Endy:Northern blot, AlkPhos end-labeled probes
The following protocol for Northern Blotting is based on getting RNA from the RNA extraction method used by Sean Moore, Agarose (as opposed to polyacrylamide) gel electrophoresis, and the Turboblotter system from Whatman. This is largely a compilation of another previous Endy lab protocol and the Knight lab's protocol for RNA gels.
- 1 Materials
- 2 Protocol
- 2.1 Running the Gel (2-6hrs, depending on size of gel)
- 2.2 Stain gel (can't do if continuing to northern)
- 2.3 Prepare the Membrane (5-15 minutes)
- 2.4 Transfer of RNA onto Membrane by Turboblotter Capillary Transfer (3-4 hours)
- 2.5 RNA Fixation with UV Cross-Linking (5 minutes)
- 2.6 Hybridization (2 hours + overnight + >1 hour)
- RNA extracted from cells (should be ~1-5µg/µl).
- SYBR Gold for staining the gel (if you're going to view it under UV).
- HPLC grade or better DMSO
RNAse free water
- make lots for rinsing glassware and electrophoresis chambers (4L for a reaction?)
- Add DEPC to final concentration of 0.1%.
- Incubate 1hr at 37°C.
- Autoclave for 15 mins at 15 psi.
This kit uses downward capillary transfer and therefore can run a transfer in <4hrs. It uses (+) charge nylon membranes, which bind RNA more tightly but also have higher background.
- From Whatman []
- Stack and Buffer trays. These can apparently be improvised using empty tip boxes (?).
- Blotting paper.
- (+)nylon membrane (Nytran SuPerCharge)
- buffer wick
10X BPTE electrophoresis buffer
This buffer has low ionic strength and does not remove glyoxal from RNA during electrophoresis and therefore does not have to be recirculated.
- The final pH of this 10x buffer is ~6.5.
- 100 mM PIPES
- 300 mM Bis-Tris
- 10 mM EDTA
- Prepare by adding the following to 90 ml of distilled H2O. (might scale up to 500ml for first time).
- 3 g of PIPES (free acid)
- 6 g of Bis-Tris (free base)
- 2 ml of 0.5 M EDTA
- Treat the solution with final concentration of 0.1% DEPC for 1 hour at 37°C
- Commercially available stock solutions of glyoxal contain both hydrated forms of glyoxal and oxidation products that can degrade RNA. These must be removed; we used the protocol as follows (See Sambrook and Russel 3rd Ed, App. 1-24):
- Run the Glyoxal over mixed-bed ion exchange resin. We used BioRad AG 501-X8 (D) resin beads (these beads need to be largely blue; if they're gold-orange, they've oxidized and should be disposed of as HazWaste. Our resin was a mix of blue and gold beads, so I ended up just using as much resin as I needed to to get the pH up. This seemed to work fine. ~~Felix Moser. Equal volumes of resin and glyoxal were placed in an epp tube; the glyoxal's pH was measured w/ pH paper strips after a few minutes. Repeat this until the glyoxal's pH is >5.5.
Glyoxal reaction mixture
- 6mL DMSO
- 2mL deionized glyoxal
- 1.2mL of 10X BPTE electrophoresis buffer
- 0.6mL of 80% glycerol
- 0.2ml of either ddH2O or ethidium bromide, depending on how or whether you're staining it.
- (divide into small aliquots and store at -70°C)
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
- got some from Sean Moore. Make sure final volume of buffer is no less than 50% volume of the sample.
- 1.5M NaCl
- For 1L: 20g NaOH, 87.66g NaCl, bring volume to 1L w/ ddH2O.
- 0.5M Tris-HCl pH 7
- 1.5M NaCl
- For 1L: 60.56g Tris, 87.66g NaCl, 800ml ddH2O, adjust pH w/ conc. HCl and bring volume to 1L w/ ddH2O.
20x SSC "Transfer" buffer
- 3M NaCl
- 0.3M sodium citrate
- adjust pH to 7.
- For 1L: 175.5g NaCl, 88.2g Na citrate (dihydrate), 800ml ddH2O, adjust pH to 7 then bring to 1L w/ ddH2O.
Running the Gel (2-6hrs, depending on size of gel)
For RNase free electrophoresis apparatus: Clean electrophoresis tanks and combs used for electrophoresis of RNA with detergent solution, rinse in H20, dry with ethanol, and then fill with a solution of 3% H2O2. After 10 minutes at room temperature, rinse the electrophoresis tanks and combs thoroughly with H2O treated with 0.1% DEPC
This protocol comes pretty much straight from Sambrook and Russel 3rd ed.:7-29.
- Set up the glyoxal denaturation reaction by combining 0.5-2 ul of RNA (up to 10 ug) with 10 ul of glyoxal reaction mixture. You MUST also treat your RNA ladder the EXACT SAME WAY so it runs the same as your samples if you want to use it for comparison.
- Incubate the RNA solutions for 60 minutes at 55°C. Chill the samples for 10 minutes in ice water and then centrifuge them for 5 seconds to deposit all of the fluid in the bottom of the microfuge tubes.
