Colony embedding protocol

 The following is a modification of a method for embedding yeast colonies for light and electron microscopy (1). After incubation of approximately 300 colonies on agar medium for the indicated time, an isolated colony (1-2 mm in diameter) and a small amount of the underlying agar medium was removed using a narrow spatula and placed face up on a microscope slide. Several drops of 2% agar (42°C) were placed on a microscope slide, the colony was immediately placed face up on the agar and several drops of agar placed on top of the colony. (Agar is kept in uncapped 2 ml tubes in 42°C water bath). The resulting agar block was trimmed with a razor blade, and placed in a 3.5 ml borosilicate screw-cap vial (Fisher 03-339-21B). All subsequent incubations and washes used 1.5 -2.0 ml and were performed in the same vial. First, colonies were fixed by incubating in 2% paraformaldehyde / 2% glutaraldehyde for 7 days at 4°C.

After about 1 week agar blocks were washed on ice by incubating twice for 15 minutes with 0.15M sodium cacodylate (pH 7.2), and then twice more for 5 minutes with OS buffer (100 mM KH2PO4, 10 mM MgCl2, pH 6.0). To allow electron microscopy of sections, 1% OsO4 in OS was added to vials to cover the agar blocks and incubated on ice in a fume hood for 1 hr, then washed twice with OS buffer on ice for 10 minutes each, and incubated in OS overnight at 4°C. As OsO4 is very toxic, these steps should be performed in a chemical fume hood, with goggles, lab coat and doubled gloves.

The following washes and dehydrations should be performed on ice. After washing two times with cold water for 10 minutes each, the blocks were washed sequentially with cold 25%, 50%, 75%, 95% and 100%(twice) ethanol for 10 minutes each, and the blocks left overnight at 4°C in 100% ethanol.

The following day ethanol was removed and agar blocks were washed with 100% room temp ethanol 5 times for 10 minutes each. The blocks were then resuspended in a 2:1 ratio of 100% ethanol: Spurr’s reagent and the vials rotated for 15 minutes, then allowed to stand for 30 minutes. After 30 minutes the solution was removed and the 2:1 ethanol:Spurr’s treatment was repeated 2 more times. After the final 30 minutes, the ethanol:Spurr’s reagent was removed and a 1:1 ratio of 100% ethanol: Spurr’s reagent was added to cover agar blocks (1.5-2.0 ml). The vials were then rotated for 15 minutes,then allowed to stand for 30 minutes after which the solution was replaced with another 1:1 100% ethanol: Spurr’s reagent and the procedure repeated. After the last 30 minutes the solution was replaced with 100% Spurr’s reagent and left to sit for 4hrs at room temp.

After 4 hrs the Spurr’s reagent was replaced with 100% Spurr’s reagent and left to rotate overnight.In the morning the Spurr’s was replaced with more Spurr’s and left rotating until the late afternoon. The solution was replaced with freshly made Spurr’s reagent and left rotating until the following day. The next morning the Spurr’s reagent was replaced again and rotated until the following day. The next morning the Spurr’s reagent was replaced and the vial rotated until the following day. Finally in the morning, each agar block was placed in a mold with 0.2 ml of Spurr’s and incubated at 60°C for four hours. After 4hrs the agar blocks were topped off with Spurr’s and incubated at 60°C for 3 days.

Colonies were sectioned using a microtome. Sections (0.5 um) from the central region of the colony were collected in a drop of distilled H2O on a glass slide and dried on a 52°C heat block for approximately 15-30 minutes, and subsequently stained with 1.0 % toluidine blue, 1% Sodium Borate for 5-15 sec. and washed under a stream of water. The slides were then dried for 15-30 minutes on a 52°C heat block covered in Permount (Fisher SP1.5-100), and examined by light microscopy.

1. Scherz, R., Shinder, V. & Engelberg, D. (2001) J Bacteriol 183, 5402-13.

4% paraformaldehyde solution
(DO ALL WORK UNDER HOOD WEAR MASK AND GLOVES)

           Heat 40ml ddH20 to 55-60° C
           Add 2 grams paraformaldehyde powder
           Add one NaOH pellet, let dissolve
           Repeat until solution clears
           Place flask on ice for 10-15 minutes to cool
           Add 5ml 10x PBS solution
           pH to 7.6 with HCL
           Fill to 50 ml
           Store at -20° C
           If precipitate forms discard and make fresh

4% glutaraldehyde
          Dilute 50% glutaraldehyde to 4% with 0.2M Na Cacodylate Buffer
          Store at -20 Celcius

0.2M Na Cacodylate Buffer
(CONTAINS ARSENIC SO DO ALL WORK UNDER HOOD, WEAR MASK AND GLOVES)

           Dissolve 8.56 grams Na Cacodylate Powder(Fisher BP 325-50) in 160ml ddH2O
          Adjust pH to 7.2 with NaOH and fill to 200ml with ddH20
          Store at 4° C

2X Osmium Buffer
          200mM KPO4H2
          20mM MgCl2*H20
          pH to 6
          Store at 4° C

OS Buffer
          Dilute 2X Osmium Buffer to 1X in dH2O
          Store at 4° C

Spurrs solution (Order numbers and source)
(DO ALL WORK UNDER HOOD,CHEMICALS ARE CARCINOGENIC)

          ERL 4221   10 grams(RT 15004)
          DER 736     8 grams(RT 13000)
          NSA           26 grams(RT 19050)

Make this fresh as early in the morning as possible since the following protocol is about 9 hrs.
Stir first 3 chemicals in plastic cup for 20 minutes under the hood with stir bar. Add 0.3 grams DMAE(RT 13300) and stir for another 20 minutes. De-gas until all bubbles are gone 1-2hrs.

 

Citation:

Piccirillo, Sarah, White, Melissa G., Murphy, Jeffrey C., Law, Douglas J., and Honigberg, Saul M., (2010) The Rim101p/Pacc Pathway and Alkaline Ph Regulate Pattern Formation in Yeast Colonies. Genetics, 184(3): p. 707-716.
http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd=Retrieve&db=PubMed&dopt=Citation&list_uids=20038633.