3D Acrylamide FISH protocol
Detailed version 4

The Short 1 page version is HERE

Detailed protocol of 3D FISH methods described by Bass et al., (1997)

Acrylamide FISH for 3D Imaging of Chromsomes and Nuclei

The Acrylamide 3D FISH Protocol was developed for use with pollen mother cells (meiocytes) of maize as a way to study chromosome and telomere distributions within intact, formaldehyde-fixed nuclei as previously described (1). Fixed cells are embedded in a thin layer of optically clear acrylamide and buffers are exchanged by addition and aspiration of 200 microliter drops. The cells remain intact in the acrylamide throughout the entire procedure, including mounting and imaging.

This protocol was adapted from Sedat lab (UCSF) protocols for 3D high-resolution deconvolution microscopy of nuclear organization and chromatin structure. The protocol below was initially designed for higher plant meiocytes, but also works with cells from root tips, shoot meristems, and vibratome-sectioned leaf material, and probably other free-hand sectioned plant material. The protocol is a little tedious with many short steps, but provides excellent preservation of the integrity the nucleus, cell, and even tissue during FISH.

The protocol also works for immunocytochemistry (not described) and could be adapted for animal, yeast, or microbial cells too.



10X Buffer A salts, pH'd (Buffer A - - - )
     10X        -----------------------------------------
     150 mM     PIPES      ULTROL CALBIOCHEM, 528132L 
     800 mM     KCl        (reagent/mol biol grade)
     200 mM     NaCl          "
      20 mM     EDTA          "
       5 mM     EGTA          "
  bring to 90% of final volume, pH with 1M NaOH to 6.8, bring to 100% final vol,
  sterile filter, store aliquots at -20C or -80C in 50 ml polypropylene 
  conicals.  Stable for 1-2 yr.

1000X PA (polyamines, spermine and spermidine)
    0.15 M      SPERMINE     Spermine tetra HCL (Sigma S-2876)
    0.5  M      SPERMIDINE   Spermidine  (Calbiochem 56766)
  make 20 ML, sterile filter into microfuge tubes (0.5 mL each)
  store -80, stable for 1-2 yr.  LABEL AS "1000X PA, date"

1000X DTT (dithiothreotol)
    1.0  M      DTT          Calbiochem (233155)
    0.01 M      NaOAc        Sodium Acetate
  keep cool dissolve, sterile filter, aliquot to microfuge tubes 0.5 mL each, 
  store -80 for 1 yr. LABEL AS "1000X DTT, date"

30% Acrylamide Stock  (30:3.3 acrylamide:Bisacrylamide) 
  sterile filter, store 4C, stable for 6 months

1.6 M sorbitol, sterile filtered & stored -20 or -80.

mounting medium (I like Vectashield, Vetor Laboratories)

16% paraformaldehyde, sealed vials (EM grade, EM Sciences)

Ammonium Persulfate (for acrylamide gels)

Sodium Sulfite, anhydrous (used instead of TEMED for acry. gels)

POST-HYB WASHES - Made fresh from stock solutions.
Wash 1  4X SSC  (standard saline citrate)
Wash 2  2X SSC, 2X PBS (phosphate buffered saline)
Wash 3  1X PBS, 0.2% tween-20
Wash 4  1X PBS, 1mM DTT (= 1X DTT from above)
Wash 5  1X PBS
Wash 6  1X PBS, 10 ug/mL DAPI
Wash 7  1X PBS 1mM DTT

Aspirator  Side arm flask set up as vacuum trap.  At end of tubing,
  attach a 3 CC syringe (without stopper or needle). Use 200 uL
  pipet tips on end of syringe as disposable aspirator tips.


MBA (Meiocyte Buffer A) is made fresh from frozen stocks.
  MBA (a.k.a. Buffer A +++) =
    Buffer A salts, 1X
    sorbitol, 0.32 M (this may vary, isosmoticum for meiocytes)
    DTT, 1X
    PA, 1X


A1. Fix anthers in MBA plus 4% paraformaldehyde.
    Make fresh and add formaldehyde just before use.
    Fix for 2h in small vessel (small petri dish or 6-well dish)
    on a ratary shaker at RT.  Anthers should be swirlng slightly
    for good fixation.  They usually float on the surface.
    Use 4-6 mL for 20-40 anthers.
A2. Wash anthers by transfer to 5 mL MBA.
    Rotary shake 15 min., RT.  Repeat wash 3 more times.
A3. Use anthers immediatley or store in MBA at 4C.
Good for FISH for weeks or a few months if you're clean.


