Materials

1. Isopentane.

2. Liquid nitrogen.

3. Model Minotome Microtome Cryostat (Damon/IEC Division) or comparable instrument.

4. Coplin staining jar or a glass slide rack (Fisher Scientific, Pittsburgh, PA).

5. Glass slides.

6. Subbed slides: Dissolve gelatin (1 g) in 1 L of hot distilled water. Cool and add 0.1 g chromium potassium sulfate. Store in refrigerator. Dip slides two to three times in the solution. Drain and dry in a vertical position. Store in a dust-free box.

7. Humidified chamber: Falcon no.1058 polystyrene Petri dish (150 x 15 mm). Line inside of dish with Parafilm and place six droplets (50 |L each) of 0.85% saline onto the Parafilm along the inner rim of the dish so as to circumscribe it. Placing the Petri dish cover on the bottom half of the Petri dish creates a humidified container.

8. Endogenous peroxidase activity blocking solution: Acid alcohol (add 1.0 mL HCl to 100 mL of 70% ethyl alcohol) or 0.1% phenylhydrazine in phosphate-buffered saline (PBS).

9. Blocking buffer: SuperBlock blocking buffer in PBS (Pierce, Rockford, IL); alternatively, use a blocking buffer consisting of 1% normal goat serum (NGS) and 1% bovine serum albumin (BSA) in PBS.

10. 10X PBS: NaCl (80 g), KCl (2 g), Na2HPO4-7H2O (11.5 g), KH2PO4 (2 g) in a final volume of 1 L.

11. Primary antibody: Rabbit antibodies to cannabinoid receptors (see Table 1).

12. Secondary antibody: HRPO-conjugated IgG goat anti-rabbit IgG (heavy and light chains, Organon Teknika Cappel, Durham, NC).

13. Antibody controls: Normal rabbit IgG control (Organon Teknika Cappel, Durham, NC) or preimmune serum from rabbit used to make the anti-cannabinoid receptor antibody; antigen added in excess to primary antibody (see Table 1).

14. Staining dishes.

15. Plastic slide box.

16. 0.2% Triton X-100/PBS: Add 200 |L Triton X-100 to 1X PBS.

17. Hanker-Yates reagent (Polysciences, Warrington, PA): Hanker-Yates mixture (7.5 mg), Tris-HCl buffer, 0.05 M, pH 7.6 (10 mL), hydrogen peroxide, 1% (0.1 mL).

18. Cresyl violet acetate (0.1%): Add 100 mg cresyl violet acetate to distilled water to make a final volume of 100 mL. Filter with a Whatman no. 2 filter paper.

19. Light microscopy aqueous mounting medium: Aquamount Aqueous Mountant (Lerner Laboratories, Pittsburgh, PA); Crystal/Mount Permanent Aqueous Mounting Medium (Biomeda Corp., Foster City, CA).

20. Permount Histological Mounting Medium (Fisher Scientific, Company, Pittsburgh, PA).

21. Large glass cover slips (No.1, 22 X 50 mm; Arthur H. Thomas Co., Philadelphia, PA).

22. 0.05 M Tris-HCl buffer, pH 7.6 (store at room temperature): Trizma HC1 (6.06 g), Trizma base (1.39 g). Bring to 1 L with distilled water.

23. 2% Formalin in PBS (50 mL): Weigh 1 g of paraformaldehyde and add to 25 mL reagent-grade water. Heat the solution to 60°C on a heating-stir plate in a chemical hood. Once the temperature has reached 60°C, immediately remove from the heat. Transfer to another stir plate and add 10 N NaOH (approximately two to three drops) until the cloudy solution clears. Allow the solution to cool. Add 20 mL of reagent-grade water and 5 mL of 10X PBS. Filter and store at 4°C for no more than 1 wk.

24. 12-mm No.1 round glass cover slips (Arthur H. Thomas Co., Philadelphia, PA), sterilized by autoclaving or soaking in 70% ethanol.

25. 150-mm Petri dishes.

