Other Protocols

Western Blots

Excised brain tissue will be resuspended in 100 ul of buffer H (50 mM beta-glycerophosphate, 1.5 mM EGTA, 0.1 mM Na3VO4, 1 mM DTT), then sonicate for 15 sec. 100 ul of 5X sample buffer will then be added and the samples heated to 90 degrees C for 10 min. After vortexing and centrifugation, 30 ul of extract will be loaded onto a 12% SDS-PAGE gel and electrophoresed using standard procedures. After protein has been transblotted, membranes (Immobilon P: Millipore) will be blocked with 10% powdered milk in PBS, then incubated (4 degrees C overnight) in PBST with primary rabbit antibody against MKP-1 or MKP02 (1:2000). Membranes with then be treated with a goat anti-rabbit IgG alkaline phosphatase-conjugated secondary antibody (1:2000), and immunoreactivity will be developed using the Western-star alkaline phosphatase detection system (Tropix). Membranes will be washed 6X with a 5% milk/PBST solution after each antibody treatment. MKP expression will be normalized to total protein by the bicinchoninic acid method (Pierce chemical).

Fixing Cells for Apoptosis Assay

1. At the end of the incubation with inducer, remove plate from incubator.

2. Add to each well (do not wash wells first) 1.0 ml 100% Ethanol (this will approx 70% EtOH if the original volume was 700 ul). Mix in by gently swirling plate or with a pipetter.

3. Store plate in 4C for at least 30 minutes to fix cells.

4. When ready to do PI assay, centrifuge the plate to sediment unattached cells (1500 rpm 7 min), then carefully aspirate the supernatant so as not to remove detached cells.

5. Add to each well 200 ul of PI+RNAse solution (see below), incubate at 37 C for at least 20 minutes to digest RNA.

6. Score apoptotic nuclei/20X field as a percentage of total nuclei/same field, on the JENA scope.

PI+RNAse solution:

1 ml PBS pH 7.3

25 ul PI (0.2 mg/ml)

5 ul DNAse-free RNAse solution

Staining Brains with Neutral Red

1. Mount brain sections (40 micron), on PDL coated slide.

2. Stain with 1% Neutral Red (in water) for 2 min.

3. Was with water several times.

4. Dehydrate with 50%, 75, 85%, 90%, 95%, 100% EtOH.

5. Clarify with 50% EthOH/50% Xylene, 100% Xylene for 1 min.

6. Seal with permount.

Coating slides with PDL:

Put 5-10 ul of PDL (5 mg/ml) on slide and spread the PDL out. Air dry.

Membrane Stripping Method #1 - Heat/Detergent

Required Equipment and Solutions:

Stripping: 100 mM 2-mercaptoethanol, 2% (w/v) SDS, 62.5 mM Tris-HCl, pH 6.7

Buffer (same as used in initial immunodetection)

Shallow tray, large enough to hold membrane.

Stripping:

1. Place the blot in stripping solution and agitate for 30 minutes at 50 degrees C.

2. Place the blot in buffer and agitate for 10 minutes. Repeat with fresh buffer.

3. (OPTIONAL) Repeat the initial detection protocol (omitting the primary antibody) to make sure that the antibodies have been inactivated or stripped from the membrane.

4. Place the blot in buffer and agitated for 10 minutes.

5. Proceed to the blocking step for the next round of detection.

Membrane Stripping: Method #2 - Low pH

Required Equipment and Solutions:

Stripping solution: 25 mM glycine-HCl, pH 2, 1% (w/v) SDS

Buffer (same as used in initial immunodetection)

Shallow tray, large enough to hold membrane

Stripping:

1. Place the blot in stripping solution and agitate for 30 minute.

2. Place the blot in buffer and agitate for 10 minutes. Repeat with fresh buffer.

3. Proceed to the blocking step for the next round of detection.

Staining for Barrel Formation

1. Prefuse mouse with 1% PFA/BS (100 ml at 15-20 ml/permin). [pump from GILSON.] needle = 25G 5/8.

2. Dissect the brain out of the skull.

3. Cut 1/3 from the front and cerebellum. Cut straight along the fissures, then use knife to peel the cortex. Further separate the cortex (from layer 1 to layer 6) from hippocampus and striatum by fine forceps and surgeon knife. Separation of striatum may be difficult and may need to be cut off by knife.

