Reusing a tested primer pair from a repository or publication can save you some time. Otherwise primer selection from scratch is similar to that for a standard qualitative PCR experiment but the product size is typically much smaller (below 200nt) and the amplification characteristics of the primer have to be rigorously tested.

Quantification methods

There are 3 common quantification methods. The standard curve method is the only one that gives you are absolute concentration. Both the Pfaffl method and the ΔΔCt method produce relative data with the Pfaffl method being superior.

Standard curve method

  • requires template at known concentration (e.g. cDNA or TA cloned PCR product)
  • requires dilution series of known template for standard curve (more wells)
  • yields absolute concentrations by comparing unknown samples to known

Pfaffl method

  • requires that primer efficiency be known but needs to be determined only once with a standard curve or a different method
  • produces relative amount (e.g. treated is 2x untreated)

(named after the inventor; see Pfaffl 2001 PMID 11328886)

ΔΔCt (delta delta Ct)

  • easiest, oldest, least reliable
  • assumes that primers for unknown and reference gene have very similar efficiency
or that v little correction is necessary (i.e. reference gene almost same level)
  • yields relative amounts

(Ct = cycle threshold; point when fluorescence reading surpasses a set baseline)

Primer efficiency estimation

A Ct difference of 1 between two samples has a different meaning depending on the efficiency of the primers used. If primers are 100% efficient, then ΔCt = 1 means one sample has twice the amount of template compared to the other. The simple ΔΔCt method, described above, often wrongly assumes perfect efficiency. It is better to experimentally verify the primer efficiency and use the Pfaffl method instead. The standard method takes the primer efficiency into account via the standard curve run with each sample. However, primer efficiencies in the standard curve dilutions and the actual samples are not necessarily the same.

Linear regression on dilution curve Ct data

Primer efficiencies can be calculated by making a dilution series, calculating a linear regression based on the data points, and inferring the efficiency from the slope of the line. For a base 10 logarithm the formulae is:

efficiency = 10^(-1/slope)

Slopes between -3.3 and -4 will thus give you estimated primer efficiencies between 100% and 78% respectively. It can happen that the calculated efficiencies are above 100% [5] [6] [7]. This may be due to incorrect template concentrations, too concentrated template, inhibition of the PCR reaction, unspecific PCR amplification, mistakes in the calculation [8], etc.

See a figure explaining the fitting process from the Hunts’ qPCR tutorial [9].

Efficiency estimation based on the kinetics of single PCR runs

Efficiency (and Ct values) can also be calculated from the fluorescence data of a single PCR run or preferably replicates of the same PCR. The Miner algorithm (PMID 16241897) is an example for this type of method and can be used online at [10].

qPCR data quality

Sources of variability: Operator

Due to the small amount of liquid handled and the sensitivity of the technique, operator variability is high. Bustin reports that the same qPCR experiment repeated by 3 people using the same reagents lead to very different copy number estimations [Bustin 2002 PMID 12200227, figure 3]:

  • person A: 8·7 × 105
  • person B: 2·8 × 105 different by a factor of 3!
  • person C: 2·7 × 103 different by a factor of 300!!

Sources of variability: Reagent lots/age

Different lots of reagents can lead to different results. Experiment repeated by same operator 5 times, same RNA sample, different kits; values are copies/μg total RNA:

  • kit 1: 13±32 × 107
  • kit 2: 5.4±1.6 × 107 – different by a factor of 2.4

Similar experiment with old (9 months 4°C) and new probe (3 months 4°C), values are copies/μg total RNA:

  • old: (5.6 ± 1.3) x 103
  • new: (3.8 ± 0.6) x 108 – different by a factor of 100’000!!

both experiments above from [Bustin 2002 PMID 12200227, figure 4]

Notes

  • The most commonly used specialist reverse transcriptase enzyme for cDNA production is AMV reverse transcriptase. It has RNase H activity (so that RNA molecules are only transcribed once) and has a high temperature stability (to reduce RNA secondary structure and nonspecific primer annealing) [1].
  • Since RNA can degrade with repeated freeze-thaw steps, experimental variability is often seen during successive reverse transcription reactions of the same RNA sample [1].
  • Reverse transcriptase enzymes are notorious for their thermal instability. Repeated removals from the freezer can degrade the efficiency of the enzyme [1].
  • Producing total cDNA from total RNA can be advantageous because
    1. cDNA is more stable than RNA so making total cDNA allows you to make multiple sequence-specific RNA measurements [1].
    2. This approach could reduce experimental variability stemming from RNA degradation [1].
  • To make total cDNA
    1. Use a polyT primer (most but not all eukaryotic mRNA) or random decamers (prokaryotic mRNA) [1].
    2. Random decamers give longer cDNAs on average than random hexamer primers [1].
    3. Use longer reverse transcription reaction times [1].
    4. Ensure that the concentration of deoxynucleotides doesn’t run out [1].
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