26. Ethanol.

27. Xylene.

28. Equipment and facilities for paraffin embedding and tissue sectioning.

29. Slide warmer (GCA Precision Scientific, Chicago, IL).

30. Liquid release agent (Electron Microscopy Sciences, Fort Washington, PA).

31. Acetone.

32. Vibratome (Vibratome Series 2000, Technical Products International Inc., St. Louis, MO) or similar instrument.

33. Hemostat.

34. Blunt 13 G and 15 G hypodermic needles.

35. Ultramicrotome for cutting thin plastic sections.

36. Tools for removing brain (Fine Science Tools (USA), Inc., Foster City, CA): 17-cm standard tough cut scissors (no. 14110-17), spring scissors angled to side with a 10mm cutting edge (no. 15006-09), 14.5-cm straight forceps (no. 11000-14), 16-cm angled forceps (no. 11080-02), Dumont forceps (no. 11295-51), standard scalpel handle (no. 10003-12).

37: Anesthesia: Administer sodium pentobarbital intraperitoneally to mouse or rat at 60 mg/kg. Verify animal death by absence of cardiac pulse and presence of fixed/dilated pupils. Pinch animal in foot to verify absence of response.

38. Rotary mixer (Penetron Mark IIIB, Sunkay Laboratories, Inc., Tokyo, Japan).

39. Perfusion instruments.

40. Tygon tubing.

41. Peristaltic pump.

42. Avidin/Biotinylated Enzyme Complex (ABC) kit (Vector Laboratories, Burlingame, CA).

43. Glass vials with caps (Fisher Scientific, Pittsburgh, PA).

44. Camel's hairbrush.

45. Poly-L-lysine: Add 25 |L of 1 mg/mL to each sterilized no.1 cover slip in a hood and allow to stand (10 min). Rinse cover slips three times with sterile distilled water and air dry.

46. Perfusion fixative (4% paraformaldehyde and 0.1% glutaraldehyde in 0.1 M phosphate buffer): Add 20 g paraformaldehyde to 200 mL distilled water and heat to 60°C. Add 1 N NaOH until solution clears and allow solution to slightly cool. Add 125 mL 0.4 M standard phosphate buffer and cool to 20°C on ice. Add 2 mL puri fied 25% glutaraldehyde and 2 mL 0.5% calcium chloride and adjust pH to 7.2. Add distilled water to make 500 mL.

47. Perfusion prewash solution (mouse, rat): Add in the following order: 0.4 M standard phosphate buffer (50 mL), NaCl (1.8 g), 0.5% calcium chloride (0.8 g), and heparin sodium (1000 IU/mL). Adjust pH to 7.2. Add distilled water to make 200 mL.

48. 0.4 M Standard phosphate buffer, pH 7.2: Add in order sodium phosphate, monobasic (10.6 g); potassium phosphate, dibasic (56 g); distilled water (800 mL). Adjust pH to 7.2 with NaOH or HCl. Add distilled water to make 1 L.

49. 0.1 M Sodium phosphate buffer: To 100 mL distilled water add sodium phosphate (Sigma-Aldrich S-0876) (3.55 g) and 0.5 % calcium chloride (1 mL). Adjust pH to 7.2 using HCl and add distilled water to make 250 mL.

50. Double strength buffer for osmium tetroxide (0.2 M phosphate buffer with 0.04 mM CaCl2 and 14% dextrose): Dextrose (14 g), 0.4 M standard phosphate buffer (50 mL), 0.5% calcium chloride, anhydrous (0.8 mL).Add distilled water to make 100 mL.

51. Osmium solution (1.0% osmium tetroxide in 0.1 M phosphate buffer with 7% dextrose and 0.02 mM CaCL and 1.5% potassium hexacyanoferrate): Double strength buffer (5 mL), 2% osmium tetroxide solution (5 mL) (0.5 g OsO4/25 mL distilled water), potassium hexacyanoferrate (0.15 g).

