Proximity Ligation Assay to Study In Situ Protein-Protein Interactions

NOTE: This protocol uses C. elegans strains in which potential interacting partners are both tagged. It is strongly recommended that a negative control strain be used, in which one tagged protein is not expected to interact with another tagged candidate interaction partner. Here, GFP alone was used as a negative control to assess background, as DLC-1 is not expected to interact with GFP in the worm. GFP-tagged OMA-1 was used as the experimental strain, as preliminary data suggest an interaction with DLC-1. Nematode strains co-expressing control and test proteins with 3xFLAG-tagged DLC-1 are referred to in this text as 3xFLAG::DLC-1; GFP and 3xFLAG::DLC-1; OMA-1::GFP (strains available upon request; more information in Table of Materials), respectively. Here, the 3xFLAG and GFP tags are used; however, other tags may be substituted as long as their antibodies are compatible with the PLA kit reagents.

1. Animal care

1. Keep worms on nematode growth medium (NGM) plates that are seeded with the OP50 strain of E. coli and maintain at 24 °C for optimal expression of GFP.
2. Passage adult worms every 2-3 days to propagate worms and keep them well-fed.

2. Preparation of synchronous culture

1. Synchronize worms by bleaching a plate of well-fed, gravid hermaphrodites. A bleaching protocol is described in Porta-de-la-Riva et al. Let the embryos hatch overnight in a centrifuge tube at 24 °C while rotating end over end in 10 mL of M9 minimal media (M9) buffer. This will produce a culture of arrested L1 larvae.
2. Incubate the tube of arrested L1 stage larvae on ice for 10 min, then top off the tube with ice cold 1x M9.
3. Use a centrifuge to pellet the larvae at 600 x g for 5 min at 4 °C. Carefully aspirate the supernatant so that only 1-2 mL of supernatant remains.
4. Resuspend the pellet of larvae and use a micropipette to transfer 2 µL of suspended larvae culture to a glass slide. Count how many larvae are present to determine the density of the larvae culture, which will help guide seeding of the worms in step 2.5.
   NOTE: A density of 10-15 L1 larvae/1 µL works well for seeding.
5. Use a micropipette to transfer the volume of larvae culture needed to seed approximately 100-120 L1 stage larvae on a 60 mm OP50 plate. For example, seed 10 µL of a larvae culture that has a density of 10 L1 larvae/1 µL of culture.
   NOTE: Do not exceed a volume of 40 µL to seed the larvae, or excess liquid will disrupt the OP50 lawn. If the culture volume exceeds 40 µL, repeat steps 2.3-2.4 to further reduce the volume and increase the density of larvae culture.
6. Grow worms at 24 °C. Record the time when L1s are seeded on plate and periodically check the stage of development to identify the ideal time for dissection.
   NOTE: At 52 h after seeding of L1s, worms cultured at 24 °C are typically in the young adult stage, which is the ideal stage for dissection for gonad-targeted PLA. However, the actual time at which the synchronized worms reach young adult stage may vary among strains and incubation temperature.

3. Dissection/gonad extrusion

NOTE: Dissection to extrude the gonad is necessary for gonad-targeted PLA to work successfully. This approach can also release embryos, which also work using this protocol for PLA (see Discussion for more information). After dissection, both the negative control and experimental samples are fixed and treated for PLA together in parallel. It is also suggested that an additional set of samples be prepared for the purpose of fluorescent co-immunostaining to demonstrate expression patterns of the protein partners of interest.

1. Pick 30-40 young adult worms into a watch glass dish containing 500 µL of 1x M9 + levamisole (2.5 mM final concentration). After collecting the worms, carefully remove and discard most of the media to remove bacteria that is transferred along with the worms.
2. Add in fresh 500 µL of 1x M9 + levamisole and use the pipette to gently draw up and dispense the media to rinse the worms. Carefully remove and discard most of the media to clear bacteria that is transferred along with the worms.
   1. Repeat this step 2x-3x until all bacteria are removed. After washes are completed, leave worms in about 100 µL of media to keep hydrated.
      NOTE: Do not let the worms sit in the media for longer than 7 min, as this will impair the extrusion of gonads during dissection. Perform washes under aid of dissecting microscope to monitor removal of media so that worms are not lost.
3. Using a glass or polyethylene pipette, transfer worms to a 25 mm x 75 mm microscope slide coated with 0.001% poly-L-lysine (slides used in this procedure have an epoxy coated perimeter, leaving three workspaces, 14 mm x 14 mm each). Remove excess media so that approximately 10-15 µL of media remains.
4. Under the aid of a dissecting microscope and using two 26½ gauge needles, place one needle over the other so that the ends form a pair of scissors. Using needles oriented in this fashion, cut worms behind the pharynx to release the germlines. Dissect all worms within 5 minutes.
   NOTE: More detail on how to perform dissections can be found in a previous publication by Gervaise and Arur.
5. After all worms are dissected, gently place a 22 mm x 40 mm coverslip over the slide so that it is perpendicular to the slide. The ends of the coverslip should hang off the slide.
6. Freeze the slides on a pre-chilled aluminum block maintained on dry ice for at least 20 min. Gently place a chilled pencil on top of the coverslip to prevent the coverslip from becoming loose due to ice expansion.

