Silver: FISH/IF

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Contents

Day 1 - Probe Preparation/Starting Cultures

Probe Preparation

  1. Each probe: mix on ice (in 0.5 ml PCR tube):
    • x µl 2 µg probe fragment (phenol free, RNA free)
    • 10 µl 10x dNTP mix
      • 50 µl 1 M Tris-HCl pH 7.8 (500 mM Tris-HCl, pH 7.8)
      • 5 µl 1 M MgCl2 (50 mM MgCl2)
      • 0.7 µl 14.3 M BME (100 mM BME)
      • 2 µl of 10 mM dATP (0.2 mM dATP)
      • 2 µl of 10 mM dCTP (0.2 mM dCTP)
      • 2 µl of 10 mM dGTP (0.2 mM dGTP)
      • 1 µl of 10 mM dTTP (0.1 mM dTTP)
      • 37.3 µl of ddH2O
    • 1 µl 1 nmol/µl DIG-dUTP (an anti-DIG antibody conjugated to FITC is applied near the end of the procedure and will light up the probe)
    • 1 µl 1 µg/µl nuclease-free BSA (stock is 20 mg/ml, so make 1 µg/µl)
    • 10 µl DNA Polymerase/DNase mix (Invitrogen)
    • x µl ddH2O to 100 µl
  2. Incubate for 3 hours at 16°C (in PCR block)
  3. Add 5 µl of 300 mM EDTA-NaOH, pH 7.4
  4. Denature the probe for 5 min at 98°C
  5. Chill on ice and add:
    • 2 µl 10 mg/ml salmon sperm DNA
    • 12 µl 3 M sodium acetate
    • 2 volumes (~240 µl) ice cold 100% ethanol
  6. Precipitate at -20°C overnight
  7. Centrifuge for 30 min, 14,000 rpm at 4°C
  8. Wash the pellet in 1 ml cold (-20°C) 75% ethanol
  9. Dry the pellet, store as dried pellet at -20°C

Starting Cultures

  1. Grow cells overnight to 0.5-1 x 107 cells/ml in 50 ml YPD or selective media
    • If strains are ade-, supplement with 20 µg/ml adenine sulfate
      • For 50 ml culture, add 500 µl of 2 mg/ml adenine sulfate

Day 2 - Fixation/Spheroblasting/Dehydration/Primary Antibody Incubation/Secondary Antibody Pre-Absorbing

  • Important notes
    • No Triton in buffers
    • Fix cells before spheroplasting, do not use coverslips – just use humid chambers (even gentle removal of coverslips will strip the cells right off your slides)
  • Before Immunofluorescence
    • Polylysine coat slides by adding 10 µl polylysine solution (1mg/ml in H2O) to each well on slide. Dilute the lab stock of polylysine from 0.3% to 0.1% to make 1mg/ml. Incubate at room temp for 10 min. Remove the polylysine and wash 2x with H2O and allow to air dry
    • Make 20% paraformaldehyde (takes ~30 min, see below)
    • Make YEPD/1.2 M Sorbitol

Fixation

  1. Fix cells in growth medium for 10 min at 30°C in 4% paraformaldehyde (PF) before spheroplasting. To a 50 ml culture, add 10 ml of 20% PF
    • For 50 ml 20% PF:
      • 10 g PF
      • 30 ml ddH2O
      • 50 µl 10N NaOH
        • Dissolve at 62°C in closed tube for about 30 min shaking
        • Once dissolved, add 10 g sorbitol and ddH2O up to 50 ml
  2. Transfer cells to 50 ml conicals (mostly fits, lose ~1 ml)
    • pellet 2,400 rpm for 5 min at RT
  3. Resuspend the pellet in 40 ml YEPD/1.2 M sorbitol, then pellet the cells for 5 min at 2,400 rpm
    • For 500 ml YEPD/1.2 M sorbitol
      • 400 ml YEPD
      • 110 g sorbitol
        • Dissolve sorbitol (needs heating)
        • Bring volume up to 500 ml with YEPD
        • Filter
  4. Resuspend the pellet in 1 ml YEPD/1.2 M sorbitol and transfer to 1.5 ml tube
  5. Pellet at 2,000 rpm for 2 min at RT and remove supernatant