52. 2% Osmium: Distilled water (25 mL, osmium tetroxide (0.5 g). Pour liquid nitrogen into a plastic cup and immerse vial of osmium to crystallize. Score the vial of osmium using a file and place into a white silicon holder. Snap vial open and put crystallized osmium into 25-mL Erlenmeyer flask with a stopper. Add distilled water to osmium under a chemical hood. Wrap Parafilm tightly around flask and stopper. Store in refrigerator. It will take at least 24 h to dissolve.

53. Sodium hydrogen maleate-NaOH buffer solutions (Sodium hydrogen maleate, NaHC4H2O4 ■ 3H2O, MW 192.11): 0.2 M solution is prepared by dissolving in distilled water 23.2 g maleic acid (requires heating), and mixing with 200 mL 1 N NaOH and diluting to 1 L with distilled water. Prepare for appropriate pH as follows: to 25 mL 0.2 M NaH maleate add x mL 0.1 M (N) NaOH and dilute to 100 mL with distilled water.

54. 0.5% Uranyl acetate (Sigma-Aldrich, St. Louis, MO): Uranyl acetate powder (50 mg), 0.05 M sodium hydrogen maleate-NaOH buffer, pH 6.0 (10 mL).

55. TAAB 812 Embedding resin: TAAB 812 (12.00 g), dodecenyl succinic anhydride (4.75 g), nadic methyl anhydride (8.25 g), DMP-30 (0.50 g). Weigh first three components into 50-mL disposable beaker using Pasteur pipets. Mix thoroughly first with stirring rod and then with magnetic stirrer for 30 min covered tightly with Parafilm. Before use, add DMP-30 to mixture, and mix thoroughly and slowly with magnetic stirrer for 30 min. Place under vacuum in a desiccator to remove air before use. Mixture may be stored in desiccator in freezer indefinitely, but allow desiccator to reach room temperature before removing.

Resin stubs: Place resin into size 00 BEEM capsules (Ted Pella, Redding, CA). Place capsules with pointed side up in a 60°C oven for 2-3 d to harden. 10% Phosphate-buffered formalin: Made up fresh from (powdered) paraformaldehyde (Sigma-Aldrich, St. Louis, MO). Add 20 g to 50 mL distilled water. Heat to 60°C with stirring and then add drops of 1 N NaOH (usually about 12 drops) until solution clears. Cool. Add 10 mL 10X PBS. Bring final volume to 100 mL with distilled water.

Glutaraldehyde (Polysciences, Warrington, PA): Bought as 25% solution. Wear gloves and dispense in a fume hood.

4% Paraformaldehyde, 0.25% glutaraldehyde in 0.1 M sodium cacodylate/HCl buffer: 10% formaldehyde (40 mL), 25% glutaraldehyde (1 mL), 0.2 M sodium cacodylate buffer (100 mL). Bring to 200 mL with distilled water. 0.1 M Sodium cacodylate/HCl: Add 2.12 g sodium cacodylate to 50 mL distilled water. Adjust pH to 7.4 with 1 N HCl and bring to a final volume of 100 mL with distilled water. Store at 4°C for as long as 6 mo.

0.2 M Sodium cacodylate/HCl: Prepare as described for 0.1 M sodium/cacodylate except that 4.24 g of sodium cacodylate are added.

Uranyl acetate, aqueous (Polysciences, Warrington, PA): Make up from powder as 0.5% aqueous solution then thoroughly mix and filter.

LR gold resin (Polysciences, Warrington, PA): Purchased as liquid resin and separate catalyst (benzoin methyl ether). The 0.1% benzoin methyl ether is added to the resin for the complete mixture.

