Vanessa cardui embryo fixation and in situ protocol, Katie Reding
Embryo collection
When the sunflower plant is placed in the butterfly cage, Vanessa tend to lay embryos on and near the plant, including on the soil surface. To avoid having embryos laid in hard to reach places, we put a piece of paper around the base of the sunflower plant. Fold a piece of paper in half lengthwise, and then again. Cut a small hole in the folded corner, and cut around the edges to make a circle. Unfold, and cut a line from the outside of the circle to the small hole in the center. Place the paper around the stem and overlap the two ends of the circle and tape together once you have created a cone around the stem.
Place the sunflower plant in the butterfly cage for as long as you want to collect embryos. Carefully remove all embryos from the sunflower, allowing them to fall to a collection tray below. Age the embryos as desired.
Embryo fixation
1.Heat fix
Place the embryos (no more than 100 uL of embryos) in a 1.5 mL tube, add about 500 uL of water. Put the embryos in a boiling water bath for 3 min, then submerge the tubes in ice, and leave for 10 min. In addition to partially fixing the embryos, during the heating step, the chorions will inflate which will create space between them and the embryos, making them much easier to remove.
2.Chorion removal
Eggshells can then be removed using one of two methods: 1) Transfer embryos to PBST and to a glass dissection dish, and remove eggshells manually; 2) Transfer embryos to a Drosophila-style embryo collection basket and submerge in 1:1 bleach:water for about 1 min, then rinse several times in tap water. The first method is more labor- and time-intensive, but is sure to work, while the second method can result in embryos disintegrated if they are left in bleach too long. The amount of time to leave the embryos in the diluted bleach may need to be determined empirically given that commercial bleach products vary in their percentages of sodium hypochlorite. If using the second method, rinse the embryos several times in PBST before proceeding to the PFA fix.
Note: If you are staining germband-stage embryos, you can also go a step further and dissect the germbands out of the yolk at this point, or you can do this later (after storage in methanol).
3.Fixation with paraformaldehyde
Thaw a fresh tube of 12% paraformaldehyde (PFA). Remove PBST from each tube of embryos and add ~1 mL of 4% PFA in PBST. Rock embryos for 20 min. Remove the PFA, and rinse the embryos 3X with PBST. Gradually transfer the embryos to methanol. We do this by removing 300 uL of PBST, adding 300 uL of methanol, allowing embryos to rock a few min, then repeating this several more times until the solution is almost entirely methanol. Then rinse the embryos 3X with 100% methanol and finally store the embryos at -20C in methanol.
4.Dissection of germband-stage embryos
If you haven’t already done so, any germband-stage embryos should be dissected to remove the germband from the yolk. You can do this any day after fixation and before you want to start the in situ. Simply transfer the embryos from methanol to PBST (using the gradual transfer method described above), move to a glass dissection dish, and dissect. Remove the yolk to leave only the germbands. Gradually transfer back to methanol and store at -20C until you’re ready to begin the in situ hybridization. At many stages the germband is concave, almost bowl-shaped at the posterior end, and it can be very difficult to remove the yolk from this area without damaging the embryo. However, you do not need to remove all bits of yolk at this time; you just need to remove the yolk that is enveloping the embryo.
In situ hybridization
Gradually transfer embryos from methanol to PBST. Remove PBST and add 4% PFA; rock for 1.5 h. Rinse several times with PBST. Add about 1 mL hybridization buffer and rock until embryos sink. Incubate at 60C for 4 h. Add probe diluted in hybridization buffer and hybridize at 60C overnight.
The next day, put 2 mL of hybridization buffer and 1 mL 2X SSC in the hybridization oven to warm up. Remove probe and perform the following washes:
Buffer Temperature (C) Time
Hyb buffer (pre-warmed) 60 30 min
Hyb buffer (pre-warmed) 60 30 min
2X SSC (pre-warmed) 60 30 min
2X SSC RT 30 min
0.2X SSC RT 30 min
Rinse the embryos 3X with PBST. Wash for 2 h at RT with 10% sheep serum. Add the anti-dig-AP antibody diluted 1:1600 in 10% sheep serum; rock for 1.5 h at RT. Remove the antibody, and rinse with PBST 3X. Leave to rock in PBST overnight at 4C.
The next day, wash 3 X 20 min in PBST, then 3 X 5 min in AP staining buffer. Remove all buffer and add 500 uL of NBT/BCIP. Place tube in the dark and check progress of stain occasionally (after 3 h or so). Time for stain development needs to be determined empirically. Once embryos are stained, remove NBT/BCIP, and rinse several times with PBST. Transfer embryos to methanol. Rinse 2X in 100% methanol, 1X in 100% ethanol, 2X in 100% methanol, and then transfer back to PBST. Remove any bits of yolk that may remain on the embryos before mounting on slides.