4. Sandwich cortex between filter paper, and then between glass slides. Put extra filter paper on either side of glass slide as well. Then hold the sandwich by rubber band.

5. Post fix in 4% PFA/PBS for 12 hours.

6. Cryoprotection by 30% sucrose/PBS

7. Wrap a glass slide with parafilm. Put the dorsal side of the cortex on the parafilm and put the glass slide on smashed dry ice. Then, bury the sample (brain slice) with dry ice powder to freeze the sample.

8. Keep it frozen and put into a 450 ml cornical tube (pre-cooled on dry ice).

9. Mount and cut by crystat.

10. Cut 30-40 um sections, and number each section to follow up.

37 degrees C in 40 mg/ml sucrose, 0.6 mg/ml DAB or PBS, 0.4 mg/ml cytochrome c from horse heart (Sigma) in PB (0.1M phosphate buffer, pH 7.4) for 2 hours.

Rinse with PBS and mount. (Do it in the dark, cover with foil.)

Golgi Impregnation Protocol

Day 1 Preparation:

1. Reserve Perfusion room, obtain an empty waste collector, and check if pump and its tubing are ready.

2. Prepare solutions: 4% Paraformaldehyde, saline solution (0.9% NaCl) 1 Liter.

4% Paraformaldehyde (for 1500 ml):

3.3 g sodium phosphate monobasic (1H2O)

33.75 sodium phosphate dibasic (7H2O)

Add ddH2O to 750 ml

pH 7.4

In a ventilation hood, place the above buffer on a hot plate and heat up to 60 degrees C. Shut down heater, add 60 g PFA, stir to dissolve, let stand for a while, add ddH2O to 1.5 L, check pH 7.4 with pH paper, filter, cover flask with parafilm, and keep in cold room until day 2.

Day 2 Perfusion:

1. Take out about 100 ml and keep it on ice, take the rest of the 4% PFA from the cold room and let it warm up to room temperature for 1 hour.

2. Set up the pump, speed 38. Tighten the tubing to about 2-3 clicks away from the maximum (this will give you a flow rate about 12-15 ml/minute. Squeeze out air bubbles.

3. Needles: 23 3/4

4. Metafane: open chest, insert needle about 4-5 mm into the left ventricle, cut right atrium, start pump with saline, perfuse with saline for 45 sec, stop pump, switch to 4% PFA. After 1 minute, you should see muscle cramp and liver turning pale. Do not clamp the needle with a hemostat; it will cause capillaries to be swollen. Perfuse for 8-10 min, stop pump, and take out the brain (be very careful; damage to the structure will result in bad cutting and failure in impregnation).

5. Put brains in 4% PFA, in cold room, until day 3.

6. Prepare 300 ml 3% potassium dichromate, overnight, room temperature, wrap with aluminum foil.

Day 3 Cutting Brains

1. Wash blades with ethanol.

2. Adjust blade vibration to 6, advance speed to 6.

3. Cut brains. Use a metal block-cutter to get smooth and even cut.

4. Glue brains onto the stage with crazy glue.

5. Cut 100 um/slice in 3% potassium dichromate in a ventilation hood. Put slices in a 24 well plate, 4-5 slices/well, with 2-3 ml of potassium dichromate.

6. Wrap with aluminum foil and leave overnight (do not leave it longer, or the tissue will be too stiff).

7. Wash and prepare Coplin jars for tomorrow.

8. Prepare (50m. Coplin jar) 1.5% AgNO3.

Day 4 Assembly and Impregnation

1. Take the slices out of the potassium dichromate, dip into distilled water, and quickly pick them up again. Put them on a slide.

2. With a Kimwipe, dry out the excessive water, but do not try it too much because the coverslips will be hard to seal with tissue, which will result in crystals all over the tissue, rendering it useless.

