recipe). Confluent HeLas, for example, will grow onto coverslips

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recipe). Confluent HeLas, for example, will grow onto coverslips Powered By Docstoc
					Sam’s general immunofluorescence advice & protocol (updated May 2007)

   1- 1-3 days before staining, plate cells on acid-washed coverslips (see
      recipe). Confluent HeLas, for example, will grow onto coverslips up to
      about 300,000 cells per small (10 mm) coverslip.

For stem cells:

Matrigel 1:500 in media (plain DMEM works fine), incubate overnight at 4
degrees, then overnight at 37 degrees, to coat the coverslips. Then wash twice
with PBS and twice with stem cell media before using.


Coat coverslips with gelatin, then plate feeder layer (MEFs or HS-27 foreskin
fibroblasts) the day before.

   2- Fixation is key. Here are the fixations I commonly test when I’m using a
      new antibody:

   • 4% formaldehyde in PBS (see recipe) for 10 minutes at room temp
   • 1% formaldehyde in PBS for 15 minutes
   • 1% formaldehyde in PBS-Tx for 10 minutes
   • 100% high-purity (HPLC grade) –20°C methanol for 10 minutes at -20°C

   Other variations include: pre-extracting with various buffers, adding inhibitors
   (such as phosphatase inhibitors to the fixative solution). In some cases it is
   necessary to combine formaldehyde and methanol fixations when using
   multiple antibodies. I always do the formaldehyde fix first, then do the
   methanol fix IMMEDIATELY after (don't rinse in between and don't let the
   coverslips dry out!).

   For example, to get good DNA AND spindle morphology, I do a 2-step fixation:

   Do NOT rinse cells prior to fixing if you’re looking for mitotic cells, as they tend
   to detach very easily. Just remove media and add the fixative.
   1% formaldehyde in PBS-Tx 15’ room temp, take off and IMMEDIATELY add
   100% cold (-20) HPLC-grade methanol, IMMMEDIATELY transfer plate to -20
   for 10 minutes, NO MORE NO LESS.
   Take off methanol and IMMEDIATELY add PBS. Be very gentle with pipetting,
   because the cold liquid will blast your cells off the coverslips, but move fast
   with methanol, or your coverslips will dry out. Proceed with primary antibody
   as usual.
Formaldehyde has to be fresh (see recipe). Do NOT overfix! AT NO TIME

Also, I NEVER fix and store samples prior to staining. I ALWAYS fix and stain
the same day. I DO NOT recommend storing samples at 4 degrees after

There’s no need to block prior to adding the primary antibody, because
you should make your primary antibody dilutions in blocking solutions!
The only time you need an extra blocking step is if you’re doing something
fancy (see separate double-labeling protocol, e.g. two rabbit or two mouse
antibodies, if needed).

Step 1, prepare cells for fixing:
Take media off of cells by aspirating gently. Cover with PBS (see recipe; this
rinse step is optional). If extracting, extract now (shouldn’t be necessary for
most proteins). Otherwise, go on to fixation.

Step 2, Fixing:
Remove PBS (or media if you skipped this) by aspirating gently and add
fixative for the appropriate amount of time.

Step 3:
Wash with PBS once and PBS-TX (PBS + 0.1% Triton x-100) once.

Step 4.
Transfer coverslips to incubation chamber (mine is a box with a wet sponge,
with a piece of glass on top, and a piece of parafilm on top of that).

Step 5, add antibody:
10 mm coverslips need ~ 25 ul of antibody mix to cover them. I always make
extra b/c you need to centrifuge it (10 minutes on top speed) to get rid of any

Make up the antibody at various dilutions in blocking solution. Different
blocking solutions may be appropriate depending on your application. You
may want to try: Ig-free BSA (1-5%) in PBS-Tx, or 5-10% serum in PBS-Tx.

