Silver: FISH/IF

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Probe Preparation

 * 1) Each probe: mix on ice (in 0.5 ml PCR tube):
 * 2) *x µl 2 µg probe fragment (phenol free, RNA free)
 * 3) *10 µl 10x dNTP mix
 * 4) **50 µl 1 M Tris-HCl pH 7.8 (500 mM Tris-HCl, pH 7.8)
 * 5) **5 µl 1 M MgCl2 (50 mM MgCl2)
 * 6) **0.7 µl 14.3 M BME (100 mM BME)
 * 7) **2 µl of 10 mM dATP (0.2 mM dATP)
 * 8) **2 µl of 10 mM dCTP (0.2 mM dCTP)
 * 9) **2 µl of 10 mM dGTP (0.2 mM dGTP)
 * 10) **1 µl of 10 mM dTTP (0.1 mM dTTP)
 * 11) **37.3 µl of ddH2O
 * 12) *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)
 * 13) *1 µl 1 µg/µl nuclease-free BSA (stock is 20 mg/ml, so make 1 µg/µl)
 * 14) *10 µl DNA Polymerase/DNase mix (Invitrogen)
 * 15) *x µl ddH2O to 100 µl
 * 16) Incubate for 3 hours at 16°C (in PCR block)
 * 17) Add 5 µl of 300 mM EDTA-NaOH, pH 7.4
 * 18) Denature the probe for 5 min at 98°C
 * 19) Chill on ice and add:
 * 20) *2 µl 10 mg/ml salmon sperm DNA
 * 21) *12 µl 3 M sodium acetate
 * 22) *2 volumes (~240 µl) ice cold 100% ethanol
 * 23) Precipitate at -20°C overnight
 * 24) Centrifuge for 30 min, 14,000 rpm at 4°C
 * 25) Wash the pellet in 1 ml cold (-20°C) 75% ethanol
 * 26) 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
 * 2) *If strains are ade-, supplement with 20 µg/ml adenine sulfate
 * 3) **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
 * 2) *For 50 ml 20% PF:
 * 3) **10 g PF
 * 4) **30 ml ddH2O
 * 5) **50 µl 10N NaOH
 * 6) ***Dissolve at 62°C in closed tube for about 30 min shaking
 * 7) ***Once dissolved, add 10 g sorbitol and ddH2O up to 50 ml
 * 8) Transfer cells to 50 ml conicals (mostly fits, lose ~1 ml)
 * 9) *pellet 2,400 rpm for 5 min at RT
 * 10) Resuspend the pellet in 40 ml YEPD/1.2 M sorbitol, then pellet the cells for 5 min at 2,400 rpm
 * 11) *For 500 ml YEPD/1.2 M sorbitol
 * 12) **400 ml YEPD
 * 13) **110 g sorbitol
 * 14) ***Dissolve sorbitol (needs heating)
 * 15) ***Bring volume up to 500 ml with YEPD
 * 16) ***Filter
 * 17) Resuspend the pellet in 1 ml YEPD/1.2 M sorbitol and transfer to 1.5 ml tube
 * 18) 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
 * 2) *For 50 ml
 * 3) **39.5 ml ddH2O
 * 4) **10 ml 0.5 M EDTA-KOH (pH 8.0)
 * 5) **500 µl 1 M DTT
 * 6) Incubate the tubes at 30°C for 10 min with no shaking, gently invert a couple times
 * 7) Collect the cells by centrifuging at 2,500 rpm for 2 min at RT
 * 8) Resuspend the cell pellet in 1 ml of YEPD/1.2 M sorbitol. To resuspend evenly, start with 500 µl
 * 9) Add lyticase to 1000 Units/ml and zymolase (100T) to 400 µg/ml of fixed cells
 * 10) *For 1 ml of cells add
 * 11) **20 µl lyticase (50 Units/µl)
 * 12) ***Resuspend 50,000 units lyticase (one bottle) in 1 ml ddH2O – gently pipet, do not vortex – makes 50 Units/µl
 * 13) **40 µl zymolyase (of 10 µg/µl)
 * 14) ***Use 10 mg/ml zymolase (stock)
 * 15) Incubate at 30°C with no shaking. Monitor spheroplast formation at 5, 10, 15, and 20 min. (~ 10 min works well)
 * 16) 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
 * 17) Harvest cells before complete spheroplasting (~80%)
 * 18) Centrifuge for 1 min at 2,500 rpm at RT
 * 19) Wash twice in 1 ml YEPD/1.2 M sorbitol.
 * 20) 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
 * 21) 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
 * 22) Take away as much liquid as possible using a pipette and let air dry for 2 min
 * 23) 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
 * 2) *For 10 ml
 * 3) **9.0 ml ddH2O
 * 4) **1 ml 10x PBS
 * 5) **100 mg ovalbumin
 * 6) *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
 * 7) 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)
 * 8) *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
 * 9) * For MAb414 1:5000 in PBS
 * 10) **1 µl MAb414
 * 11) **5 ml PBS/1% ovalbumin (see above)
 * 12) Incubate for 1 hr at 37°C in humid chamber or overnight at 4°C
 * 13) *Humid Chamber
 * 14) **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
 * 4) *Important when continuing with FISH as the primary and secondary antibodies tend to dissociate under the harsh conditions of in situ
 * 5) *For 1 ml
 * 6) **200 µl 20x SSC
 * 7) **200 µl 20% paraformaldehyde (made fresh)
 * 8) **600 µl ddH2O
 * 9) Wash each spot 3 x 3 min in 4x SSC (10 µl per spot)
 * 10) *For 1 ml
 * 11) **200 µl 20x SSC
 * 12) **800 µl ddH2O