B1.  Wash slides (plain glass) with 70% ethanol until 
     squeaky clean using a Kimwipe.
B2.  Put on right kind of music (music from the hearts of space)
B3.  Remove dust/lint with blower or wipe with lens paper.
B4.  Label slide corner with a diamond marker.
B5.  Paint a ring of Sally Hansons Clear Hard as Nails nail 
     polish in middle of slide.  Usually 2 big drops.
     Leave a 2 x 1 cm open area for acryl pad.
B6.  Let sit to dry at room temp, with air blowing on it for 
     3-4 hours.  Can store in square petri dishes (Falcon 1012).
B7.  Prepare 22 x 30 x 1.5 coverslips by rubbing clean with 
     dry lens paper.  The no. 1.5 matches the optics of the
     deconvolution microscope.
 - make acrylamide mix and extrude meiocytes while slide rings dry

C)  MAKE 3X ACRYLAMIDE MIX (MBA, 15% Acryl) (C1-C6)

** acrylamide is a neurotoxin, (gloves & goggles) **

C1.  In a clean/new 15 mL conical, mix the following:
     5    mL    30% Acryl Stock (30:3.3) (to 15% final)
     1    mL   10X BufferA Salts (to 1X final)
     0.01 mL   1000X PA (to 1X final)
     0.01 mL   1000X DTT (to 1X final)
     2    mL   1.6 M Sorbitol (to 0.32 M final)
     1.98 mL	dei H2O
     10 mL final, vortex well.
C2.  Aliquot into 1.5 mL microfuge tubes, 0.5 mL/tube.
C3.  Degas for 1 minute in a speedvac (spin vac), RT, lids open.
C4.  Make 1ml 20% APS (ammonium persulfate) (0.2 g / 1 mL H20).
C5.  Make 1ml 20% NaS (Sodium Sulfite, anhydrous) ( same ).
C6.  Wash hands, have lunch.


D1.  Prepare a working surface by placing a circle of parafilm in
     the inverted lid of a standard plastic petri dish.
D2.  Place a 50 uL drop of MBA on this surface.
D3.  Add 5-8 fixed anthers for each slide.  
     Make sure the anthers and the meiocytes always stay submerged.
     Avoid dehydration (a lid over droplet reduces evaporation).
D4.  Work under a dissecting microscope.  To extrude the meiocyte
     column, hold an anther by one end with a forceps.
     Cut off the other end with a small sharp blade (Moria).
D5.  Hold the anther at the uncut end with the forceps, gently tap
     or squeeze out the column like toothpaste, using the blunt 
     side of the blade & moving it forward towards the cut open end.
     Watch the meiocyte column come out.  This takes some practice.
     Extrusion improves after 2 weeks of anther storage (really).
D6.  Once all the anthers are done - discard large non-meiocyte
     tissues/clumps before embedding in polyacrylamide.


E1  To transfer meiocytes from the petri dish to the slide, prepare 
    a 200 uL pipet tip by cutting off the end and then sucking up
    and expelling a solution of 20 ug/mL BSA in MBA.
    This will coat the plastic to prevent cells from sticking.
E2. While watching under the dissecting scope, suck up 13 ul of
    meiocytes.  Place them in the center of the nail polish ring 
    on the dry clean slide.
E3. Work fast and wear gloves from this point on:  
    To a tube containing 0.5 mL 3X Acryl. Mix, 
    add 25 uL 20% APS,  and 25 uL 20% Sulfite. Vortex briefly, 
    then quickly add 6.5 uL of this activated mix to the drop of 
     meiocytes on the slide. (acry will gel in tube in 20-30 sec).
E4. Stir with pipette tip just 2-3X around, then swirl mix on 
    the slide by hand, vigorously 20 sec.
E5. Mount a coverslip by placing the coverslip on the clean lab
    bench, turn the slide upside down, and pick up the coverslip. 
    -boink-  Ideally there will be no bubbles, and the coverslip
    will be nicely centered over the nail polish ring.
E6. Let acryl polymerize, coverslip side up, for 20 to 25 min., RT.
E7. Remove the coverslip by inserting a razor blade at a corner.
    There may be unpolymerized polyacrylamide at the edge because
    oxygen inhibits the reaction, but this is OK, just suck it up.
E8. The pad will stick to either the coverslip, or the slide.  
    If it sticks to the coverslip, prepare a new clean and labeled
    slide, add a small drop of water, and place the coverslip pad
    side up on the clean slide.  With a pipet tip, gently press down
    the coverslip, and suck off any excess water with an aspirator.
    Do not get bubbles under this coverslip directly under the pad.
    If the pad sticks to the slide, it is great.
E9. Aspirate off excess unpolymerized acrylamide.
E10. Add a drop of MBA to the slide, swirl, & aspirate.  Do this a
    few times to clean up the mess; results improve with more
    complete exchange of buffers.
E11. Add a drop of MBA and place on rotary shaker.  You can leave 
    them here until all the slides are prepared.  If pad wrinkles
    or lifts here, or at any other point, you can either ignore it,
    or correct it using a eyelash glued to a handle.