- While the samples are incubating, clean electrophoresis tank if necessary, and pour a 1.5% agarose gel in 1X BPTE (1.05 g agarose in 70 mL buffer). When set, cover the gel with sufficient buffer.
- Add the formamide loading buffer (>equal volume buffer) to the glyoxylated RNA samples, and without delay, load the glyoxylated RNA samples into the wells of the gel.
- Carry out electrophoresis at 70 Volts (in 14cm gels); ie. run gel @ 5V/cm.
- Trim away areas of the gel to be stained with sybrGold. DO NOT trim the area BELOW the loading dye, since many RNA's run much lower than the dyes; trim the gel later once you know where all your RNA is. (Membrane should be cut to match the size of the gel.) Wrap gel to be stained in sarran wrap and store at 4°C until post-transfer gel is ready to be stained.
- Soak gel 30 min in "Denaturing Buffer" (0.5 M NaOH/1.5 M NaCl (~400 mL)). Agitate occasionally.
- Rinse the gel in ddH2O and Soak gel 20 min in "Neutralizing Buffer" (0.5 M Tris (pH 7.4)/ 1.5 M NaCl (~400 mL))
7 and 8 are optional steps for improving the transfer of long RNAs, esp from >1% gels. These steps are the from the Turboblotter protocol
Stain gel (can't do if continuing to northern)
- Prepare fresh 1:10,000 dilution in RNase free water of SybrGold or SybrGreen (SybrGreen recommended by Sean, since it's specific for RNA; ppbly doesn't make a big difference, though).
- Ensure pH is 7.0-8.5.
- Pour into staining tray.
- Place gel in plastic staining container.
- Shield from light.
- Agitate gently for 10-40 mins at room temperature.
- Image with gel box. (Place a clear ruler next to gel to more accurately assess length.)
Prepare the Membrane (5-15 minutes)
NOTE: Be gentle with the membrane. The number of times a membrane can be stripped and re-probed is usually limited by physical damage to the blot.
- Cut a piece of membrane to the dimensions of the agarose gel. Max dimensions for hybridization in 50mL tubes: 8 x 9 cm (circumference x diameter). If membrane is larger, sandwich between sheets of nylon mesh to allow buffer to penetrate overlap.
- Wet the membrane by carefully laying it on top of Milli-Q water in a shallow tray. (Do not immerse the Immobilon-Ny+ membrane in liquid on the first liquid exposure. If you wet both sides, air can become trapped in the pores and form bubbles.)
- Agitate the tray gently once the membrane is wet to completely immerse the membrane.
- Transfer the membrane to a second tray containing transfer buffer (20 x SSC).
- Equilibrate the membrane at least 5 minutes.
Transfer of RNA onto Membrane by Turboblotter Capillary Transfer (3-4 hours)
NOTE: Refer to Fig. 1 when setting up the TurboBlotter System.
- Place stack tray of transfer device on bench, making sure it is level.
- Place 20 sheets of dry GB004 blotting paper (thick) in stack tray.
- Place 4 sheets of dry GB002 blotting paper (thin) on top of stack.
- Place one sheet of GB002 blotting paper, prewet in 20xSSC transfer buffer on stack.
- Place transfer membrane on stack. Smooth bubbles by rolling a clean glass pipette over the surface. Do not touch with gloves.
- Cover the membrane with agarose gel, cut the gel to the size of the membrane, making sure there are no air bubbles between the gel and the membrane.
- IF the gel is smaller than the membrane and the surrounding blotting paper, cover the area surrounding the gel w/ PLASTIC (eg. Sarran wrap, cut-out parts of tip boxes) so that the transfer buffer does not soak through the areas where the blotting paper above the gel touches the blotting paper below it.
- WET the top surface of the gel with transfer buffer and place 3 sheets of presoaked (in 20x transfer buffer) GB002 Blotting Paper, in on top of the gel.
- Attach the buffer tray of the transfer device to the bottom tray using the circular alignment buttons to align both trays.
- Fill the buffer tray with 125 ml transfer buffer for 7 x 8 cm to 11 x 14 cm transfers; (200 ml for 12 x 21 cm to 20 x 25 cm transfers).
- Start the transfer by connecting the gel stack with the buffer tray using the precut, PRESOAKED buffer WICK (included in each blotter stack), in transfer buffer. Place the wick cover on top of the stack to prevent evaporation. Make sure the edges of the wick are immersed in the transfer buffer.
- Continue the transfer for 3 hr. Additional transfer time may be required for gels thicker than 4 mm or larger-size nucleic acids. (Try 4 hr, since gel is 1.5%)
- Disassembly: mark edges of gel and lane borders onto blot with pencil.
NOTE: Do not place any other weight on top of the wick cover during transfer. This is unnecessary and may inhibit transfer by crushing the pore structure of the agarose gel.
RNA Fixation with UV Cross-Linking (5 minutes)
- It is not necessary to allow the blot to dry completely prior to UV cross-linking.
- Place the blot on a sheet of clean filter paper to prevent contamination if you plan to place the UV light source above the blotted RNA. (If you plan to place the membrane on a UV transilluminator, clean the surface with Milli-Q water and a Kimwipe.)
