Immunofluorescence

Stock Solutions
10X Egg Buffer   NPG-Glycerol  Solutions made fresh the day of use: 
 * 250 mM HEPES pH 7.4
 * 1.18 M NaCl
 * 480 mM KCl
 * 20 mM EDTA
 * 5 mM EGTA
 * Filter sterilize and store at room temperature
 * 2 g N-propyl gallate (Sigma)
 * 50 g Glycerol (easier to weigh than to measure by volume)
 * Place on nutator and dissolve by agitating overnight, and store at room temperature.
 * Methanol (-20°)
 * 2 M TRIS-base (not pH’ed)

EBT (1 ml) optional: (these are found in the -20° and contribute to the maintenance of chromosome structure during IF; if you use these, reduce the water to 848 µl)
 * 110 µl 10X Egg Buffer
 * 10 µl 10% Tween
 * 30 µl 0.5 M NaAzide
 * 850 µl milliQ water
 * 1 µl .5M spermine
 * 1 µl .15M spermidine

 Fix (1 ml) 

for 2% formaldehyde (1% formaldehyde final) for 7.4% formaldehyde (3.7% formaldehyde final) (i.e. DAPI staining 1X PBST (1 L) Mounting Medium (0.5 ml) Mix this solution using a P-1000 tip that has been cut with a razor blade, vortex to mix further, then spin for 1-2 minutes at top speed to remove bubbles.
 * 100 µl 10X Egg Buffer
 * 54 µl 37% Formaldehyde
 * 846 µl milliQ water
 * 100 µl 10X Egg Buffer
 * 200 µl 37% Formaldehyde
 * 700 µl milliQ water
 * 100ml 10X PBS
 * 900ml milliQ water
 * 1ml Tween 20
 * 35 µl 2M TRIS (not pH-adjusted)
 * 15 µl milliQ water
 * 450 µl NPG-glycerol

Protocol
Before you start: make all your “day of” solutions and place a metal block on dry ice so it starts cooling.

1. Dissect age-matched adults (typically 18-24 hours post-L4). Pick worms into a 30µl drop of EBT on a No. 1 (18mm2 ) coverslip on top of a glass slide. Use a scalpel blade (we use Feather brand #11) to cut the heads and/or the tails off of the worms to extrude the gonad.

2. Pipet 30µl of Fix solution into the drop of dissected worms. Pipetting up and down a few times may help release more gonads. Let worms fix for 4-5 minutes.

3. Pipet off excess liquid leaving around 15µl remaining. Pick up the drop by lightly touching a HistoBond (75x25x1mm from Lamb) microscope slide to the top of it. Wick away excess liquid from the edges of the coverslip using a torn piece of Whatman paper. The more liquid you remove, the better to worms will stick to the slide.

4. Freeze crack samples. Freeze the sample by placing it on the ice block for at least 30 seconds. If you are dissecting multiple slides, you can leave your slides on the ice block indefinitely (not really, just until all your slides are dissected; this is not a point in the protocol where you can stop for several hours). Repeat steps 2 through 5 until all samples are dissected.

5. Carefully flick off the cover slip by catching the edge with a fresh razor blade.

6. Place the slide immediately in -20° Methanol for 1 min, then move to a Coplin jar of PBST at room temperature.

7. Wash slides 3x (10 min/wash) by moving slides to fresh Coplin jars of PBST.

8. Block slides by moving them to Coplin jar with PBST plus block (0.5% BSA in PBST with .02% azide; stored in 4° refrigerator). Incubate at room temperature for 30 minutes.

9. Primary Antibody: Apply 50-75µl of 1° antibody diluted in PBST plus block onto a Parafilm coverslip, invert your slide and touch it to the 1° antibody dilution. Invert slide again, picking up coverslip, and incubate in a humid chamber for 2 hours at room temperature or overnight at 4°C.