3. Assemble the "sandwich." Crazy glue the 4 corners of the coverslip, drop it down slowly, use the end of the paint brush to gently squeeze out air so that the coverslip and the tissue have a perfect contact.

4. Add 1.5% AgNO3 (every Coplin jar needs 50 ml).

5. Leave 2-3 days (1 day too short, 4 days too long) in a dark cabinet.

Day 7 Mounting Slides

1. Wipe the slides very clean, examine quickly under microscope, and pick impregnated tissues.

2. Disassemble "sandwich" (move coverslips sideways; don't lift it up -- easy to tear tissues).

3. Put tissues in ddH2O.

4. Put in 95% EtOH for 2 min, then 100% EtOH for 1.5 min.

5. Put in xylene until the tissues are clear (usually takes 15 sec).

6. Transfer tissues to slides. Do not dry tissues out because they will shrink and warp up. Simple get rid of the excessive xylene. Add ample amount of Depex Mounting Medium (electron Microscope Sciences, Cat #13514). Coverslip.

7. Let them dry in a ventilation hood for at least 14 days. Block light by covering them with aluminum foil.

Protocol for DIG In Situ Hybridization

Sections

- Rapidly freeze brain tissue in -50 degrees C to -60 degrees C isopentane.

- Make 14 um cryo-sections, store at -80 degrees C.

Making RNA Probe

In vitro transcription (reagents from Roche):

2 ul 10 x RNA labeling mix

2 ul 10 x transcription buffer

1 ul RNase inhibitor

2 ul SP6 or T7 or T7 RNA polymerase

Linearized DNA template and H2O up to 20 ul

37 degrees C for 2 hours

DNase 2 ul, 37 degrees C for 15 min

2 ul EDTA (0.2 M), 1 ul 2% agarose minigel check

LiCl precipitation: 2.5 ul 4 M LiCl (from Ambion) + 75 ul ice-cold ethanol; -80 degrees C, overnight; dissolve in 50 ul depc-H2O + 1 ul RNase inhibitor; make aliquots and store at -80 degrees C

Hybridization

Hybridization buffer: 50% formamide, 5 x SSSC, 5 x Denhardt's, 250 ug/ml yeast RNA, 10 mM DTT, 100 ug/ml sperm DNA

Set up a 100 degrees C heat block, and 58-68 degrees C oven. Warm up hybridization buffer.

1. Take slides out from -80 degrees C. Air dry (can use hair dryer). Demarcate the tissue area using an ImmEdge pen from Vector Laboratories.

2. 4% ice-cold PFA for 10-20 min

3. Wash with PBS twice for 5 min each

4. 0.3% H2O2 in H2O, 30 min (if you use HRP later)

5. Wash with PBS three times for 5 min each

6. 0.1 ug/ml Proteinase K (Roche) in 10 mM Tris-HCl, pH 7.5, 10 min (25 degrees C)

7. Rinse in 10 mM Tris-HCl

8. 0.1 M Triethanolamine, 2 min (stirring). (Prepare in advance: 600 ml of water plus 7 ml of triethanolamine, stirring). Add 1.5 ml acetic anhydride (dropwise) (keep stirring), 8 min (Note: mix well but don't do damage to the tissues).

9. Wash with PBS three times for 5 min each.

10. Incubate in hybridization buffer at 58-68 degrees C, 1-2 hours (remember to add denatured salmon sperm DNA to 100 ug/ml) (on paper towels soaked in 50% formamide + 5 x SSC, warmed up to 58-68 degrees C).

11. Hybridization (hyb buffer + 0.5 to 2 ul probe/ml buffer) at 58-68 degrees C, overnight (at least 12 hours) (remember to add denatured salmon sperm DNA).

12. Wash 5 x SSC, room temperature, 5 min

13. 2 x SSC rinse, then 10 ug/ml RNase A 30 min, room temperature.