Spheroblasting

  1. Resuspend the cells in 1 ml of 100 mM EDTA-KOH pH 8.0 and 10 mM DTT
    • For 50 ml
      • 39.5 ml ddH2O
      • 10 ml 0.5 M EDTA-KOH (pH 8.0)
      • 500 µl 1 M DTT
  2. Incubate the tubes at 30°C for 10 min with no shaking, gently invert a couple times
  3. Collect the cells by centrifuging at 2,500 rpm for 2 min at RT
  4. Resuspend the cell pellet in 1 ml of YEPD/1.2 M sorbitol. To resuspend evenly, start with 500 µl
  5. Add lyticase to 1000 Units/ml and zymolase (100T) to 400 µg/ml of fixed cells
    • For 1 ml of cells add
      • 20 µl lyticase (50 Units/µl)
        • Resuspend 50,000 units lyticase (one bottle) in 1 ml ddH2O – gently pipet, do not vortex – makes 50 Units/µl
      • 40 µl zymolyase (of 10 µg/µl)
        • Use 10 mg/ml zymolase (stock)
  6. Incubate at 30°C with no shaking. Monitor spheroplast formation at 5, 10, 15, and 20 min. (~ 10 min works well)
  7. To check, mix 4 µl of cells with 4 µl 1% SDS on a glass slide and observe the number of cell "ghosts" under microscope. This is very hard to do – usually just go on timing
  8. Harvest cells before complete spheroplasting (~80%)
  9. Centrifuge for 1 min at 2,500 rpm at RT
  10. Wash twice in 1 ml YEPD/1.2 M sorbitol.
  11. Resuspend spheroplasts in 0.8 ml YEPD. This concentration of cells should be such that only one layer of non-confluent cells will adhere to the slide
  12. Leave a drop on each spot (~10 µl) of a super-Teflon (pre-polylysine coated) slide for 5 min to allow the spheroplasts to adhere to the glass surface. Do not throw away unused spheroplasts – keep on ice for later pre-clearing of secondary antibody
  13. Take away as much liquid as possible using a pipette and let air dry for 2 min
  14. Check that cells are there by light microscopy

Dehydration

  1. Perform methanol and acetone washes in Coplin jars (blot off extra liquid from ends of slides)
  2. Put the slides in -20°C methanol for 6 min
  3. Transfer the slides to -20°C acetone for 30 sec
  4. Air dry for 3-5 min

Primary Antibody Incubation

  1. Cover each spot with 10 µl of 1x PBS/1% ovalbumin for at least 10 min
    • For 10 ml
      • 9.0 ml ddH2O
      • 1 ml 10x PBS
      • 100 mg ovalbumin
    • After this step the cells should appear transparent and the nucleus can be seen as a dark spot. This is an indication of good spheroplasting
  2. Pipet off PBS/1% ovalbumin and cover each spot on the slide with 10 µl of the appropriate antibody diluted in PBS (antibody originally used for NPC staining was MAb414)
    • Found that a strain containing a myc-tagged NPC component gave much better NPC staining at the end when using an anti-myc antibody rather than MAb414
    • For MAb414 1:5000 in PBS
      • 1 µl MAb414
      • 5 ml PBS/1% ovalbumin (see above)
  3. Incubate for 1 hr at 37°C in humid chamber or overnight at 4°C
    • Humid Chamber
      • We use tip boxes – soak paper towels and place them under the tip holder - place slide on top of tip holder – tape top and bottom together to seal the box

Secondary Antibody Pre-absorbing

  1. Use the remaining fixed spheroplasts by washing them 3 x 1 ml in cold PBS and resuspending them in 1 ml of cold PBS
  2. Dilute the secondary antibody (594(RED) anti-mouse for MAb414 - stock is usually 1 mg/ml) 1:1000 in the spheroplast suspension and incubate for 1 hr on a rotating wheel in the dark
  3. Centrifuge at top speed, collect the supernatant (containing the secondary antibody) and store at 4°C

Day 3 - Washes/Secondary Antibody Incubation and Fixation/RNase A Treatment

  • Make fresh 20% paraformaldehyde before starting

Washes

  1. Carefully remove the slide from the humid chamber
  2. Wash each spot 3 x 5 min (10 µl) in 1x PBS at RT

Secondary Antibody Incubation and Fixation

  1. Pipet off washes and cover each spot with 10 µl of the pre-absorbed secondary antibody and incubate at 37°C in the humid chamber in the dark for 1.5 hr
  2. Wash each spot 3 x 5 min in 1x PBS at RT
  3. Post-fix the cells by adding drops (10 µl) of 4x SSC + 4% paraformaldehyde for 20 min at RT
    • Important when continuing with FISH as the primary and secondary antibodies tend to dissociate under the harsh conditions of in situ
    • For 1 ml
      • 200 µl 20x SSC
      • 200 µl 20% paraformaldehyde (made fresh)
      • 600 µl ddH2O
  4. Wash each spot 3 x 3 min in 4x SSC (10 µl per spot)
    • For 1 ml
      • 200 µl 20x SSC
      • 800 µl ddH2O

