Martin Lab / George Washington University
Jan 2022 update
Dissection and storage of Heliconius wings for in situs
***Important***
1) All the steps on ice to protect RNA from degradation. The time of fixation will also depend on temperature, and the times provided here are for tubes standing on ice.
2) W = wash : time indicated in minutes. Each wash : 1mL in 1.5mL Eppie for larval wing disks ; 2-3mL if using NetWells for 12-well plates (Sigma-Aldrich catalog# CLS3477-48EA). The NetWells help a lot for in situ of pupal wings, but are not necessary for fixation and storage, where 2mL screwcap cryotubes are preferred (e.g. USA Scientific catalog# 1420-9700) should fit most Heliconius pupal wings
3) Be aware that MeOH will dissolve Sharpie inscriptions on the tube : take extra care during the washes and transport (seal 1.5mL eppies with parafilm, transparent tape covering your writing on the sides of tubes).
4) Ship 1.5mL eppie tubes with the lid tighly wrapped in parafilm, otherwise it might pop up in altitude.
***What to bring on a dissection trip in the tropics***
On site you will need access to
Bring with you:
Tips :
- Only dissect 4-8 individuals per session. I spend 3-4 minutes per individual and can do 10 individuals max, but it gets a little bit crazy at the end when you have to start washing the first samples while still dissecting the last ones.
- Fast your 5th larvae for at least 2hours without food so they lose gut pressure
- Avoid coffee : makes you shiver.
- RNAse free conditions -> wear gloves, avoid sneezing and keep in mind that your skin and everything it touches is potentially dangerous for the sample quality!
- 5th instar : 1.5mL eppie tubes.
- day 3 Pupae : 2mL screw cap tubes (e.g. USA Scientific catalog# 1420-9700)
day 3 = 52-60h after pupation, but note that development is temperature development so keep the pupae OUTSIDE, next to the insectaries, to be consistent with what we have done so far.
FW well attached on cuticle in 2mL vials.
HW : fixed on the pupa, and it is ok if there is some membrane on it.
Then, after the 2nd wash in PBT, dissection of the HWs (membrane ok but better if removed)
Dissection/Fixation/Washes
***ON ICE***
- put PBS, PBT, MeOH33%-100% on ice, label your tubes
- Freshly prepare 1 or 2mL of Fix per specimen (1mL = 750 uL PBS 50mM EGTA, 250uL formaldehyde 37%)
- Clean Dissection area (EtOH, eventually, tools cleaned with RNAseZAP).
- Put tubes on Ice, prepare a clock/timer + paper to keep track of fixation time
- Anesthetize larva/pupa in ice cold water.
- Dissection in PBS. The procedure depends on the stage:
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for 5th instar Larvae
- Pin the larva in Head + Pseudopod (abdominal legs)
- Make a sub-cutaneous incision running antero-posteriorly from segments T2 to T3. Wing Disks sit under the cuticle, on a lateral line between the spiracles (and are attached to the tracheal trunk).
- If the larva has been 5th instar for more than 36 hours, the wings should be fairly obvious and developed.
-Be careful not touching the gut or it might pop.
- Enlarge the opening, cut the wing disk.
- Avoid keeping to much white substance (it will stick to the tip and tube)
- pipette the wing with a cut 20ul tip.
- Go Ahead. Fix the 4 wings in a same fix tube (Eppie 1.5mL). If the gut leaks, change the PBS. Record the time when you have done the 4 wings. Be sure the tube is on ice.
- Fixation on ice 30’-40’ (so if you wash at time 30’, your wings should be 30’-40’), mix by inverting tube a few time every 5 or 10 minutes
- Clean tools and dissection dish when you are done.
Note: there is no Tween in this step, therefore the disks tend to stick on the vial walls, so detach them by pipeting. If it becomes a problem, you can pre-lubricate your tip with PBT
- W, W, W’ in PBT (short washes of 30’’ to 5’ ; if needed, jst do two quick washes to stop fixation and then these samples can wait on ice until all your samples have been washed twice. Do a third wash and dehydrate in MeOH everything once you have stopped fixation for every tube)
- Be sure you are not loosing any wing disk while pipetting (happens often at early stages, in which case you can use a 12- or 24 well culture dish to visualize the solution you eject)
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for Day 3 (52-60h) pupae
-Pin the pupa on the side in abdomen + eyes ; cut spikes and cremaster
- Remove the strip of spikes that cover the legs with a forcep. That should come off nicely.
- With scissors or forceps, run inside the opening you just made at the level of the legs to detach the forewing from its central site of attachment (also loosens membrane)
- with scissors, gently cut all around the Forewing. The Forewing shape is nicely drawn in Heliconius so just follow this landmark, but if you go inside, you may damage the forewing. Your scissors should be flat.
- Critical Step : Very gently open the forewing cuticle on one side using forceps, like the trunk of a car. With your scissors in your right hand, gently separate the forewing form the hindwing and membrane. The objective is to obtain the forewing almost perfectly flattened on its cuticle.
- Put the forewing in the FW fix tube on ice, do the other forewing.