10. Remove the Parafilm coverslip by placing each slide in a Coplin jar of PBST and letting it float off (this is easiest when the level of PBST is higher than the top of the slides). Remove the coverslips and throw them away. Wash slides a total of 3x (10 min/wash) by moving slides to fresh Coplin jars of PBST.

11. Secondary Antibody: Apply 50-75µl of 2° antibody diluted in PBST onto a Parafilm coverslip, invert your slide and touch it to the 2° antibody dilution. Invert slide again, picking up coverslip, and incubate in a humid chamber for 2 hours at room temperature in dark.

12. Wash and DAPI stain samples. Remove the Parafilm coverslip as in step 11. Wash slides 10 min in a fresh Coplin jar of PBST, then move to Coplin jar of PBST plus 0.5µg/ml DAPI (add 5 µl of 5mg/ml stock to 50ml PBST in a Coplin jar) and incubate for >10min. Finally, wash slides >15 min in a fresh Coplin jar of PBST.

13. Mount slides. Pipet 10-15µl of Mounting Medium onto a clean No. 1 ½ (22mm2 ) coverslip. Remove the slide from buffer, remove as much buffer as possible without dessicating the tissue sample by wiping with a kim wipe and/or aspirating. BE CAREFUL NOT TO WIPE OFF YOUR WORMS!!! Invert the slide and touch it to the mounting medium. Invert the slide again, and carefully aspirate off any excess mounting medium. Seal with clear nail polish (we use Sally Hansen Hard-as-Nails).

Additional Information
DAPI staining of meiotic nuclei:
 * 1° antibodies are stored at -20° in 50% glycerol. Most of the antibodies we use are still in serum and can be used at 1:250. Additional aliquots of most antibodies are stored at - 80°. If you need to pull out an antibody from the -80°, let it thaw on ice and dilute it with an equivalent volume of 100% glycerol (for a final concentration of 50%) before use
 * 2° antibodies are stored at 4° and can be used at 1:500. Additional aliquots are stored at -80° and should be thawed on ice before use. When determining what 2° antibodies to use, keep in mind the animal in which the secondary was generated (most of ours are from donkey and goat) and whether an cross reactivity has been minimized by incubation with serums from other animals. This information can be found on the information sheets of these antibodies in the maroon binder or online at http://igene.invitrogen.com/antibody/filter.do (for Alexa Fluor 488 conjugated antibodies) and http://www.jacksonimmuno.com/Catalog/wholeigg.asp (for Cy3 and Cy5 conjugated antibodies)

To only visualize meiotic chromosomes, i.e. for counting bivalents in diakinesis, one should age the animals so diakinesis nuclei are maximized (48 hours at 20° or 72 hours at 15° post L4). Dissect these animals and fix in 3.7% formaldehyde for five minutes, following the protocol until step #6. Skip to step #12 (DAPI staining) and complete the protocol.

To test a new antibody, two parameters should be tested:

1. fixation conditions for most of our antibodies, a 5 minute fix in 1% formaldehyde works. However, you may need to vary formaldehyde concentrations for your antibody of interest. In addition, some antibodies (i.e. Upstate’s dimethyl-Lys9-histone H3) work better on tissue that has not been fixed with formaldehyde. In this case, you would dissect your animals, skip step #3 (forgoing the addition of fix) and continue the rest of the protocol. Your meiotic nuclei might not be so pretty, but you may be able to see where your protein is localized. Another alternative fix protocol is to skip formaldehyde fixation, freeze crack your samples and incubate in -20° Methanol for 10 minutes, then -20° Acetone for 10 minutes, let them dry on your benchtop and then move onto the incubation in block.

2. antibody concentration typically when testing a new antibody, you vary antibody concentration ten fold, so the first time you perform IF you should have 3 dilutions: 1:50, 1:500 and 1:5000. If the antibody is affinity purified (and therefore at much higher concentration than in serum), you may need to go even higher (i.e 1:10,000 and 1:20,000 dilutions are common).