RNase A Treatment

  1. Apply 4x SSC + 20 µg/ml RNase A to each spot
    • For 1 ml
      • 200 µl 20x SSC
      • 2 µl 10 mg/ml RNase A (10 mg/ml, breboiled)
      • 800 µl ddH2O
  2. Incubate overnight at RT (in the dark – humid chamber)

Day 4 - Dehydration/Probe Hybridization

Dehydration

  • Use Coplin jars for ddH2O wash and dehydration
  1. Wash slides in H2O
  2. Dehydrate slides in Coplin jars containing 70%, 80%, 90%, and 100% ethanol (-20°C) for 1 min each
  3. Air dry
  4. Add 10 µl per spot 2x SSC + 70% formamide (cover ALL the spots – lots of liquid)
    • For 1 ml
      • 100 µl 20x SSC
      • 700 µl deionized formamide (100%)
      • 200 µl ddH2O
  5. Incubate at 72°C for 5 min
    • Our trick is to place the slide on top of an aluminum block that is partially submerged in a 72°C water bath. Allow a few drops of water to spread under the slide by capillary action – in between the glass and aluminum – for better heat conductance
  6. Dehydrate slides in Coplin jars containing 70%, 80%, 90%, and 100% ethanol (-20°C) for 1 min each
  7. Air dry
  8. Check that you still have cells, unfortunately this is not a joke :)

Probe Hybridizing

  1. Make hybridization solution
    • For 800 µl of hyb solution
      • 500 µl deionized formamide
      • 200 µl 50% dextran sulfate
      • 100 µl 20x SSC
  2. Resuspend each pellet with 40 µl ddH2O
  3. Combine resuspended pellet with 160 µl of hyb solution
    • 200 µl total gives probe concentration of 10 ng/µl if started with 2 µg
      • The optimal concentration of probe depends on the sequence and must be determined empirically. Put hyb solution (without probe) on other spots to keep enough moisture around
  4. Apply 10 µl of the pellet/hyb solution mix to each spot
  5. Incubate for 10 min at 72°C
  6. Incubate for 24-60 hr (greater than 40 usually) at 37°C in dark
    • Place in humid chamber with several drops of hyb solution (without probe) in the wells to maintain internal humidity. Tape everything shut to avoid evaporation

Day 5 - Washes/Secondary Antibody Incubation Round 2/Coverslip

Washes

  1. Preheat 0.05x SSC at 40°C (water bath) – also preheat aluminum block in water
    • For 1 ml
      • 2.5 µl 20x SSC
      • 997.5 µl ddH2O
  2. Remove slides from humid chamber and wash twice with 0.05x SSC for 5 min at 40°C (10-20 µl drops on each spot) – place on partially submerged aluminum block in water bath
  3. Keep the hyb chamber in the 37°C incubator (keep it warm – will need it at 37°C again later)
  4. Incubate spots in BT buffer (0.15 M NaHCO3, 0.1% Tween 20, pH 7.5) + 0.05% BSA for 2 x 30 min at 37°C in the dark – don’t immerse slide, just place drops on individual spots
    • 1 ml BT buffer + BSA
      • 150 µl 1M NaHCO3
      • 1 µl Tween-20
      • 25 µl 20 mg/ml BSA
      • 824 µl ddH2O

Secondary Antibody Incubation Round 2

  1. Add secondary Alexa-594 anti-mouse antibody 1:1000 for MAb414 (for refreshing the IF signal) and add sheep anti-digoxigen diluted 1:100
    • 5 ml BT buffer
      • 750 µl 1M NaHCO3
      • 5 µl Tween-20
      • 4.245 ml ddH2O
    • For 1 ml antibody mix:
      • 989 µl BT buffer
      • 1 µl Alexa-594 anti-mouse
      • 10 µl sheep anti-DIG Fab fragments (is only 0.2 µg/ml)
  2. Incubate for 1 hr at 37°C in humid chamber in the dark
  3. Wash 5 x 3 min in BT buffer
  4. Add 15 µl antifading solution per spot (1x PBS, 50% glycerol, 24 mg/ml DABCO, pH 7.5)
    • Use 2 x PBS and dilute 1:1 with 100% glycerol

Coverslip

  1. Cover with a coverslip, avoiding air bubbles
  2. Seal with clear nail polish, allow to dry and add a second coat of nail polish
  3. Keep the slides at 4°C in the dark
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