Indirect Immunofluorescence

Paraformaldehyde method (microtubules)

Reagents

           

PHEM buffer pH 6.9  (for 250 ml)

            60 mM PIPES  (4.54 g)

            25 mM HEPES  (1.5 g)

            10 mM EGTA  (950 mg)

            2 mM MgCl₂  (100 mg)

            (dH₂O to 250 ml)

 

Practical notes: 

            The PIPES will not dissolve until the pH is raised to at least 6.  Add PIPES to 225 ml of  dH₂O.  Raise the pH to 6.9 (with 5M NaOH), add the remaining reagents, and add dH₂O to bring to 250 ml.  Raise to a final pH of 6.9 (with 5M NaOH).

 

Phosphate-buffered saline (PBS)  pH 6.9

            9 g NaCl

            1.66 g Na₂HPO₄·7H₂O

            820 mg NaH2PO₄·H₂O

            bring to 1 L with dH2O

            Adjust the final pH to 6.9 (with 5M NaOH).

 

Fixative:  3% Paraformaldehyde, 0.2-0.5% Triton X-100, in PHEM buffer.

            40 mls PHEM buffer pH 6.9 (see above)

            120 µl Triton X-100

            10 mls 16% Paraformaldehyde solution (Electron Microscopy Sciences #15710)

 

Practical notes: 

            Use a wide-tip 200 µl micropipet tip or a 1000 µl micropipet tip for measuring the Triton X-100 (it is viscous).  Add Triton X-100 to PHEM buffer and shake or stir well for several minutes before adding paraformaldehyde. Aliquot and freeze (15ml disposable centrifuge tubes work well, 4 to 6 mls in each).  Thaw before needed.

 

Reagents to be made the day of use

 

Antiquenching mounting medium (Sanders, 1995).

DABCO 

            250 mg DABCO (Sigma D-2522) in 1 ml dH2O, add to 9 ml glycerol.  This can be easily mixed (by inversion) in a 15 ml centrifuge tube. 

 

1% BSA-PBS (for each sample; increase as needed)

            30 mg BSA (Bovine Albumin:  Sigma A-7906) in 3 mls PBS.  Shake to dissolve. 

 

0.1% BSA-PBS (for each sample; increase as needed)

            30 mg BSA in 30 mls PBS.  Shake or stir to dissolve.

Polylysine-coated coverslips

1)  Fill a coverslip staining jar (small Coplin jar) with 9 mls poly-L-lysine solution           (0.1% w/v in H₂O, Sigma P8920).  Alternatively, put 20 ml poly-L-lysine in a      50 ml centrifuge tube.  The polylysine can be stored in the coverslip staining     jar or centrifuge tube for at least several weeks.

2)  Soak the coverslip in polylysine for 5' (up to 7 coverslips in the staining jar:  4             straight across, and 3 on the diagonals;  one at a time is best in the centrifuge    tube).

3)  Set on edge, on absorbent paper to dry.  Use within a few hours.

 

Preparation of Samples

Protocol outline

1)  Pellet cells in 15 ml centrifuge tube.

2)  Wash cells with PHEM buffer.

3)  Fix cells.

4)  Pellet cells and remove fixative.

5 & 6)  Two rinses with 0.1% BSA-PBS.

7)  Add 1 ml 0.1% BSA-PBS and transfer to microfuge tube.

8)  Pellet cells.

9)  Add primary antiserum.

10)  Two rinses with 0.1% BSA-PBS.

11)  Add secondary antiserum.

12)  Pellet and remove antiserum.

13)  Counterstain.

14)  Two rinses with 0.1% BSA-PBS.

15)  Resuspend in 300-1200 µl 0.1% BSA-PBS and allow to settle on coverslips.

16)  Meanwhile, prepare slides.

17)  Wick away buffer and place onto prepared slides.

18)  Blot edges and seal with Permound

 

Protocol in detail

1)  Pipet 2-5 mls of cells (at 200,000 cells per ml) into a 15 ml centrifuge tube. Centrifuge (IEC clinical:  #4 for 2'), decant supernatant, and resuspend the pellet (by finger flicking) in the liquid that remains.

2)  Add 5 ml PHEM buffer to wash cells.  Centrifuge, decant supernatant, and resuspend in the liquid that remains.

3)  Working in a fume hood, add 1-2 mls of paraformaldehyde fixative to cells, and let sit at room temperature for 30'.  Increase the amount of fixative if the sample is large—the ratio of fixative to pelleted sample should be about 4 to 1 or greater.  Cells may be fixed for as long as 60', beyond that, however, the detergent begins to cause too much cell degradation.

4)  Centrifuge, decant supernatant, and loosen pellet by finger flicking.  Dispose of supernatant (containing paraformaldehyde) appropriately.

5)  Add 5 mls 0.1% BSA-PBS for 5' to rinse cells,.  Centrifuge, decant supernatant, and resuspend. 

6)  Repeat step 5, but do not resuspend the pellet by flicking. 

7)  Add 1 ml 0.1% BSA-PBS, mix with the pipetor, and transfer solution to a microfuge (Eppendorf) tube. 

8)  Centrifuge (IEC clinical #4 for 2' or equivalent) and aspirate supernatant.  Resuspend the pellet in the liquid that remains by flicking.