The source of the serum matters- it has to be heat-inactivated, and should
match the source of your secondary antibody, but I’ve found that fetal calf
serum (the kind we use for tissue culture) works just as well as almost any
other kind, and it’s cheaper. Under no circumstances should your serum
match the animal your primary originated in, unless you want a negative
Sterile-filter (that's 0.22 micron filters) your blocking solution and store
at 4 degrees. If you find you start getting background suddenly one day, it’s
probably because your blocking solution has gone bad (gets contaminated
with bacteria, etc). I make mine in 10 ml batches so that I can be sure to use
it up before it goes bad.

Primary Antibody Dilutions:
Dilutions matter, just like for western blots. A good rule of thumb:
- for affinity-purified peptide antibody fractions, start around 1:25.
- For commercial antibodies, start around 1:50-1:100.
- For high-titre antibodies, like human autoimmune sera, start around
I’ve never met a good primary antibody that worked well diluted more than
1:5000 at most.
My default trials for a first-run (non-peptide) antibody are usually 1:50, 1:100,
1:500, and 1:1000. This is an important parameter and it’s worth your time to
test it early with new antibodies.
For 10 mm coverslips, I use 25 ul of antibody mix. Always make a little extra!
Centrifuge sufficient diluted antibody in blocking solution for 10
minutes on high (14,000 rpms) in a tabletop microcentrifuge. DO NOT
SKIP THIS STEP if you want your staining to be clean!

Step 6, incubate in primary:

I do my incubations at 37 degrees for 30 minutes. You shouldn’t need to
go longer at this temperature, kinetics is on your side! If the antibody solution
is evaporating off your coverslips, your box isn’t sealed well enough or your
sponge isn’t wet enough. Some people incubate overnight at 4 degrees, but
this can increase background (nonspecific binding) since it is less stringent. I
have never found it to be useful.

Step 7, wash:

I use a set of ceramic coverslip holders from Coors, USA (Thomas Scientific,
part # 8542E40). They’re about $100 each, and each holds 12 coverslips. I
put the holder in a small Tupperware or beaker and cover it with PBS-Tx just
enough to have the coverslips covered, THEN add the coverslips to the
holder. Do this gently, or you’ll wash your cells off just from breaking the
surface tension of the buffer.

Wash 2x in PBS-Tx, for 5-10 minutes each, with VERY GENTLE agitation
on an orbital shaker. Just enough to keep the buffer circulating, not enough to
make the coverslips bang back and forth against each other.
Step 8, add your secondary:

Make it up in blocking solution and centrifuge for 10 minutes.

Seocndaries for deconvolution: I only use pre-absorbed secondaries from
Jackson Immunological, the multilabeled variety (ML). I do this because
they’re the cleanest ones available (last time we checked). I use my
secondaries at 1:50-1:100.

Secondaries for confocal: I use highly cross-absorbed Alexa secondaries from
Molecular Probes (now Invitrogen) because they’re the brightest and the most
stable. They’re not remotely as clean as the Jackson antibodies, especially on
the human stem cells. AVOID the anti-mouse 546, it seems to have really
severe nonspecific cross-reaction.

At this point, you can add your DAPI (1 mg/ml in water, store at –20°C
indefinitely, use at 1:100-1:1000 depending on cell type) or Hoechst or ToPro
or other DNA stain. Just remember: don’t use a green secondary if you’re
looking at GFP also!

Incubate 30 minutes at 37 degrees, as for primary.

Step 9, wash again.

This time, do one 10-minute wash in PBS-Tx, then one 10-minute wash
in plain PBS to get rid of the detergent. Optional: wash 1 minute in distilled
water to get rid of PBS.

Step 10, mounting:

Air dry your coverslips if you want to. One advantage to this is that the
coverslips will last longer; the drawback is if you want to look at 3D structure,
the cells sometimes collapse if you let them dry too long & get flattened out.

Mount using 2 µl Slowfade per 10 mm coverslip (glycerol & water-based
antifade reagent from Molecular Probes). You'll need more for thick stem cell
growth (up to 10 ul for EBs).