4. Fixation/blocking

1. When ready for fixation, flick off coverslips with a pencil or other blunt-edged tool and immediately dip the slide into a jar containing fresh, ice-cold methanol (chilled to -20 °C) for 1 min.
2. Gently wipe the edges of the slide that surround the sample so that the next reagent is held by surface tension around the sample. Apply 150 µL of fixative (2% formaldehyde in 100 mM KH₂PO_4, pH = 7.2) for 5 min at RT.
   NOTE: We have also tested a methanol/acetone fixation procedure and found that it is compatible with the PLA reaction.
3. Touch the slide to a paper towel at a perpendicular 90° angle to let the fixative run off the slide and absorb into the paper towel. Block slides 2x for 15 min at RT in a Coplin jar with 50 mL of 1x PBS/1% Triton X-100/1% bovine serum albumin (PBT/BSA).
   NOTE: Coplin jars or other types of staining jars are recommended for this blocking step and the washing steps below in sections 6-9. These provide sufficient volumes for efficient exchange of blocking or washing buffer with the sample.
4. Block slides with a PBT/BSA solution containing 10% normal goat serum. Gently wipe edges that surround the slide and apply 100 µL of the solution to the slide. Incubate for 1 h at RT in a humid chamber.
   NOTE: This step is highly recommended for staining with the αFLAG primary antibody. The humid chamber is constructed by securing glass pipettes with tape in the tray for the slides to lay on as they incubate. Dampened task wipes (Table of Materials) are placed in the tray to raise the internal humidity of the tray to prevent evaporation. The lid and tray are covered in foil to protect the samples from light during the light-sensitive steps.
5. Place the slide on a paper towel to let PBT/BSA/10%NGS solution run off the slide and gently wipe the edges of the slide. Use the blocking reagent (Table of Materials) to block slides. Apply one drop to the 14 mm x 14 mm space. Incubate slides for 1 h at 37 °C in a humid chamber.

5. Primary antibody incubation

NOTE: To obtain the best PLA results and minimal background, the dilution factor of the primary antibodies may require optimization (see Discussion for more details). Additionally, the primary antibodies should be raised in different hosts that match the specificity of the secondary antibodies used for PLA.

1. Place slide on a paper towel to let blocking reagent run off the slide and gently wipe the edges. Use the antibody diluent (Table of Materials) to dilute the primary antibodies. Apply 40 µL of primary antibody solution per 14 mm x 14 mm space.
2. Incubate slides in a humid chamber overnight at 4 °C.

6. PLA probe (secondary antibody) incubation

NOTE: For steps 6-9, use wash buffers A and B at RT. If the buffers are stored at 4 °C, then let them warm to RT prior to using.

1. Wash slides 2x for 5 min with 50 mL of 1x wash buffer A (Table of Materials) at RT in a Coplin jar. Set the Coplin jar on an orbital shaker set to 60 rpm.
2. Place slide on a paper towel to let wash buffer run off the slide and gently wipe the edges. Prepare a 40 µL solution containing PLUS and MINUS probes (diluted 1:5 with antibody diluent). Apply the solution to each 14 mm x 14 mm space.
3. Incubate slides in a humid chamber for 1 h at 37 °C.

7. Ligation

1. Wash slides 2x for 5 min with 50 mL of 1x wash buffer A at RT in a Coplin jar. Set the Coplin jar on an orbital shaker set to 60 rpm.
2. Dilute the ligation buffer (Table of Materials) 1:5 with ultrapure water. Use this buffer to dilute the ligase (Table of Materials) 1:40 to prepare a working stock of ligation solution.
   1. Place the slide on a paper towel to let the wash buffer run off the slide and gently wipe the edges. Apply 40 µL of the ligation solution to each 14 mm x 14 mm space.
3. Incubate slides in a humid chamber for 30 min at 37 °C.

8. Amplification

NOTE: Using detection reagents with red fluorophores (Table of Materials) results in the least amount of background in C. elegans tissue.

1. Wash slides 2x for 5 min with 50 mL of 1x wash buffer A at RT in a Coplin jar. Set the Coplin jar on an orbital shaker set to 60 rpm.
2. Dilute the amplification red buffer (Table of Materials) 1:5 with ultrapure water. Use this buffer to dilute the polymerase (Table of Materials) 1:80 to prepare a working stock of amplification solution and protect from light.
   1. Place the slide on a paper towel to let the wash buffer run off the slide and gently wipe the edges. Apply 40 µL of the amplification solution to each 14mm x 14 mm space.
3. Incubate slides in a humid chamber for 1 h and 40 min at 37 °C. Make sure the humid chamber is covered with foil to protect the samples from light.

9. Final washes

1. Wash slides 2x for 10 min with 50 mL of 1x wash buffer B (Table of Materials) at RT in a Coplin jar. Set the Coplin jar on an orbital shaker set to 60 rpm.
2. Wash slides 1x for 1 min with 50 mL of 0.01x wash buffer B at RT in a Coplin jar. Set the Coplin jar on an orbital shaker set to 60 rpm. This buffer is prepared by diluting wash buffer B with ultrapure water.

10. Coverslip mounting

1. Let the excess wash buffer run off the slide onto a paper towel and wipe off any residual buffer remaining on the epoxy-coated perimeter of the slide.
2. Add 10 µL of mounting medium (Table of Materials) to sample and gently lay a coverslip on top, allowing for the mounting medium to spread out.
3. Paint around the edge of the coverslip with nail polish to seal the coverslip and slide. Be gentle with application of nail polish to avoid moving the coverslip, which will damage the germlines. Let the nail polish harden for at least 20 min at RT, while slides are protected from light, before viewing it under a microscope.