- Put the whole pupa, with the hindwings opened, in the HW fix tube on ice
- Record the time
- Fixation 55’-65’ (so the first FW is usually a little bit more fixed than the others)
- W, W in PBT (short washes of 30’’ to 5’ ; if needed, do two washes to stop fixation and then wait for the next samples)
- Dissect HWs in cold PBT. Pin the fixed pupa, remove the membrane, gently run around the HW with scissor to detach it from underneath, and gently remove the HW.
-Pipette the hindwing with a cut 1000uL tip and put back in the HW tube.
- W in PBT
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Progressive dehydration:
- remove ~80% of the PBT. Add cold MeOH 33% gradually. Let on ice 2’-5’, inverting the tube a few times.
- repeat the same process with MeOH 66%
- repeat the same process with MeOH 100%
- Final wash in MeOH 100%, store in freezer at -20C degrees
(note: MeOH will freeze at -80C or on dry ice… you can ship MeOH samples on wet ice if they are properly marked and parafilm but protect your writing from MeOH leaks with clear tape…)
-------------Solutions -----------
Check that your solutions are clean of growth or dust before using.
PBT = PBS + Tween20 (Detergent). Tween avoids the wings to stick to the tube, but should be avoided before fixation
dH2O : always useful . 1L of distilled H2O ; autoclave, store @RT
PBS 1X. Solution PBS10x (eg, InVitrogen) or tabs.
Dilute in dH2O, autoclave, store @RT or 4C
PBS 50mM EGTA
For 50mL: 5mL PBS10X + 5mL EGTA 0.5M + 40mL dH2O
PBT = PBS 0.01% Tween 20
for 50 mL : 50mL PBS + 200uL Tween20 25%
Fix (always prepared the same day of dissection). For each 1mL:
750 uL of PBS 50mM EGTA : 250uL of formaldehyde 37%
Note: order a fresh bottle of formaldehyde (for instance Sigma 252549-25ML) instead of using a bottle of unknown age…
MeOH mixes, made in Falcon 50mL tubes
15mL MeOH + 30mL PBS = 33% MeOH
30mL MeOH + 15mL PBS = 66% MeOH
45mL MeOH = 100% MeOH
Tween20 25% 12mL in 15mL Falcon tube
9mL sterile dH2O + 3mL Tween 20 (very viscous, so just pour it) (Sigma#P-1379)
Mix gently until homogenous, Protect from light, Store @RT
EGTA 0.5M, 100mL:
19.02g in 90mL sterile dH2O. Should completely dissolve at RT once at pH=8 (adjust pH with NaOH). Filter, Autoclave, Store @ RT
In-Situ Protocol for butterfly wing disks
Adapted from the Grace Panganiban / Carroll lab protocol and Nipam Patel Lab protocol
L5 = 5th instar larva
P = pupa (so far, I have tried only after 36h of pupation and before the ommochromme stage)
R = rinse ; W = wash
Tips :
- I perform all the steps before the oven on ice, but I invert the tubes occasionally. I perform rocking during and after the oven steps, but not during the incubation with the DIG antibody (as this creates tracheal staining artifacts)
- 5th instar wing disks : use 1.5mL vials
- Pupae : FW well attached on cuticle ; HW properly detached; -12 well plates?
Day 1
***ON ICE***
SAME-DAY DISSECTIONS
- Freshly prepare Fix. (750 uL PBS 50mM EGTA, 250uL formaldehyde 37%)
- Clean Dissection area. Cool the fix vials, PBS and PBT on ice
- Anesthesize larva/pupa 5’ max at -20C.
- Dissection in PBS. Removal of peripodial membranes for Pupae.
- Fixation 25’-40’ for L5 ; 40’-70’ for P. Note: there is no Tween in this step and the disks tend to stick on the vial walls, so detach them by pipetting. Fixation in PBT is probably OK, but I haven’t tried yet.
- R, 4x W5’-15’ in PBT
OR
STORED TISSUES
- Rehydrate progressively in cold MeOH 66%, MeOH 33%, PBT ; 2-5 minutes for each wash. Add solutions slowly to tubes or wells that still have some remnants of the previous solution to allow gradual rehydration
- R, 4x W5’-15’ in PBT]
- proteinase K 2.5ug/mL in PBT : Larval wings 2’ , Pupal wings 5’ (with cold buffer)
Add 2.5uL of 1ug/uL frozen aliquot to 1mL of ice cold PBT (see solution section)
Digest while rocking the tube by hand for 90 s, remove the proteinase K, and add the Stop Solution at 2 min. Digestion rate is sensitive to temperature.
- R, W5’ in Stop Solution. Rinse with 2x W5’ in PBT.
- Larval wings : Dissection of peripodial membrane in cold PBT (fine forceps, Dumont Biology #5). Dispatch the samples.
- Post-fix 20’, rinse with 4x W5’-15’ in PBT.