RNase A Treatment

 * 1) Apply 4x SSC + 20 µg/ml RNase A to each spot
 * 2) *For 1 ml
 * 3) **200 µl 20x SSC
 * 4) **2 µl 10 mg/ml RNase A (10 mg/ml, breboiled)
 * 5) **800 µl ddH2O
 * 6) Incubate overnight at RT (in the dark – humid chamber)

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)
 * 5) *For 1 ml
 * 6) **100 µl 20x SSC
 * 7) **700 µl deionized formamide (100%)
 * 8) **200 µl ddH2O
 * 9) Incubate at 72°C for 5 min
 * 10) *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
 * 11) Dehydrate slides in Coplin jars containing 70%, 80%, 90%, and 100% ethanol (-20°C) for 1 min each
 * 12) Air dry
 * 13) Check that you still have cells, unfortunately this is not a joke :)

Probe Hybridizing

 * 1) Make hybridization solution
 * 2) *For 800 µl of hyb solution
 * 3) **500 µl deionized formamide
 * 4) **200 µl 50% dextran sulfate
 * 5) **100 µl 20x SSC
 * 6) Resuspend each pellet with 40 µl ddH2O
 * 7) Combine resuspended pellet with 160 µl of hyb solution
 * 8) *200 µl total gives probe concentration of 10 ng/µl if started with 2 µg
 * 9) **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
 * 10) Apply 10 µl of the pellet/hyb solution mix to each spot
 * 11) Incubate for 10 min at 72°C
 * 12) Incubate for 24-60 hr (greater than 40 usually) at 37°C in dark
 * 13) *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

Washes

 * 1) Preheat 0.05x SSC at 40°C (water bath) – also preheat aluminum block in water
 * 2) *For 1 ml
 * 3) **2.5 µl 20x SSC
 * 4) **997.5 µl ddH2O
 * 5) 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
 * 6) Keep the hyb chamber in the 37°C incubator (keep it warm – will need it at 37°C again later)
 * 7) 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
 * 8) *1 ml BT buffer + BSA
 * 9) **150 µl 1M NaHCO3
 * 10) **1 µl Tween-20
 * 11) **25 µl 20 mg/ml BSA
 * 12) **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
 * 2) *5 ml BT buffer
 * 3) **750 µl 1M NaHCO3
 * 4) **5 µl Tween-20
 * 5) **4.245 ml ddH2O
 * 6) *For 1 ml antibody mix:
 * 7) **989 µl BT buffer
 * 8) **1 µl Alexa-594 anti-mouse
 * 9) **10 µl sheep anti-DIG Fab fragments (is only 0.2 µg/ml)
 * 10) Incubate for 1 hr at 37°C in humid chamber in the dark
 * 11) Wash 5 x 3 min in BT buffer
 * 12) Add 15 µl antifading solution per spot (1x PBS, 50% glycerol, 24 mg/ml DABCO, pH 7.5)
 * 13) *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