From Wikionchus

Contents 1 Intro 2 Finney-Ruvkun protocol for larvae and adults 2.1 Fixation, permeabilization and immunofluorescence staining 2.2 Fixation 2.3 Reduction 2.4 Oxidation 2.5 Staining: 2.6 Buffers and Solutions 3 Modified Drosophila protocol for Pristionchus embryos (Schlager-Meyer stain) 4 Links

Intro

There are two protocols for Antibody staining in Pristionchus. Finney-Ruvkun for larvae/adults - first applied and optimized for Pristionchus by Kolotuev and Podbilewicz. A modified Drosophila protocol for embryos. We have recently developed a separate protocol for staining embryos. We did that because the eggshell and vitelline membrane of Pristionchus embryos are not efficiently permeabilized with the Finney-Ruvkun protocol. This protocol is a modified standard "Drosophila" protocol. The two protocols are mutually exclusive: While "Finney-Ruvkun" is good for larvae and adults, it is not satisfactory for embryos. The embryo protocol ("Schlager-Meyer" :) is good for embryos up to the 3 fold stage but performs badly for anything older.

Finney-Ruvkun protocol for larvae and adults

Fixation, permeabilization and immunofluorescence staining

(Protocol by Mike Finney, 1991, copy-paste of a protocol given to the lab by Benjamin Podbilewicz)

This method is used in C.elegans for immunoflourescent staining of adherens junctions at the cell’s apical domain. It is a very convenient way for staining large quantities of the nematode. The main problem with the staining in "P. pacificus" is its harsh cuticle, compared to that of C.elegans. I tried to optimize this protocol by performing the experiment under different conditions: 1) changing the number of the freezing - thawing cycles during permeabilization; 2) performing this procedure for the time range from 30min to 2h. The reason behind changing those specific parameters was to get a better cuticle perforation in order to permit the staining materials to get into the nematode. During the freezing – thawing cycles the frozen solution particles tear mechanically the cuticle, and literally break this guard cover. The time of thawing is important because of the fixative action of the para – formaldehyde. This polymeric chemical builds the net, which helps to keep the morphology of the organs similar to real, before the chemical treatment. Time in this case may be an important feature.

The best staining was obtained with a thawing time of 1h, one round of procedure in 2% of para – formaldehyde. Procedure description:

All the reactions were performed in 1.5ml eppendorf tubes at room temperature, unless specified. In all cases the spins in microfuge were performed at 5000rpm for 2min.

Fixation

• 1. Wash the worms from the unstarved plates with M9. Leave it on ice to digest the

bacteria and to sink .

• 2. Add cold 1xMRWB and 20% para-formaldehyde* to a final concentration of 2%

(total volume , 1ml).

• 3. Mix well with the nematode pellet. Freeze in liquid nitrogen (dry ice/ethanol is also

a possibility).

• 4. Permeabilization-melt on ice and incubate with occasional agitation for 1-2hr.

(Frozen samples may be processed immediately or stored for a long time.)

• the para-formaldehyde has to be freshly prepared.

Reduction

• 1. Wash the worms twice with Tris Triton buffer.
• 2. Incubate the worms in Tris Triton buffer with 1% β-mercaptoethanol for 2hr. at

37°C with occasional agitation.

• 3. Wash the worms once in 10-15volume 1xBO3 buffer (for the eppendorf tube 1.5 ml

is enough).

• 4. Incubate in 1xBO3 buffer with 10mM dithiothretol (DTT) for 15 min at R.T. with

agitation.

Oxidation

• 1. Wash the worms once in 10-15 volume 1xBO3 buffer.
• 2. Incubate the worms in 1xBO3 buffer with 0.3% H2O2 for 15 min at R.T. with gentle

agitation, when the tubes are closed tightly with parafilm .

• The worm pellet is very loose after this step! Be careful not to lose too many worms while removing the supernatant.
• 3. Wash the worms once in 10-15 volume 1xBO3 buffer.
• 4. Wash 3 times for 15 min with antibody buffers B.

(Samples may be stored at 4°C in antibody buffer A)

Staining:

• 1. Incubate fixed worms with a primary antibody (MH27 diluted in antibody buffer

A 1: 300) at 4°C overnight on the rocker platform.

• Anti ds-DNA antibodies are good positive controls.
• 2. Wash the worms with an antibody buffer B 3 times for 15 min.
• 3. Incubate with 200 μl of the secondary antibody (FITC 1:200 in PBS milk) for 2hr.

at R.T. (In the case that PI is used, add RNase with a final concentration of 200μg/ml).

• 4. Wash the worms once with antibody buffer B.
• 5. Add to the worms 200μl of PI and/or DAPI at a final concentration of 1:1000 both.
• 6. Incubate for 15min R.T.
• 7. Wash the worms with antibody buffer B 3 times for 15 min.
• 8. Wash once with antibody A.
• Place the worms on a glass slide, coated with polylysine, allow excess solution to air dry, and mount using moviol.