F1.  Prepare Prehyb. Buffer (50% good formamide, 2X SSC).
     Ten mL is enough for up to 8 slides, and 8 good slides is 
     4 slides too many (for deconv data collection, that is).
F2.  Aspirate off the solution very well from the slides.
F3.  Wipe the slides very clean with lens paper.
F4.  Add a drop of Prehyb buffer to the pad, rotary shake;  
     you may have to manually cover entire pad surface with buffer
     using pipette tip.
F5.  Aspirate and repeat for 2-3 changes of prehyb, 5 min each.


(Hyb. Buffer is probe in 50% good formamide, 2X SSC)

G1.  Dilute concentrated probe stock into Prehyb Buffer   
     Generally dilute to 1-3 ug/ml of fluorecently labeled oligo
     (or 0.13 uM) final concentration.
     [Note, Fluorescent oligo probes should be diluted to 
     500 or 100 ug/mL and stored in small aliquots (20-50 uL) 
     at -80 (amber microfuge tubes are a good idea).]
G2.  Dim the lights from now on to reduce photo-bleaching.
G3.  Apsirate & lenspaper remove the final prehyb.
G4.  Add 30 uL probe mix to the pad.  Use the tip to spread the
     liquid over the whole surface of the pad.  Let sit at room 
     temp at least 10-15 minutes.
G5.  Aspirate off probe mix.  Add another 30 ul probe mix to a 
     22 x 30 x 1.5 coverslip placed on the lab bench.  Mount on 
     the pad (boink).  Turn right side up and insure that the 
     coverslip is centered over the pad.  Air bubbles are OK, 
     but try to avoid them
G6.  Seal the coverslip on with rubber cement. (Heavy goopage is OK)
G7.  Place slide on a slide warmer at 30-40o C.  Slides can wait 
     here until they all catch up.  Wait at least until rubber 
     cement is pretty dry (about 5 minutes)
G8.  Pre-warm the containers (37C) that you will use to incubate the
     slides in overnight by placing them on the slide warmer 
     (avoid letting the slides drop below hyb temp after denaturation).
G9.  Heat denature by placing slides on a accurate heat block or PCR
     block set at 92 or 96 for 6 minutes.  A good way is to have
     duplicate slides, and denature one at 92, one at 96.
     [Note:  Do a temp series experiment from 80-100 for yourself.
     the best temp for hybs is a few degrees below whatever temp
     shows cell/nucleus/chromosomal disruption or damage.]
G10. Place in pre-warmed containers, store 37C dry incubator
     overnight (in dark).

H)  WASH and MOUNT (H1-H12)

H1.  Prepare wash and DAPI (wash6) solutions:
H2.  Remove slides from 37o C incubator.
H3.  Roll off the rubber cement.
H4.  Remove the coverslip by inserting a razor blade in the corner.
H5.  Wash and DAPI stain by adding appropriate solution, 
     placing slides in a foil-covered box on a rotary shaker, RT,
     for about 5 minutes per wash, remove by aspiration.
  WASH 1.  3 times
  WASH 2.  3 times
  WASH 3   3 times
  WASH 4.  3 times
  WASH 5.  3 times
  WASH 6.  2 times, second time for 20 min, RT.
  WASH 7.  3 times.
     Aspirate the last wash very well,
     clean and dry the slide with lens paper.
H6.  Mount in Vectashield (Vector labs, Burlingame, CA  H-1000):
H7.  Add 100 ul Vectasheild, swirl around, aspirate, repeat 2 times.
     (aspiration is slower, tilt slide to help excess run down)
H8.  Clean a coverslip with lens paper, and blow dry with dust free
     and static free blower.  
     Place on a clean piece of lens paper on the bench.
H9.  Mount by placing Vectashield on the clean coverslip, 
     (30 uL for 22x30 or 20 uL for 22x22 coverslip).  
     Invert the slide, and mount coverslip upside down. (-boink-).
H10. Gently push down coverslip with new clean blunt end of 1 mL 
     blue tip, no harder than a pushing a tack on your skin 
     (helps to flatten out wrinkles in the acrylamide).
H11. Seal with nail-polish (Sally Hansons Clear Hard as Nails).  
H12. Let air dry, image within 24h or store at -80.


Thanks to Lisa Harper for the writing the first version of the detailed 3D Acrylamide FISH protocol.

(1) Bass HW, Marshall, WF, Sedat JW, Agard DA, and Cande WZ (1997) Telomeres cluster de novo before the initiation of synapsis; a 3-dimensional spatial analysis of telomere positions before and during meiotic prophase J. Cell Biol. 137(1):5-18.

3D images from this technique.