Pollen Tube Stain

Ben Carter, 9/24/2014

Adapted from Plant Cell 2005 Feb;17(2):584-96

Pollination

 * 1) Pollinate flowers in the same manner as when performing a cross. Cover pistils with the plastic bag as done when crossing. Allow to pollen tubes to develop for the desired amount of time. Suggested time points for observation: 4, 12, 24, and 48h after pollination.
 * 2) Harvest silique using forceps. Leave most of the silique stem attached to the silique.

Tissue Fixing
Note: It is easiest to use a glass pipette when removing fluids from siliques.
 * 1) Rock pistils in 1mL of 3:1 ethanol/acetic acid for 2h at room temperature. Multiple siliques of the same sample can be processed in a single microfuge tube if desired.
 * 2) Pipette away the 3:1 solution and rock five times in 1mL of Milli-Q water for 5min each time. Leave about 1/5thof the washes in the tube to minimize physical damage to the siliques.
 * 3) Rock pistils in 1mL 8M NaOH for 5min and then leave them stationary overnight at room temperature to soften them.

Staining and Imaging

 * 1) Rock pistils five times in 1mL of Milli-Q water for 5min each time. Pipette away the water.
 * 2) Rock pistils for 5min in 1mL of 0.1M Phosphate Buffer. Pipette away the buffer. Add 1mL Staining Solution and rock for 1min at room temperature. Incubate at room temperature for 3h in the dark.
 * 3) Pipette away stain solution and rock pistils in 1mL of 0.1M Phosphate Buffer twice for 5min each time.
 * 4) Remove pistils from buffer and place on a microscope slide in 100µL of glycerol. Cover with a glass cover slip. Image using the fluorescence microscope under UV light excitation.

0.1M Phosphate Buffer

 * 11.41g of K2HPO4 (Potassium phosphate, dibasic)
 * 450mL of Milli-Q water
 * Adjust pH to 11 using KOH
 * Adjust volume to 500mL
 * Filter sterilize, store at 4°C

Staining Solution

 * 5mL of 0.1M Phosphate Buffer
 * 20mg of aniline blue (0.1% final concentration)