Lead citrate (Polysciences, Warrington, PA): Made up from three stock solutions, which are: solution A) trisodium citrate (37.7g/100 mL); (solution B) lead nitrate (31g/100 mL); (solution C) NaOH (4 g/100 mL). Make up in a 1.5-mL Eppendorf tube: To 0.64 mL distilled water, add 0.12 mL solution A. Mix, then add 0.08 mL solution B and mix. To the mixture then add 0.16 mL solution C and mix. Centrifuge and dispense from top. Note: Do not exhale over lead citrate mixture, as CO2 will result in formation of lead carbonate precipitate. Modified phosphate-buffered saline (MPBS): BSA 10.0 g/L; Na2HPO4 (anhydrous) 0.524 g/L, KH2PO4 (anhydrous) 0.092 g/L; NaCl 8.76 g/L; NaEDTA 0.372 g/L; NaN3 0.2 g/L, Tween-20 500 |L/L. Mix thoroughly and adjust pH to 8.2. Filter immediately before use (0.22 |im). Note: Use as diluent for all antisera and gold probe as well as for all buffer rinsing steps. Methanol.

Propylene oxide (Polysciences Inc., Warrington, PA).

Benzoin methyl ether (Polysciences Inc., Warrington, PA).

Nickel grids (300 mesh) (Polysciences Inc., Warrington, PA; Ted Pella, Redding, PA).

Gelatin capsules (Polysciences Inc., Warrington, PA; Ted Pella, Redding, PA).

Goat anti-rabbit IgG-polygold (10 nm) (Nanoprobes Inc., Stony Brook, NY;

Aurion, 6702 AA Wageningen, The Netherlands; BBI International: distributed in

United States by Vector Laboratories Inc., Burlingame, CA).

72. Formvar (Polysciences Inc., Warrington, PA).

73. Near-ultraviolet lamp set-up (Thorn projector lamp, A1/209 FDX, 12 V 100 W).

74. 0.85% Saline: Add NaCl (8.5 g) to 100 mL distilled water and mix. Bring to a final volume of 1 L with distilled water.

75. Vecastain STAIN ABC kit (Vector Laboratories, Burlingame, CA).

76. DAB substrate kit for peroxidase (Vector Laboratories, Burlingame, CA).

3. Methods

3.1. Processing of Cells and Tissues for Light Microscopy

3.1.1. Procedure for Preparation of Cryostat Tissue Sections

1. Precool isopentane in liquid nitrogen. Dip freshly dissected tissue block (5 x 5 mm pieces) into cold isopentane. Frozen tissue blocks can be stored in sealed vials at -70°C in the presence of a few drops of isopentane to prevent drying. Caution: Do not allow either isopentane or liquid nitrogen to come in contact with skin! Do not inhale vapors!

2. Transfer frozen tissue blocks to a cryostat and allow temperature to equilibrate to -20°C for 30 min.

3. Mount the tissue block on the cryostat stub with embedding medium such as Tissue-Tek O. C. T. Compound (Sakura Finetek U.S.A. Inc., Torrance, CA) or TBS Tissue Freezing Medium (Triangle Biomedical Sciences, Durham, NC).

4. Trim surface of block.

5. Cut sections (5-20 |im) of unfixed tissue in a cabinet cryostat (-20°C ) directly onto subbed slides. Allow the sections to air dry overnight at room temperature in a dark, dust-free place. Alternatively, sections on slides may be stored at -70°C to -80°C (2-3 mo) in a sealed plastic slide box. Stored slides should be allowed to warm to room temperature before use.

6. Pretreat slides with Triton X-100 to increase permeability of tissue sections: Place slides in a rack and insert rack in PBS containing 0.2% Triton X-100. Incubate for 20 min at room temperature.

7. Wash slides in PBS (four times, 5 min each, room temperature). Make certain to remove all of the detergent.

8. Rinse slides with distilled water and allow to air dry. Do not include Triton X-100 or any other detergent in any of the subsequent steps.

9. Treat tissue sections with endogenous peroxidase inhibitor solution (4 min).

10. Rinse tissue sections in reagent quality water (5 min).

11. Treat tissue sections with 150-200 ||L blocking solution (e.g., SuperBlock) for 1 h at room temperature. If processing immune cells or tissues, include 1% NGS in the blocking buffer to block Fc receptors present on the surface of some immune cells.

12. Drain off excess blocking solution by touching edge of slide to a paper towel. Transfer slides to a humidified chamber.