Store the stained worms in aluminum foil covered eppendorf tubes or the slides in tightly closed boxes at 4°C.

• Worms may also be mounted on 2% Agar pads, the same that are used for Nomarski microscopy.

Buffers and Solutions

• 20% para-formaldehyde solution:

Weigh approximately 250mg of dry para-formaldehyde in eppendorf (work under the air hood with gloves). Multiply the weight (in mg) by 4.5 and add that volume in ml of 5mM NaOH. Place in 65°C bath for 30 min, with occasional agitation. Spin for 1min.Use the solution immediately!

• 2xModified Ruvkuns witches brew (MRWB):

KCl 160mM NaCl 40mM Na2EGTA 20mM Spermidine HCl 10mM Na PIPES pH 7.4 30mM Methanol 50% (add before working)

• Tris Triton buffer:

Tris-Cl pH 7.4 100mM EDTA 1mM Triton X-100 1%

• 40xBO3 buffer ( pH 9.2 at 25 mM ):

H3BO3 1M NaOH 0.5M

• Antibody buffer A:

1xPBS 1%BSA 0.5%Triton X-100 0.05% Na Azide 1mM EDTA

• Antibody buffer B:

The same as antibody buffer A except for 0.1% BSA.

Modified Drosophila protocol for Pristionchus embryos (Schlager-Meyer stain)

• 1. Grow 30 6-cm plates of worms to mixed stage
• 2. You do not need to bleach to collect eggs. Larvae and adults will not hinder the staining - they will just not be stained.
• 3. Wash your worms off the plates with water. Wash at least 3 times with large volumes to remove as many bacteria as possible. Be paranoid about this. The fewer bacteria - the better.
• 4. Resuspend worms in 25 ml water and split into 5 14-ml Falcon tubes, containing 5 ml each.
• 5. Add water and "Clorix" (household bleach) to 5 different dilutions in the same total volume (10 ml, ratios refer to the dilution of bleach in the total volume): 1:2, 1:4; 1: 8, 1:16, 1:32. Work fast and concentrate ! Note that this step is used to pre-bleach (not destroy) the eggshell. Therefore the short incubation time! This step is done to permeabilize the eggshell for heptane - which is later used to dissolve the vitelline embrane. This sometimes leads to weird, strong signals in partially dissolved cuticles of larvae and adults. These are false-positive because they are also seen in 2° only negative controls.
• 6. Incubate exactly 5 minutes on gel rocker/shaker (slow mixing).
• 7. Immediately centrifuge in Heraeus centrifuge: 2 min, 2.5 krpm, brake setting: 2.
• 8. Wash twice with room temperature PBS - work fast.
• 9. Take off supernatant to 2 ml total volume (worms in PBS).
• 10. Add 2.6 ml n-heptane. Heptane is used to dissolve/permeabilize the vitelline membrane.
• 11. Add 666 µl 4% PFA (freshly depolymerized in PBS), final conc. 1%.
• 12. Place on horizontal shaker, shake at 250 rpm for exactly 5 minutes (Fast shaking is required to maximize the interphase of the PBS and heptane phases).
• 13. Spin as above.
• 14. Take off supernatant close to the pellet.
• 15. Wash with 5 ml MeOH abs. twice.
• 16. Store/Freeze in 2ml MeOH or proceed with staining.
• 17. Wash with PBS 3 times.
• 18. Use 200 µl aliquots for one staining. Centrifugation steps are implicit. 2 min, 2krpm in Eppendorf centrifuge. "Soft" setting ON.
• 19. Block on rotator in 500 µl PBSTB for at least 1 hour.
• 20. Incubate in 1° Ab in 300 µl total volume. For embryos anti-alfa tubulin or anti-ds-DNA are very good positive controls. If anti-ds-DNA is used then no DAPI is needed later on. Incubate over night at 4°C. Use a rotator for all incubation and washing steps. Always include at least one negative control for each 2°Ab used - a sample with no 1°Ab. Run one sample for each initial dilution of Clorix. Usually 1:2 is too strong - embryos burst. Usually 1:32 is too weak - embryos are not permeabilized.
• 21.Wash 3-5 times in 500 µl TBSB for 15 minutes each.
• 22. Incubate in 2° Ab for 2 h at room temperature. I use GaM-Cy3 to detect positive controls (like for example mouse anti ds-DNA) and GaR-FITC to detect my self made rabbit antibodies.
• 23. Wash at least 4 times in TBSB for 15 minutes each.
• 24. Resuspend in Vectashield (around 50 µl) and mount for microscopy on Agar pads.

Links

Back to Pristionchus pacificus Protocols page.