Immunostaining

Phalloidin is a stain that allows for the visualization of F-actin, allowing us to visualize muscle structure. α-BTX and SV2 label acetylcholine receptors and synaptic vesicles, respectively, allowing for visualization of neuromuscular junctions (NMJs). The following protocol allows for the visualization of muscle structure and neuromuscular junctions in zebrafish embryos and larvae using phalloidin, α-BTX, and SV2. This protocol is optimized for the staining of 1-10 embryos per tube at 8 days post-fertilization or younger.

Procedure 

Note: When rocking or storing embryos, always place tubes horizontally (on their side) for best results. Tubes should be placed perpendicular to the rocking motion, such that the embryos rock from side to side in the tube as opposed to rocking from cap to bottom. 

Note: A pipette pump and glass pipette tip can be used for addition and removal of all solutions, except for addition of phalloidin and primary and secondary antibody solutions. 

Phalloidin-only Staining

1. Fix fish in 4% paraformaldehyde (PFA) in microcentrifuge tubes. Add enough liquid so embryos are covered (~0.5 ml). Between 1-10 embryos can be placed per tube. Label tubes accordingly.
   1. Embryos can be rocked in PFA at room temperature (with tubes on their side) for 4 hours or stored on their side in a 4°C fridge overnight.
2. Remove 4% PFA from tubes containing the embryos and dispose it in the appropriate waste container. Rinse embryos 5 times for 5 minutes each in PBS-0.1% Tween 20® (PBS-Tw; Bio-Rad, Hercules, CA).
   1. Add ~0.5 ml PBS-Tw to embryo tubes, rock tubes on their side for 5 minutes, then remove and discard PBS-Tw. Repeat 5X.  Gently pipetting the embryos up and down with a glass pipette tip in the tube with each solution change can aid in washing and removal of solutions. This can also prevent embryos from sticking together if >1 embryo is in each tube. Remove as much liquid as possible from the microcentrifuge tube after the last wash.
3. Add ~0.5 ml PBS-2% Triton-X-100® (Fisher Scientific, Waltham, MA) to each tube (enough solution to cover the embryo). Rock at room temperature on their side for 1.5 hours. 
4. Remove PBS-2% Triton-X-100® from tubes and add 1:20 dilution of Alexa Fluor phalloidin (488 or 546) in PBS-Tw (10 µl volume per tube). Rock for 4 hours at room temperature or store at 4°C overnight on their side completely covered (in the dark). From this point on all steps must occur in the dark with tubes covered (e.g. under a box lid or wrapped in aluminum foil).
   1. If there is a large number of tubes, a master mix can be made with 1 µl phalloidin for every 19 µl of PBS-Tw. Then 10 µl master mix can be added per tube.
   2. If there is a small number of tubes, 19 µl PBS-Tw can be added per tube. Then, 1 µl phalloidin can be added to each tube.
5. Remove the phalloidin/PBS-Tw mix, and rinse embryos 5X for 5 minutes with PBS-Tw. Store embryos in ~0.5 ml PBS-Tw  at 4°C until imaging can occur.