- 2x W5’ in 50:50 PBT:PreHyb
*** Not on ice. Rocking if possible ***
Day 2
- 6xW5’-30’ in PreHyb @63C, letting the last wash go overnight
Day 3
- W5’ in PreHyb @63C, transfer @RT
- W5’ in PreHyb:TBT 50:50 (now all the protocol is done @RT)
- R, 4x W5’ in TBT-BSA on nutator
- W30’ in TBT-BSA
- Incubate with Antibody anti-DIG (Roche) 1:3000 in TBT-BSA for 2h at RT
- 3xW5, 7xW15’ in TBT-BSA (important!). Use a nutator, and TBT-BSA kept on ice so the samples stay a little bit cold to preserve Alkaline Phosphatase activity.
- Let the last wash go overnight at 4C (or continue with day 4 if you can stay at least 2 more hours in the lab. However, you will probably have less control on the stainings since an overnight staining at 4C will is usually necessary)
Day 4
Solutions : check that your solution are clean of growth or dust before using. I often use filtered and autoclaved dH2O (instead of DEPC H2O) since it seems to be enough to maintain RNase-free conditions.
PBS 1X
Solution 10x (eg, InVitrogen) or tabs.
Dilute in dH2O, filter, autoclave, store @RT or 4C
PBT = PBS 0.1% Tween 20
for 50 mL : 50mL PBS + 200uL Tween20 25%
Fix (freshly prepared). For each 1mL:
750 uL of PBS 50mM EGTA + 250uL of formaldehyde 37%
Proteinase K : stock = Promega 20mg/ml Proteinase K, catalog number MC5005
- dilute your stock 20x and dispatch in well-marked 1ug/uL aliquots stored at -20C.
- for the digestion, add 2.5uL to 1mL of ice cold PBT (400x dilution)
Digest while rocking the tube by hand for 90 s, remove the proteinase K, and add the Stop Solution at 2 min.
Post Fix (freshly prepared). For each 1mL:
150 uL formaldehyde 37% + 850 uL PBT.
Stop Solution = 2mg/mL glycine in PBT
For 50mL :
10mL glycine 10g/L (this stock solution can be filtered, autoclaved and kept @4C)
5mL PBS 10x
200uL Tween20 25%
fill up with sterile dH2O until 50mL
PreHyb 50 mL:
10mL sterile dH2O
25mL formamide
12.5mL 20X SSC(check that pH=4.5 prior to addition)
200uL Tween20 25%
500uL 10mg/mL Salmon Sperm DNA (heat denature 5’ @80C prior to addition)
lower pH to pH5-6, fill up to 50mL with sterile dH2O. Store @-20C.
Hyb 50 mL:
5mL sterile dH2O +5mL glycine 10g/L
25mL deionized formamide
12.5mL 20X SSC
200uL Tween20 25%
500uL 10mg/mL Salmon Sperm DNA (heat denature 5’ @80C prior to addition)
lower pH to pH5-6, fill up to 50mL with sterile dH2O. Store @-20C.
20x SSC, 1L:
900mL dH20 + 175.3g NaCl + 88.2g Sodium Citrate, dihydrate
Once at room temperature, lower pH to 4.5. Fill-up to 1L. Filter, Autoclave, Store at 4C or RT
TBS 10X 500mL:
125mL Tris 1M pH 7.5 + 40g NaCl + 1g KCl
add dH2O to mix and bring up to 500mL. Filter, Autoclave and store at 4C
TBS 1L:
100mL TBS10X + 900mL H2O
I usually autoclave some bottles of this dilutions the day I prepare 10X stock as well
TBT = TBS (1X) 0.1% Tween20. 50 mL :
50mL TBS + 200uL Tween20 25%
TBT-BSA (freshly prepared) 50mL:
50mL TBT ; Check that pH=7.5, and then add 0.05g BSA, vortex thoroughly
Alkaline Phosphatase Buffer, 50mL (freshly prepared)
250 uL of MgCl2 1M
5mL of NaCl 1M
5mL of Tris 1M (pH9.5)
200 uL Tween20 25%.
Staining Solution : dispatch BM Purple (stored at 4C) in Eppie Tubes, pellet the precipitate at max. speed 2min on a table top centrifuge, use the supernatants for Alkaline Phosphatase staining.
1M Tris pH=7.5, 1L:
121.1 g of Tris Base dissolved in 900mL H2O.
Once at room temperature lower pH to 7.5 (about 50mL of 12.1M HCl may be required)
Fill-up to 1L, filter, autoclave and store @RT
1M Tris pH=9.5, 1L:
121.1 g of Tris Base dissolved in 950mL H2O.
Once at room temperature adjust pH to 9.5
Fill-up to 1L, filter, autoclave and store @RT
Tween20 25% 12mL in 15mL Facon tube:
9mL sterile dH2O + 3mL Tween 20 (very viscous, so just pour it) (Sigma#P-1379)
Vortex, Protect from light, Store @RT
EDTA 0.5M, 100mL:
18.612g in 90mL sterile dH2O. Should completely dissolve at RT but pH=8 (adjust pH with NaOH). Filter, Autoclave, Store @4C or RT
EGTA 0.5M, 100mL:
19.02g in 90mL sterile dH2O. Should completely dissolve at RT but pH=8 (adjust pH with NaOH). Filter, Autoclave, Store @4C or RT