Non-radioactive Southern blot Protocol for telomere length analysis in plants

Note:

Steps in Days 1-3 are essentially modifications of the current radioactive TRF protocol used routinely in the Shippen lab (Texas A&M University, TX, USA)

Day 1.

1.Use CTAB method for plant genomic DNA extraction.

·  Use nanodrop or Qubit to quantify DNA. Adjust DNA to 1 µg/µl with HPLC H2O.

2. Digest up to 50 µg of plant genomic DNA with Tru1I (important!) overnight (add 150-200 µl of mineral oil to the top of the tube). Tru1I is from Fermentas/ThermoFisher (cat # ER0981).

Digestion set-up: 200 µl

Buffer R (or buffer Tango) 20 µl

HPLC H2O 128 µl

Tru1I (Fermentas) 2 µl

DNA (1 mg/ml) 50 µl

Add 150 µl Mineral oil to the top (carefully!).

Incubate at 65°C

(Optional) After several hours add another 1 µl of enzyme - be careful when penetrating the mineral oil layer - do not lose your enzyme and do not accidentally add oil to your sample.

Incubate at 65°C Overnight.

Day 2.

1. Next morning, prepare new tubes with 200 µl Isopropanol. Transfer O/N digest from under mineral oil into the new tubes and label them (make sure not to introduce mineral oil from the digestion step into your precipitation reaction: don't let the tip touch the new tube). Mix by inversion and spin at max speed for 20 min.

2. Remove sup with pipet, add 100 µl 70% EtOH, spin 5 min at max speed. Remove EtOH with P200 pipet, let sit for 10 sec, remove remaining liquid again with P20 pipet. Air dry for ~15-20 min.

3. Resuspend in 20 µl of clean HPLC water. Incubate at 37°C for 1 h to resuspend. If necessary, use pipet carefully to help resuspend DNA.

4. Pour 1% agarose gel in 1x TAE. For TRF, we run very long gels (25cm in length, ~250 ml agarose) with 10 µl GelRed (or EtBr).

5. Add 4 µl 6x loading dye to each DNA sample. Let sit in 37C for 15-20 min to help resuspend remaining precipitate. Load DNA samples onto the gel in the following order: 1) (optional) regular DNA markers (such as 1 kb+ DNA Ladder from ThermoFisher Scientific, cat #10787018); 2) 5 µl of DIG-labeled markers (DIG-labeled DNA molecular weight Marker III, Roche catalog #11218603910, or DIG-labeled DNA molecular weight Marker VII, Roche catalog #11669940910), or a combination of both; 3) DNA samples to analyze in the order from left to right: WT first, then mutants one by one; 4) 5 µl of DIG-labeled markers again.

Important!!! Markers should be on both sides of the gel to ensure proper analysis with TeloTool!

6. Run at 50-60 V overnight (~ from 4 pm till 9.00. am). Double check the gel before you go home to make sure everything is fine.

Day 3.

1. Take GelRed (or EtBr) picture to make sure the digest worked (a uniform smear of digested DNA appears in all lanes).

2. Submerge gel in standard Denaturing buffer in big tray and rock for 30 min.

Denaturing buffer 1 l 0.5l

NaCl 87.6 g/l 43.8g

NaOH 20 g/l 10 g

3. Dump buffer and rinse gel with distilled water.

4. Submerge gel in standard Neutralization buffer and rock for 30 min.

Neutralization buffer 1 l

NaCl 87.6 g/l

Tris 60.5 g/l pH 7.0 using HCl

5. Dump buffer and rinse gel with distilled water.

6. Set up a transfer pyramid with 10X SSC (see transfer set-up below). Transfer for 3-6 hrs up to O/N.

20x SSC 1l

NaCl 175.3 g/l

Na- Citrate 88.2 g/l pH 7.0 using HCl –NaOH

Transfer pyramid set up: (soak the Whatman chromatography paper and membrane in 10xSSC before setting up the transfer):

Weight (~300 to 500g is enough)

Sponge soaked with 10xSSC

Plastic borders

Whatman paper (2 pieses)

Gel (from top to bottom)

Hybond N+membrane

Whatman paper (2 pieses)

Paper towels

Empty tray

Amersham Hybond-N+ membrane, cat #RPN119B (GE Healthcare)

7. Cross-link 2 times on Auto in Stratalinker UV Crosslinker (Stratagene).

If doing regular Southern with radioactive probe, proceed with hybridization with 32P labeled (TTTAGGG)4 probe overnight. Wash next morning with 2X SSC, 0.1% SDS twice for 10 min each, then with 0.2X SSC, 0.1% SDS once for 20-30 min. Place membrane face down in Saran or Cling or similar type plastic wrap, expose. Develop next day. If all worked, stop here.

Next steps are for Non-radioactive assay only!

8. Pre-hybridize with 20 ml (or more) Pre-hyb buffer. Rotate in a hybridization oven for 1h at 42°C. Dump out.

Pre-hybridize buffer 0.5l

BSA 0.5g

SDS 20% 175 ml

1M Na2HPO4 (28.4g/200ml) 101.7 ml

1M NaH2PO4 ( 24g/200ml) 23.3 ml

9. Add pre-hyb buffer with non-radioactive DIG-TELO probe (10 µl of 10 µМ Oligo 5-DIG) and incubate O/N at 42°C.

5'-DIG primer: (Digoxigenin)TTTAGGGTTTAGGGTTTAGGGTTTAGGG-3'

Day 4.

1. Put the buffer with DIG probe to a 50 ml Falcon tube to be re-used later. Store at -20.

2. Wash with 2x SSC/0.1% SDS for 10 min at 42°C. Dump. Repeat.

3. Wash with 0.2x SSC/0.1% SDS for 25 min at 42°C. Dump. Repeat.

4. Incubate with Blocking solution for 1 h at RT.

Blocking solution 100 ml DIG1 1l

DIG 1 100 ml Maleic acid 8.8g

BSA 5g NaCl 11.3g

NaOH 7.5g

To make DIG1, dissolve everything in 100 ml water, adjust pH to 7.5 with NaOH, then adjust volume to 1 L., store at 4°C. Do not autoclave

5. Incubate with Antibody solution for 30 min at RT.

Antibody solution 25 ml

First, spin down the tube with Antibody (anti-DIG-AP Fab fragments from Roche cat # 11093274910).

To get 1:20.000 dilution:

1x Blocking solution 25 ml

Anti DIG antibody 1.25 ul

6. Prepare 800 ml Washing buffer (DIG2). Incubate membrane with DIG2 for 45 min twice at RT with 400 ml DIG2 each.

Washing buffer (DIG2) 400ml

DIG 1 400 ml

Tween-20 1.2g

7. Prepare 200 ml Detection Buffer (DIG3), save 3 ml for use with CPD* (Immune Star AP Substrate cat # 170-5018, Bio-Rad, USA). Do 5 min wash at RT.

Detection Buffer (DIG3) 200ml

100mM Tris pH 9,5 20 ml 1M Tris pH 9,5 ( 12,1g to 100ml)

100mM NaCl 1.17g

8. Place membrane in plastic sleeve, squeeze out excess DIG3. To the 3 ml of Detection buffer you saved in step 7 add 3 ml of CPD* substrate.

Add to membrane, cover to avoid any light and leave for 5 min at RT.

Then squeeze out excess liquid from membrane, seal the plastic sleeve and develop.