14. Wash twice in 0.1 x SSC, 72 degrees C, 30 min

15. Wash in 0.1 SSC, room temperature, 5 min

16. TNT (from PerkinElmer TSA kit), 5 min

17. TNB, 30 min

18. anti-DIG-HRP, 1:2000, 1-2 hours, 37 degrees C

19. Wash twice in TNT, 10 min

20. Add TSA-biotin, 10 min

21. Wash three times with TNT, 10 min

22. Streptavidin-Cy3, 1:500, room temperature, 1 hour or ABC followed by DAB (Vector Laboratories)

Poly-D-Lysine/Laminin plates for Primary Neuron Cultures

Solutions

Poly-D-lysine from Sigma (P1024, hydrobromide, mol wt > 300,000)

Laminin from GIBROL: Natural Mouse Laminin

Coating Plates:

1. Thaw 1 aliquot each of poly-D-lysine, laminin solution

2. Add the two solutions to 18.9 ml sterile H2O to make total 20 ml. (Final concentration: 66.7 ug/ml poly-D-lysin, 6.7 ug/ml laminin)

3. Use 2 ml mixture per 60 mm plate, 0.5-1 ml mixture per 35 mm plate, 2 ml per 35 mm plate with glass coverslips. Place plates at 37 degrees C tissue culture incubator 2 hours - overnight.

4. Rinse 3 x with sterile H2O and use to plate cells or store at 4 degrees C. (This corresponds to 3.3 ug poly-D-lysine/cm2, 0.33 ug laminin/cm2.)

Note:

1. For plastic, coating can be as short as 2 hours. For glass coverslips or for transfection, overnight.

2. If cells are to be plated onto plastic dish and not for transfection, can omit laminin from the above, and use poly-D-lysin only.

3. This method worked well for Cx, Hip. cultures on plastic in any medium.

4. Use sterile T.C. bottles to make up coating solution. However, do not let it sit too long before adding to plates, otherwise the glass bottle will be coated.

Coating glass coverslips in plates, if the above protocol does not work:

1. Thaw 1 aliquot of poly-D-lysine, add 9 ml sterile pBS to make total 10 ml, use 2 ml mixture per 35 mm plate with 5-6 of 10 mm coverslips placed firmly on the bottom of the plates. Use pasteur pipettes to push the coverslips if needed. Wrap plates in Saran wrap. Place plates at 4 degrees C overnight. Wash plates three times with sterile H2O.

2. Thaw one aliquot of laminin, add H2O to 10 ml. Use 2 ml mixture per 35 mm plate with 5-6 of 10 mm coverslips. Place plates at 4 degrees C overnight. Wash plates three times with sterile H2O.

Note: This type of method worked for SCG neuron cultures.

Preparation of Coverslips

Glass coverslips: Bellco German glass

Soak in Ethanol, 2 hours - overnight with mixing

Rinse thoroughly in dH2O, mixing and many changes.

Autoclave, dry load.

Keep sterile. Open in the hood only.

Preparation for Ice Slices

1. Decapitation and take out the brain and immerse to ice-cold neurobasal (or low calcium buffer).

2. Cut off cerebellum, and glue the brain (by super-glue) to the vibrotome block.

3. Put the block in the chamber (filled with ice cold neurobasal and ice).

4. Cut 500 um think slices.

5. Fix for 6 hours in 6% PFA/PBS.

6. Leave in 30% sucrose/PBS overnight.

7. Cut 50 um slices by microtome (or 40 um).

8. Block (BSA, PBST serum, room temperature for 1 hour.

9. Primary antibody in cold room overnight. Use mesh well.

10. Wash with PBST 6 times.

11. Secondary antibody in cold room overnight.

12. Wash with PBST 6 times.

13. Use gel-mount.

Keywords

Adenylyl cyclase (AC), amyloid precursor protein (APP), Alzheimer’s disease (AD), brain derived neurotrophic factor (BDNF), cAMP, cAMP-responsive element (CRE), CREB, extra-cellular regulated protein kinase (ERK), FE65, long-term potentiation (LTP), long-term depression (LTD), memory formation, neuroplasticity, local protein synthesis.