9)  For primary staining, add 200 µl (or up to 1 ml) diluted primary antiserum (in 1% BSA-PBS) to the pellet.  Time will vary depending on the antiserum, and incubation at 31°C or 37°C may be beneficial. (30 to 60 min at 31°C is typical for the antisera we use.) Two different primary antisera can be combined in this step.

10)  After incubation, centrifuge, remove the supernatant by aspiration and resuspend the pellet.  Rinse the sample (centrifuge, aspirate, resuspend) twice with 1 ml 0.1% BSA-PBS each time. 

11)  For secondary staining, add 200 µl  to 1 ml diluted secondary antiserum (in 1% BSA-PBS) to the pellet.  As with primary staining, time will vary depending on the antiserum, and incubation at a higher temperature may be beneficial. 

12)  After incubation, centrifuge, remove the supernatant by aspiration and resuspend the pellet. 

13)  If counterstaining with Sytox nuclear stain, add 1 ml 01% BSA-PBS to pellet along with 5 µl of 1:1000 Sytox for 5'. 

14)  After 5', rinse the sample (centrifuge, aspirate, resuspend) twice with 1 ml 0.1% BSA-PBS each time.  If not counterstaining, simply rinse twice. 

15)  Add 300 µl per intended coverslip (typically 1200 µl per sample for 4 coverslips).  Place 300 µl on each poly-L-lysine coated coverslip and allow to settle (in a moist chamber if needed) for 20 minutes.

16)  During the 20 minutes, prepare slides.  To prevent crushing of the cells, put four tiny drops of Permount on the slide, approximating where the corners of the coverslip will fall.  Put one or two drops of antiquenching mounting medium on the slide. 

17)  After 20 minutes, wick away the liquid from the coverslip with pieces of torn filter paper.  Blotting from two sides simultaneously will allow a maximum number of cells to adhere to the coverslip.  Quickly (before it begins to dry) place the coverslip onto the prepared slide. 

18)  Blot excess moisture from the edges of the coverslip with pieces of torn filter paper, and seal the edges of the coverslip with Permount.

 

Practical notes:

The number of washes given above is minimal, as is the time for staining.  The number of washes could be increased to reduce non-specific staining (as many as six between and after staining).  The length of time for staining could be increased (from 4 hours to overnight) to aid in binding, or so that more dilute antisera can be used. 

A moist chamber can be made by placing a circle of filter paper in the lid of a Petri dish and moistening it with H2O.  Parafilm-M or filter paper in the bottom of the dish will prevent the coverslips from sticking to the bottom. 

Antisera generated against different epitopes can require totally different (even antithetical) fixation procedures (e.g., paraformaldehyde versus ethanol).  It is worth noting that the fixative one uses when screening for effective antisera can define and limit conditions for later analysis.

                                                                                                                            

 

 

Triton-X/  ETOH Method.

(For special, cortical epitopes)

 

Reagents:

 

TRIS-Buffered Saline (TBS)

            Make stocks A (200 mM TRIS) and B (200 mM HCL).  For 1 Liter TBS:

            100 mL Stock A

            44 mL Stock B

            9 g  NaCl

            dH₂O to  1 Liter.

 

Adjust pH to 7.4.  Store in fridge.

 

 

Fixative:

            35-50% Ethanol

            add 120 microliters TX-100 to 100 mL of 35% ETOH.

            Store 2-6 degrees C. 

            Ice while using!

 

Practical Notes:  use wide tip 200 microliter micropipette or a 1000 microliter tip to measure TX-100 (viscous).

 

DAPI counter stain:  1 microgram / mL.

 

Antiquenching compound.

 

Reagents to be made day of use:

 

1% BSA in TBS.

            Add 20 mg BSA to 1 mL TBS.  Shake to dissolve.

 

0.1 % BSA/TBS.

            Add 200 mg of BSA to 200 mL TBS.  Shake or stir to dissolve. 

 

Protocol:

 

1) Pipette 2-5 mL of cells (200K) into a 15 mL centrifuge tube.  Chill on ice 5 min.

            Spin  IEC #4 2 min.  Decant, resuspend pellet by flicking.

 

2) Add 5 mL TX-ETOH (iced) keep on ice 10-30 min. 

 

3) RT  Add 5 mL 0.1 % BSA-TBS/ spin/ pellet/ flick. 

 

4) Repeat (no flick).

 

5) Add 500 microliters of cells in BSA  to coverslip, settle for 20 min, in  moist chamber.

 

6) Blot BSA/TBS, add 200 microliters primary antiserum in 1% BSA to form drop, float           coverslip face down.  45 min at 37 C.

 

7) Rinse 2 X in 0.1 % BSA.

 

8) Float coverslip on secondary antiserum (200 microliters).

 

9) Wash 2 X.  (5 min each).  (DAPI in first).

 

10) Drops of nail-polish to form feet on slides.  Mount in Vectashield.

 

Double Fixation:

 

1) Wash cells in Phem buffer.

 

2)  Fix 5 min in 1-2 mL 3% paraformaldehyde fixative.

 

3) Add 10 mLs BSA-PBS.  Spin/ pellet/ flick. 

 

4) Add 5 mL cold 15% ETOX/ TX100.    Ice for 10 min.  Then add 10 mLs BSA/PBS.

 

5) Spin/ decant/ rinse PBS…. Stain.