10:20 A – 7:15 P, 8:30 P – 1:45 A
cell staining
- remove samples from 40C, place in RT 2x SSCT for first rinse out.
- check staining of control slide on STORM2 TIRF- no staining except in dead cells (should compare to unstained cells)
- check staining of B secondary mediated stain on STORM2 – no staining except in dead cells (should compare to unstained cells)
reasons for failure?
- probes no good / positive control gone bad?
- issue with protocol execution?
- too much NaBrH4 ?
- dead cells suck out too much probe?
- too little secondary?
- try resuspending aliquot of orig probe in hybe buffer. measure conc on nanodrop.
- checked unlabeled cells — yes dead cells are brighter, but nothing compared to labeled cells — so the dead cells do seem to eat probe.
Troubleshooting cell staining
- EtOH precipitating Cy5-short secondary. Resuspend in hybe buffer and try at higher concentration for hybes
- try BX-C primary + secondary labeled (recent stock from Brian).
- try En1 at high conc. (Previously successful on embryos: 8 uL en probe in 40 uL dilution buffer, used 30 uL on coverglass)
- try AATAT with short P1 tail (100 nmol in 100 uL) and Cy5-short secondary
- try AATAT with long P2 tail (30 nmol in 30 uL) + new S2-Cy5 secondary (arrived today)
- try AATAT direct label as positive control
- negative control
Concentrations
- AATAT with short P1 tail (100 nmol in 100 uL) = 1000 pmol per uL. dilute 1:20 in hybe = 50 pmol. use 1 uL of this in 30 uL reaction. (according to nanodrop was only ~2000 ng/uL 137 ng/uL)
- try AATAT with long P2 tail (30 nmol in 30 uL) dilute 1:20 in hybe = 50 pmol. use 1 uL of this in 30 uL reaction.
- reconcentrated short secondary 1: 334 ng/uL = 33 pmol /uL. use 1.7 uL in 30 uL reaction
- S2-Cy5 ~3000 ng/uL = 284 pmol/uL
Probe labeling
- resuspend probe at 100 nmol/100 uL at ethanol precipitate 15 uL (~15 nmol), keep dry.
Brian and Fred’s ‘3D’ Fish protocol:
- (Plate cells on polylysine coated coverslips)
- (allow cells ~1hr to attach)
- (Fix cells 10′ in 5% PFA)
- (wash out PFA in PBT)
- Incubate for 10’ in 1X PBS + 0.5% v/v Triton X-100 at RT
- Incubate for 2’ in 1X PBS at RT
- Incubate for 30’ in 1X PBS + 20% v/v glycerol
- Immerse the slide in liquid nitrogen using forceps and allow to freeze completely (~15
seconds) - Allow the slide to thaw face-up on a paper towel
- Repeat steps 8 and 9 two additional times
- Incubate for 1’ in 1X PBS at RT
- Incubate for 5’ in 1X PBS at RT
- Incubate for 5’ in 1X PBS at RT
- Incubate for 5’ in 0.1 M HCl at RT (425 uL of 36% HCl in 50 mL = .1 M)
- Incubate for 1’ in 2X SSC at RT
- Incubate for 1’ in 2X SSC at RT
- Incubate for 1’ in 2X SSC at RT
- Incubate for 35’ in 2X SSC + 50% v/v formamide at RT
- Tap slides dry on a paper towel (never allowing the face of the slide containing the cells
to come into contact with the paper towel) - Create a hybridization cocktail consisting of 12.5 μl of 2X hyb. mix (4X SSCT + 20% w/v
dextran sulfate), 12.5 μl of formamide, and 20-40 pmole of each Oligopaints probe set
per slide. - Add the hybridization cocktail (typically 26-28 μl after adding the probe sets) to a 22×22
cm coverslip, invert the slide onto the coverslip, and seal with rubber cement - Allow the rubber cement to dry for 5’ at RT
- Denature the slides for 3’ at 78°C on a water-immersed heat block
- Hybridize overnight at 42°C in a humidified chamber
- The next day, remove the coverslip and wash for 15’ in 2X SSCT (2X SSC + 0.1% v/v
Tween 20) at 60°C - Incubate in 2X SSCT for 10’ at RT
- Incubate in 0.2X SSC for 10’ at RT
Seminar
- Prof Hans Blom presentation (Royal Inst. Tech in Sweden)
Intro
- single protein detection (~1970?)
- single fluor detect Los Alamos (1990)
receptor clusters
- sodium channels cluster in neurons
- dopamine and NKA receptors co-cluster in head of neural spine but not in other parts of spine or neuron. Observe differences in clustering and differences in inter-cluster difference.