Phalloidin and NMJ stain

1. Fix fish in 4% paraformaldehyde (PFA) in microcentrifuge tubes. Add enough liquid so embryos are covered (~0.5 ml). Between 1-10 embryos can be placed per tube. Label tubes accordingly.
   1. Embryos can be rocked in PFA at room temperature (with tubes on their side) for 4 hours or stored on their side in a 4°C fridge overnight.
2. Remove 4% PFA from tubes containing the embryos and dispose it in the appropriate waste container. Rinse embryos 5 times for 5 minutes each in PBS-0.1% Tween 20® (PBS-Tw; Bio-Rad, Hercules, CA).
   1. Add ~0.5 ml PBS-Tw to embryo tubes, rock tubes on their side for 5 minutes, then remove and discard PBS-Tw. Repeat 5X.  Gently pipetting the embryos up and down with a glass pipette tip in the tube with each solution change can aid in washing and removal of solutions. This can also prevent embryos from sticking together if >1 embryo is in each tube. Remove as much liquid as possible from the microcentrifuge tube after the last wash.
3. Add 125 µl of 1 mg/ml collagenase in 1X PBS per tube and rock on their side for 1.5 hours at room temperature for permeabilization. 
4. Remove collagenase from the tubes containing the embryos. Add a 1:500 dilution of α-BTX-647 and 1:20 dilution of phalloidin in antibody block (Ab block; 5% BSA [Fisher Scientific], 1% DMSO [Sigma-Aldrich], 1% Triton-X-100, 0.2% saponin from quillaja bark [Sigma-Aldrich] in 1× PBS). Add 50 µl of master mix per tube. Cover tubes and rock on their side at room temperature for 2 hours with the tubes covered. From this point on all steps must occur in the dark with tubes covered (e.g. under a box lid or wrapped in aluminum foil).
   1. Example master mix dilution:
      1.  1 μl α-BTX + 25 µl phalloidin 546 + 474 μl Ab block 
5. Remove the α-BTX and phalloidin mix from tubes. Rinse embryos 5X with PBS-Tw, rocking them on their side for 5 minutes between rinses (~0.5 ml PBS-Tw per rinse).
6. Remove PBS-Tw and add ~0.5 ml Ab block to each tube so embryos are covered with solution. Incubate covered tubes on their side overnight at 4°C.
7. Remove Ab block. Add SV2 antibody in 1:50 dilution in Ab block (25 µl volume master mix per tube). Rock for 6-8 hours on their side at room temperature (covered under a box lid or wrapped in aluminum foil), then incubate at 4°C for 48 hours. 
   1. Example master mix dilution:
      1. 1 μl SV2 + 49 μl Ab block 
8. Remove the SV2 antibody mix, and rinse embryos 5X for 5 minutes in PBS-Tw. Add ~0.5 ml Ab block to each tube and rock on their side at room temperature for 8 hours. 
9. Add 1:500 dilution of secondary antibody (GAM) in Ab block and incubate on their side at 4°C overnight (50 µl of master mix to each tube)
   1. Example dilution:
      1. 1 µl GAM + 499 µl Ab block
10. Remove secondary antibody from embryos by rinsing tube 5X for 5 minutes each with PBS-Tw. Then fill tubes with ~0.5 ml PBS-Tw and store at 4°C until imaging.

Imaging should occur within 2 weeks of immunostaining for best results (the sooner the embryos are imaged, the better the results). Phalloidin-488 or -546 and GAM-488 or -546 were used interchangeably with no differences in staining observed.

Imaging

1. Once embryo staining is completed, deyolk the embryos.
   1. Place the fixed embryo in 1X PBS, and use deyolking tools (e.g. insect pins super glued to the ends of glass capillary tubes) to gently remove the yolk sac from the zebrafish embryo while looking through a dissecting microscope.
2. Mount the deyolked embryos in a glass-bottom plate (e.g. 24-well glass bottom plate) using 0.5% low-melt agarose (Boston BioProducts, Ashland, MA) in 1X PBS. 
   1. Add 0.5 g Agarose low gelling temperature (Boston BioProducts, NC9731636) in 10 mL 1X PBS. Heat the solution in a 50 ml conical flask in a microwave. 
   2. Allow agarose to slightly cool (such that when it is taken up by the pipette, the pipette tip does not fog up), and pipette agarose to coat the bottom of the glass-bottomed plate. 
   3. Add your embryo to the agarose in the glass dish. Use a poker (e.g. fishing line super-glued to a glass capillary tube) to adjust and arrange the embryo as needed. 
   4. Zebrafish were mounted anterior left and dorsal up to ensure the same side of each embryo was imaged. 
3. Fluorescent images were captured using a 25X water objective. Appropriate lasers for each respective stain were used. Imaging focused on the 12^th myotome of each embryo. A system optimized z-stack (z-step size of between 0.57- 0.8) was obtained for each embryo, ranging from the bottom surface of the embryo (closest to the glass bottom of the dish) to the notochord (about half-way through the depth of the embryo body). The same imaging parameters were used for each embryo, for consistency.
   1. For phalloidin-only imaging, a format of 1024 x 1024 pixels, scanning speed of 200 Hz, and line average of 2 works well.
   2. For phalloidin and NMJ imaging, a format of 4096 x 4096 pixels, scanning speed of 200 Hz, and line average of 2 yields the best results. 

