Alexander Blue

Alexander’s (1969) stain is a reliable way to score pollen viability using light microscopy; grains that are viable will stain a dark blue or purple, grains that are dead will stain pale turquoise blue.

Alexander, M.P. (1969) Differential staining of aborted and non-aborted pollen. Stain Technol. 41, 117–122.

Stock solution (store in the dark at room temperature):

• 10 ml 95% ethanol,
• 5 ml 1% malachite green in 95% ethanol,
• 5 g of phenol,
• 5 ml 1% acid fuschin in H₂O
• 0.5 ml 1% orange G in H₂O
• 2 ml glacial acetic acid,
• 25 ml glycerol and
• 50 ml H₂O

Working solution, dilute 1:50 in H₂O.

Arabidopsis culture:

• seed sterilization : 

-  sterilization solution (SS): Dilute 1 volume of bleach (commercial ca. 2.6%) in 3 volume of water, add a  drop/20 ml of Tween (BTS: prepare it fresh). Dilute the previous solution 1/10 in 95% Ethanol. 

- Add 1 ml of SS to 1,5ul  eppendorf tube containing ca.40 ul of seeds seeds, and incubate 5 min at room temperature with constant, gentle agitation. Rince twice with 10 ml 95% ethanol. Let the seeds sediment, and carefully remove as much ethanol as possible. (It's possible to invert the tubes, but be careful !).  Leave the tubes open under a sterile hood overnight. 

- alternate short protocol for immediate seeding: use BTS instead of SS incubate 5 min, spin down, rinse with 1 ml of water and repeat the rinsing 4 times. Plate the seeds on Agar and leave 2-3 days at 4C for stratification. You can also store the seeds in the cold room for few days in the Eppendorf tube.

• in vitro culture medium: 

We use Arabidopsis medium diluted 1/2 (AtM/2), without sucrose (Estelle & Sommerville, 1987, MGG 206 : 200). 

Autoclave (120*C, 20 min) 

Keep at -20*C. 

• Sprinkle the surface-sterilized seeds on a 14 cm agar plate containing Arabidopsis culture medium (AtM/2), covered with a round filter paper (Whatman 3MM). Seal the plates with a gas-permeable surgical tape. 

• Synchronize germination by a cold treatment at 4¡C for 48 hours. 

• Place in the growth chamber under the following conditions : photoperiod 16 h day (100-150 µE/m2/s) / 8 h night ; temperature 20¡C day / 15¡C night ; humidity 70%. 

• After 10-15 days of culture, plantlets (2-leaf rosettes) are ready for DNA isolation. Each plate should yield 3-6 g fresh weight. 

• Gently scrape the plantlets from the filter paper using a razor blade. The plantlets are weighted, and frozen in liquid nitrogen for future use. 

DNA extraction, CTAB Reference: Doyle & Doyle (1990), Focus 12 : 13-15

• Prepare an adequate amount of Extraction Buffer + β-mercaptoethanol, preheat at 60*C
• Weigh samples, and freeze them in liquid nitrogen. Grind 2 / 3 / 5 g of fresh or frozen tissue to a fine powder, in liquid nitrogen with a mortar and pestle.
• Transfer to a 30 ml tube, wait until the nitrogen has evaporated. Add 7,5 / 11 / 20 ml preheated EB, mix well, and cap the tube. Incubate 30 min at 60*C with occasional swirling.
• Extract with 7,5 / 11 / 20 ml ml Chloroform : isoamylalcool (24:1), centrifuge (swinging bucket rotor : 5000 rpm, 10' ; other : 10000 rpm, 10'), and recover the aqueous phase.
• Add 6 / 8 / 14 ml isopropanol, and mix gently to precipitate the nucleic acids. Centrifuge (10000 rpm, 10') and decant the supernatant.
• Wash the pellet with 10-20 ml of ethanol 70%, centrifuge (10000 rpm, 5'), decant supernatant.
• Dissolve pellet in 500 µl TE, transfer into an Eppendorf tube. Add 50 µl sodium acetate 3 M, 500 µl isopropanol, centrifuge 5', decant, wash with 70% ethanol, dry on the bench.
• Dissolve DNA in 100 µl TE / g of fresh tissue extracted. Expect a concentration of 100-200 ng/µl. Store at -20*C.

Quick extraction for high-throughput PCR-based genotyping

DNA loading buffers

Dyes are used in  buffers for loading DNA samples into gel electrophoresis wells and tracking migration during electrophoresis. Sucrose (40%), Ficoll (15%), or glycerol (30%) can be used in place of each other. In a 0.5–1.4% agarose gel in 0.5X TBE, xylene cyanol migrates at approximately 4kb, bromophenol blue at approximately 300bp and orange G at approximately 50bp. Choose from the following options, I personally like the last one:

10X Xilene Cyanol/Bromophenol Blue Loading Dye

• Dissolve in 25 ml of H₂O:
• 100 mg of Xilene cyanol
• 100 mg of Bromophenol Blue
• 5 ml of 10% SDS
• 50 ml of glycerol

10X Orange G Loading Dye

• Dissolve in 40 ml of H₂O:
• 20 g of Sucrose
• 100 mg of Orange G and bring up to 50 ml with H₂O.

6X Blue/Orange Loading Dye

• Dissolve in 73 ml of H₂O (100 ml final):
• 400 mg Orange G (0.4%)
• 30 mg Bromophenol blue (0.03%)
• 30 mg Xylene cyanol (0.03%)
• 15 ml Ficoll® 400 (15%)
• 1 ml of 1 M Tris-HCl,pH 7.5 (10 mM)
• 10 ml of 0.5 M EDTA pH 8.0 (50 mM)