- Expose the side of the blot with the bound RNA to a UV light source (254 nm). We used one of the Sauer lab's UV handheld light sources for 5min.
Hybridization (2 hours + overnight + >1 hour)
This part highly depends on the type of labeling one is doing; protocols vary widely in buffers used, types of probe preparation, and hybridization protocol. We used the GE Healthcare AlkaPhos kit; the protocol below is essentially copy-pasted from the kit spec sheet.
Need the following solutions:
- Hybridization Buffer
- add to the hybridization buffer solution: NaCl to 0.5M, blocking reagent to 4% (add agent slowly while continuously stirring... takes a while to dissolve). Finished buffer can be stored @ -20*C in aliquots.
- Primary Wash Buffer
- 2M Urea, 0.1% SDS, 50mM Na phosphate pH7, 150mM NaCl, 1mM MgCl2, 0.2% blocking reagent (kit specific);
- Recipe for 1L: 120g Urea, 1g SDS, 100ml of 0.5M Na phosphate pH=7, 8.7gg NaCl, 1ml of 1M MgCl2, 2g blocking reagent.
- Can be stored 1wk @ 4*C.
- 20x Secondary Wash Buffer (Stock)
- 1M Tris base, 2M NaCl; Can be stored at 4*C for 4mo.
- Recipe for 1L: 121g Tris base, 112g NaCl.
- For working dilution: dilute 1:20 and add 2ml/L of 1M MgCl2 t final conc of 2mM Mg in the bufer. This buffer should not be stored.
Hybridization and Wash:
- If blot dimensions are less than 8 x 9 cm, place blot in a 50 mL Falcon tube, RNA facing in. Make sure the blot doesn't overlap itself. 50 mL conical tubes fit in the Sauer Lab "Bambino" mini-hybridization oven. If blot is too large for a 50 mL Falcon tube, use large glass tubes and Baker Lab hybridization oven.
- Pre-heat the rquired volume of prepared hybridization buffer to 55*C. The volume of buffer should be equivalent to 0.25ml/cm2 of membrane; this may be reduced to half that volume for large blots hybridized in plastic bags or in bottles. (For our 9x9cm blot in the 50ml Falcon tube, we used 10ml of hybridization buffer).
- Place the blot into the hybridization buffer and prehybridize for @ lest 15 min @ 55*C in a shaking water bath or hybridization oven. We used the Sauer Lab's "Bambino" hybridization oven.
- Add the lableled probe to the buffer used for the pre-hybridization step. Typically use 5-10ng/ml. Avoid placing the probe directly on the blot. Try taking out a small aliquot of buffer and mixing that with the probe before returning the mixture to the bulk of the hybridization buffer.
- Hybridize @ 55*C overnight. Stringency can be adjusted by altering the hybridization temp between 50*C and 75*C.
- Preheat the primary wash buffer to 55*C.
- Transfer the blot to the an excess of primary wash buffer and wash for 10min @ 55*C, with gentle agitation.
- Repeat wash with primary wash buffer for 10min @55*C.
- Transfer blot to new container and wash with secondary wash buffer for 5 min @ room temp.
- Repeat wash with secondary wash buffer for 5min @ room temp.
- Drain excess wash buffer from the blot and place the blot face-up on a clean, non-absorbent flat surface. DO NOT allow blot to dry.
Detection with ECF substrate:
ECF is widely preferred as a fluorescent detection agent for this assay, based on people we talked to in the Baker lab ~~Felix Moser
- Pour entire contents of the bottle containing the detection buffer into the bottle which contains the ECF detection reagent. Shake the bottle gently for about 10min to fully dissolve the ECF substrate.
- Pipette ECF substrate on to the blots (~25µl/cm2) and incubate for 1-5min. DO NOT MOVE the blot during the incubation. Transfer the blots directly on to a fresh sheet of Sarran wrap. Fold the plastic over the top of the blots to prevent drying. Incubation can be extended for up to 20min to increase the signal obtained, but too long an incubation will result in signal diffusion.
- Incubate @ room temp in the dark (eg. a drawer or under a styrofoam box or foil) for the needed time. The optimal time for your particular system can be found by rescanning @ various times. For high target levels an acceptable result may be obtained after 1hr. Scanning up to 4hrs after addition of substrate will provide a much stronger signal suited to lower target applications.
- Place the bag containing the blot on to the flat bed fluorescent scanning instrument. Water placed between the lower surface of the bag and the glass will greatly improve the image obtained.
- Scan the blot using an appropriate emission filter as available and according to the guidelines for use of the scanning instrument. ECF has a broad excitation spectrum; max excitation @ 430nm and max emission @ 560nm.
- Incubate blots in absolute alcohol (>99%) @ Room temp with agitation (1ml/cm2) twice for 10 min.
- Incubate blots in 0.5% SDS soln @ 60*C for 60min.
- Rinse blot in 100mM Tris pH8.0 for 5min @ room temp.
- Membrane should be kept moist between reprobings (in Saran wrap @ 4*C).
- Reprobe blot as per hybridization protocol above.