Embryo collection
When the sunflower plant is placed in the butterfly cage, Vanessa tend to lay embryos on and near the plant, including on the soil surface. To avoid having embryos laid in hard to reach places, we put a piece of paper around the base of the sunflower plant. Fold a piece of paper in half lengthwise, and then again. Cut a small hole in the folded corner, and cut around the edges to make a circle. Unfold, and cut a line from the outside of the circle to the small hole in the center. Place the paper around the stem and overlap the two ends of the circle and tape together once you have created a cone around the stem.
Place the sunflower plant in the butterfly cage for as long as you want to collect embryos. Carefully remove all embryos from the sunflower, allowing them to fall to a collection tray below. Age the embryos as desired.
Embryo fixation
1.Heat fix
Place the embryos (no more than 100 uL of embryos) in a 1.5 mL tube, add about 500 uL of water. Put the embryos in a boiling water bath for 3 min, then submerge the tubes in ice, and leave for 10 min. In addition to partially fixing the embryos, during the heating step, the chorions will inflate which will create space between them and the embryos, making them much easier to remove.
2.Chorion removal
Eggshells can then be removed using one of two methods: 1) Transfer embryos to PBST and to a glass dissection dish, and remove eggshells manually; 2) Transfer embryos to a Drosophila-style embryo collection basket and submerge in 1:1 bleach:water for about 1 min, then rinse several times in tap water. The first method is more labor- and time-intensive, but is sure to work, while the second method can result in embryos disintegrated if they are left in bleach too long. The amount of time to leave the embryos in the diluted bleach may need to be determined empirically given that commercial bleach products vary in their percentages of sodium hypochlorite. If using the second method, rinse the embryos several times in PBST before proceeding to the PFA fix.
Note: If you are staining germband-stage embryos, you can also go a step further and dissect the germbands out of the yolk at this point, or you can do this later (after storage in methanol).
3.Fixation with paraformaldehyde
Thaw a fresh tube of 12% paraformaldehyde (PFA). Remove PBST from each tube of embryos and add ~1 mL of 4% PFA in PBST. Rock embryos for 20 min. Remove the PFA, and rinse the embryos 3X with PBST. Gradually transfer the embryos to methanol. We do this by removing 300 uL of PBST, adding 300 uL of methanol, allowing embryos to rock a few min, then repeating this several more times until the solution is almost entirely methanol. Then rinse the embryos 3X with 100% methanol and finally store the embryos at -20C in methanol.
4.Dissection of germband-stage embryos
If you haven’t already done so, any germband-stage embryos should be dissected to remove the germband from the yolk. You can do this any day after fixation and before you want to start the in situ. Simply transfer the embryos from methanol to PBST (using the gradual transfer method described above), move to a glass dissection dish, and dissect. Remove the yolk to leave only the germbands. Gradually transfer back to methanol and store at -20C until you’re ready to begin the in situ hybridization. At many stages the germband is concave, almost bowl-shaped at the posterior end, and it can be very difficult to remove the yolk from this area without damaging the embryo. However, you do not need to remove all bits of yolk at this time; you just need to remove the yolk that is enveloping the embryo.
In situ hybridization
Gradually transfer embryos from methanol to PBST. Remove PBST and add 4% PFA; rock for 1.5 h. Rinse several times with PBST. Add about 1 mL hybridization buffer and rock until embryos sink. Incubate at 60C for 4 h. Add probe diluted in hybridization buffer and hybridize at 60C overnight.
The next day, put 2 mL of hybridization buffer and 1 mL 2X SSC in the hybridization oven to warm up. Remove probe and perform the following washes:
Buffer Temperature (C) Time
Hyb buffer (pre-warmed) 60 30 min
Hyb buffer (pre-warmed) 60 30 min
2X SSC (pre-warmed) 60 30 min
2X SSC RT 30 min
0.2X SSC RT 30 min
Rinse the embryos 3X with PBST. Wash for 2 h at RT with 10% sheep serum. Add the anti-dig-AP antibody diluted 1:1600 in 10% sheep serum; rock for 1.5 h at RT. Remove the antibody, and rinse with PBST 3X. Leave to rock in PBST overnight at 4C.
The next day, wash 3 X 20 min in PBST, then 3 X 5 min in AP staining buffer. Remove all buffer and add 500 uL of NBT/BCIP. Place tube in the dark and check progress of stain occasionally (after 3 h or so). Time for stain development needs to be determined empirically. Once embryos are stained, remove NBT/BCIP, and rinse several times with PBST. Transfer embryos to methanol. Rinse 2X in 100% methanol, 1X in 100% ethanol, 2X in 100% methanol, and then transfer back to PBST. Remove any bits of yolk that may remain on the embryos before mounting on slides.