Cytological Methods

2.6.3.1 ‘Hard’ Cytological Spreading for Aggregate Analysis

4.5 ml of meiotic culture was spun down on a bench centrifuge and resuspended to 500 µl with 1.0 M pH7.0 D-Sorbitol (Sigma). 12 µl of 1.0 M DTT (Sigma/Fermentas) and

7 µl 10 mg/ml Zymolyase (MPBio) was added and cells were spheroplasted by incubation at 37°C for 20 to 45 min with agitation. Spheroplasting success was determined by taking a few microliters to a microscope slide and adding an equivalent volume of 1.0% (w/v) Sodium N-Lauroylsarcosine (Sigma); most cells should lyse after a few seconds as the exposed membrane is disrupted by the detergent. After 3.5 ml of Stop Solution was added, cells were spun down and resuspended in 100 µl Stop Solution and distributed between 4 slides, which had been cleaned with ethanol. 20 µl of fixative was added to each slide followed by 40 µl of 1% Lipsol (Bibby Sterilin) and light mixing. 40 µl of fixative was added and the mixture spread across the slide. The spreads were incubated at room temperature (RT) for 30 min in damp conditions, then allowed to air-dry at RT. Once dry, slides were washed in 0.2% (v/v) PhotoFlo Wetting Agent (Kodak) then in dH2O and

allowed to air-dry slightly at RT. Slides were washed once in 0.025% Triton X-100 for 10 min at RT and twice in PBS for 5 min each at RT. Slides were blocked in 5% Skimmed

Milk (Sigma) in PBS for 1-4 h at 37°C. Excess liquid was removed and slides laid horizontally in damp conditions. Primary α-Rad51 antibody (Santa Cruz), 150 µl per slide made up in 1% Skimmed Milk in PBS at 1:200, was added and the slides incubated at 4°C overnight. The slides were washed three times in PBS, 5 min each at RT, and incubated

67 in secondary AlexaFluor594 goat anti-mouse antibody (Life Technologies), 150 µl per slide made up in 1% Skimmed Milk in PBS at 1:1000, for 1-2 h at RT in damp conditions.

Slides were washed three times with PBS, 5 min each at RT. Cover slips were affixed using Vectashield mounting medium with DAPI (VectorLabs), sealed with clear varnish and imaged on a DeltaVision microscope (z=12-15, Exposure times: RD-TP-RE=1.0 s, DAPI=0.05-1.0 s). Images were deconvolved by SoftWoRx software (standard settings) and the number of cells with No Signal, Rad51 Foci or Rad51 Aggregates counted.

Aggregates were identified by pixel width using ImageJ software; this limit was generally 6 pixels = 0.39 µm, however due to potential variation in spreads performed on different days, the pixel limit was determined by visual inspection of several slides and then fixed for all timecourses spread under those same conditions.

2.6.3.2 ‘Gentle’ Cytological Spreading for Spindle Pole Body (SPB) Analysis In order to correlate the DAPI and SPB signals and to observe separation of tetO/TetR signals, a gentler spreading technique was employed to reduce the risk of the DAPI signal being disturbed by the spreading process. This process involved an additional initial step:

500 µl 37% (v/v) formaldehyde (Sigma) was added to 4.5 ml of meiotic culture and incubated at room temperature (RT) for 30 min. Cells were spun down and washed with 5 ml 1% (w/v) potassium acetate (Sigma). Cells were then spun down and treatment continued as per ‘Hard’ Spreading: cells were resuspended in Sorbitol, spheroplasted with DTT and Zymolyase, resuspended in stop solution and distributed over clean slides, fixative and lipsol were added and the mixture spread out, then slides were incubated in damp conditions, dried at RT, and washed in PhotoFlo Wetting Agent and dH2O.

68 Cover slips were affixed using Vectashield mounting medium with DAPI (VectorLabs), sealed with clear varnish and imaged on a DeltaVision microscope, (z=12-24, Exposure times: FITC=1.0 s, RD-TP-RE=1.0 s, DAPI=0.05-1.0 s). Images were deconvolved by SoftWoRx software (standard settings) and analysed by counting the number of tetO/TetR signals or SPBs per cell, using GFP-tubulin as an additional signal where necessary, and the number of DAPI signals per cell.

2.6.4 Techniques for Protein Extraction from S. cerevisiae

2.6.4.1 TCA Protein Extraction

Samples containing 5-10 OD of cells were spun down, washed in 1 ml ddH2O, and flash frozen in liquid nitrogen. The pellet was stored at -80°C for at least 2 h, then resuspended in 150 µl Buffer D and incubated on ice for 15 min. To each sample, 150 µl of 55%

Trichloroacetic acid (TCA) (Sigma) was added and cells incubated for a further 10 min.

Cells were spun down for 10 min then resuspended in 250 µl of Buffer H, with 10 µl 25x protease inhibitor stock (Roche). If necessary, 10 µl 1.5M Tris HCl pH8.8 was added to maintain pH, as shown by the blue indicator in the suspension. Cells were heat-shocked at 65°C for 10 min and spun down.

2.6.4.2 Native Protein Extraction

Samples containing 5-25 OD of cells were spun down, washed in 1ml Protein Lysis Buffer or ddH2O, and flash-frozen in liquid nitrogen. The pellet was stored at -80°C for at least 2 h, then resuspended in 200 µl Protein Lysis Buffer. An equal volume of acid-washed glass beads (Sigma) was added to the suspension and the cells beaten on a Bead Beater for 3-5 cycles of 20-60 s, resting on ice for 5 min in between each cycle. The lysate was

69 separated from the beads by puncturing the base of each tube and centrifugation, collecting the flowthrough in fresh tubes. The lysate was spun down at 4°C for 10 min and the supernatant removed to a fresh tube. Protein concentration was determined by Bradford Assay.

2.6.4.3 Protein Concentration Determination by Bradford Assay

A standard protein sample, 10 mg/ml BSA (New England Biolabs), was diluted 1:10 then a range of volumes from 0-10 µl added to 1 ml filtered Bradford Assay Reagent (Biorad).

Absorbances of the known standards were measured at 595 nm and a standard curve generated of Absorbance vs Concentration. Volumes of the unknown protein samples, produced by native extraction, were added to 1 ml Bradford Assay Reagent, their absorbances measured at 595 nm and the protein concentration determined using the equation generated from the gradient of the standard curve.