All fluorescent images were captured using a Leica SP8 confocal microscope.

Materials and Equipment:

• 1.5-2 ml microcentrifuge tubes
• 4% paraformaldehyde (PFA)
• PBS-0.1% Tween 20®
• PBS-2% Triton-X-100®
• antibody block (Ab block; 5% BSA [Fisher Scientific], 1% DMSO [Sigma-Aldrich], 1% Triton-X-100, 0.2% saponin from quillaja bark [Sigma-Aldrich] in 1× PBS)
• 1 mg/ml collagenase (diluted in 1X PBS)
• Alexa Fluor Phalloidin-488 or -546 (Invitrogen, Eugene, OR)
• alpha-bungarotoxin-647 (Invitrogen)
• SV2 (DSHB, Iowa City, IA)
• Alexa Fluor GAM-488 or -546 secondary antibody
• Lab rocker
• Glass Pasteur pipettes
• Pipette pump 
• Micropipettes
• Micropipette tips
• Agarose low gelling temperature (Boston BioProducts, NC9731636)
• Glass bottomed dish (e.g. 24-well glass bottomed plate)

Reagent Recipes

• Phosphate buffered saline (PBS)
  • 10x PBS
    • To a 1 L bottle, add:
      • 74 g NaCl
      • 19.4 g Na₂HPO₄∙7H₂O
      • 4.37 g NaH₂PO₄∙H₂O
      • ~800 ml dH₂O
    • Stir until dissolved. Add dH₂O to bring volume up to 1 L. Autoclave the solution.
  • Dilute 10x PBS to 2x PBS or 1x PBS with dH₂O. Resulting solutions can be stored at room temperature.
  • PBS 0.1% Tween-20
    • Add 1 ml of Tween-20® to 1 L of 1x PBS, and mix. Store at room temperature.
  • PBS 2% Triton X-100®
    • Add 20 ml Triton X-100® to 1L of 1x PBS and mix the solution. Store at room temperature. 
• 4% Paraformaldehyde (PFA)
  • Add the following to a 50 ml conical tube:
    • 2 g PFA
    • 15 ml dH₂0
    • 10 drops of 1M NaOH
  • Dissolve PFA into solution by placing it into a water bath (a beaker filled with water) with a stir bar on a hot plate.
  • Add dH₂0 to fill the conical tube up to the 25 ml line.
  • Filter the solution into a clean conical tube using filter paper and a funnel.
  • Add 12 drops of 1 M HCl to the conical tube, put the lid on and mix the solution.
  • Add 25 ml 2x PBS to the conical tube. 
  • Check the pH so the solution is between 7.2 and 7.4. If the pH is not within this range add drops of 1 M HCl or 1 M NaOH as needed. Store the resulting solution at 4°C and use the solution within 1 week of making it. 
• Antibody (Ab) block
  • To a 50 ml conical tube, add:
    • 2.5 g bovine serum albumin (BSA)
    • 40 ml 1x PBS
  • Gently heat the tube and rock until BSA is dissolved (e.g. hold the tube under a running faucet with hot water)
  • Add the following to the conical tube:
    • 0.5 ml DMSO
    • 0.5 ml Triton X-100®
    • 0.1 g saponin
    • Add 1x PBS until the resulting volume in the tube is 50 ml.
  • Store at 4°C and use the solution within 1 week of making it.