Note: I don’t like Vectashield. ProLong is good if you’re using Alexa or Cy-
labeled secondaries, but it cures to a hard polymer, so keep that in mind if
there’s any chance you’ll want to un-mount and restain! Some mounting
media will work better for red and not at all for blue or green channels, so I
recommend finding one that works for your secondaries.

Seal edges with a 1:1 mix of plain (e.g. Wet ‘n Wild, $1.00) clear nailpolish
Modifications for Cellomics (multiwell dishes with no coverslips):
1. Make more antibody mix than you would for coverslips.
E.g. for each 10 mm coverslip, I make 25 ul of antibody mix.
Even though we grow 10 mm coverslips in 24 well dishes without much extra
space, you'll need 75-100 ul of antibody mix to cover the wells so they don't
dry out during staining.

2. Add a post-fix step with 2% formaldehyde for 5 minutes at room temp.
Remove post-fix and store in PBS + 0.1% azide.
Cover with sticky clear tape (from PCR machine), the lid, and aluminum foil.
Best to image the same day you stain, because the plates do grow bacteria
when stored at 4 degrees for a long time (and the bacteria will eat your
epitopes and stains!)


10x PBS recipe (from Current Protocols in Molecular Biology- Red Book)
10x ingredients                       1x
80 g NaCl                             137 mM
2 g KCl                               2.7 mM
11.5 g Na2HPO4•7H2O            43 mM
2g KH2PO4                             1.4 mM
1 L H2O

For PBS-Tx, make a 20% stock of Triton-X-100 in PBS, then add 5 ml of that per 1
litre of PBS. Filtering this will take a long time, so I usually don’t: I sterile-filter my 1x
PBS and PBS-Tx because it seems to help a lot with avoiding background signal.

8% formaldehyde stock for fixation (dilute before using):

Add solid PFA to 80% in ddH2O (start with 80g in 700 ml ddH2O)
Heat to 95°C in the hood for 1 hour- do NOT let it boil!
Cool and bring up to 800 ml.
Add 10 N NaOH dropwise until solution clears (~300 ul per liter)
Add 10x PBS to a final concentration of 2x (200 ml)
Sterile-filter & store in aliquots at -20°C indefinitely.

To use: Melt at 55-65 degrees for ~ 10 minutes or until clear. Then add an equal
volume of water to give 4% formaldehyde in 1x PBS. Further dilutions should
be made in 1x PBS or PBS-Tx.
   Acid-washed coverslips:

    For some cell types, simply autoclaving and washing with ethanol can be sufficient.
However, coverslips typically have chemical residues on them leftover from
manufacturing, which can interfere with cell attachment and growth. Acid-washing also
seems to help give the coverslips some ‘charge’ so the cells can attach. Finally, for high-
resolution imaging, you want to get rid of any dust- autoclaving alone won’t help with

   Also note that, for imaging analysis purposes, the best ones to use are No. 1.5 (~
   0.17mm thick) coverslips, not No. 1.0 (thinner) coverslips. The bonus is that the
   thicker ones are easier to handle.

   1- In the chemical hood, wash by incubating in concentrated HCl 1-2 hours. Pour
      this directly into a large beaker, DO NOT use plastic pipettes! You only need
      enough to cover the coverslips, but you should be able to swirl them around
      without having them dry out.
   2- wash with ddH2O 5 times
   3- boil in dH20 15 minutes (in the hood!)
   4- wash again with ddH2O 5 times
   5- wash 3-4 times with 100% ethanol
   6- separate and dry on kimwipes in a TC hood (otherwise they stick together), and
      store in a sterile Petri dish or tissue culture dish.

   I have found this method to be sufficiently sterile for most applications, but if you're
   growing cells with no antibiotic, it's probably a good idea to use fresh ethanol and/or
   flame them before using.