13. Cover tissue slice with the primary anti-cannabinoid receptor antibody (see Table 1). It is preferable initially to use at least four dilutions of affinity purified antibody (1:10, 1:25, 1:50, 1:100) in blocking buffer. Use a 1:25 dilution of normal rabbit

IgG (1 mg/mL) in blocking solution as a control. Incubate for 1 h at room temperature in a humidified chamber. Do not let the sections dry out in this or subsequent steps.

14. Gently drain the primary antibody solution off the slides. Wash slides (five times, 5 min each) at room temperature in PBS. Use a large volume for the wash (e.g., 50 mL per slide).

15. Cover tissue slice with the secondary antibody. Initially, use a 1:32 dilution of goat IgG anti-rabbit IgG-HRPO diluted in blocking solution. Subsequent experiments may require titration of the HRPO-conjugated secondary antibody. Dilutions of 1:32-1:64 work well if the primary antibody used is directed against extracellular amine terminal domains of cannabinoid receptors, while dilutions of 1:100-1:1000 work well for antibodies directed against the entire receptors. Incubate for 1 h at room temperature in a humidified chamber at room temperature.

16. Gently drain the secondary antibody solution off the slides by touching the edge of the slide to a paper towel. Wash the slides (five times, 5 min each) at room temperature in PBS.

17. Incubate slides in 0.05 M Tris-HCl buffer, pH 7.6 for 2-3 min.

18. Develop with Hanker-Yates reagent (Polysciences, Warrington, PA) in the dark by placing a sheet of tin foil over the humidified incubation chamber. Development usually is complete by 15 min. However, intensity of development may vary depending on the primary antibody used and the amount of cannabinoid receptor present in tissue. A general approach is to place a negative control (e.g., tissue slice treated with normal rabbit IgG as the primary antibody) side by side with a positive control (e.g., tissue slice known to contain the cannabinoid receptor at a relatively high concentration such as a brain slice that contains high levels of CB1). The development reaction is terminated when the positive control qualitatively exhibits a twofold level of staining at the macroscopic level. Note: A metallic layer may appear on the surface of the reaction solution. If this occurs at the termination of the reaction period, gently flush this layer off the surface with 0.05 M Tris-HCl buffer, pH 7.6. Do not allow the metallic layer to come in contact with the tissue, as it will form precipitate.

19. Wash slides in 0.05 M Tris-HCl buffer, pH 7.6, for 2-3 min.

20. Rinse slides briefly in distilled water. Note: Slides may be counterstained with 0.1% cresyl violet acetate (2-3 min for cells on a cover slip; 10-20 min for a tissue section) at this point.

21. Remove excess water around the tissue section by touching edge of slide to a paper towel. Proceed below to step 22 for mounting in aqueous mounting medium or to step 23 for mounting in inorganic medium.

22. Aqueous medium mounting: Mount with Aquamount under a large coverslip. Allow to harden overnight in a refrigerator. If using Crystal/Mount, do not add a coverslip. Instead, add three drops of the Crystal/Mount directly onto the tissue section. Rotate the slide so as to distribute the medium over the entire tissue. Place in a horizontal position in an oven at 40-50°C (30 min). Remove slides and allow to cool. Examine directly under the light microscope.

23. Organic medium mounting: Dehydrate the sections by 5-min washes with graded concentrations of ethanol (70%, 95%, and two changes of absolute ethanol). Immerse slides in 100% xylene (two changes, 5 min each). Apply a large cover slip (22 x 50 mm) using a few drops (e.g., 25-50 ||L) of permanent mounting medium suitable for light microscopy such as Permount (Fisher Scientific, Pittsburgh, PA).

3.1.2. Procedure for Processing Formalin-Fixed and Paraffin-Embedded Tissue

3.1.2.1. Processing of Brain Tissue

The following protocol is designed for either murine or rat brain but can be adapted for other tissues.

1. Store tissue in 10% phosphate-buffered formalin (freshly prepared from paraformaldehyde at room temperature for 7 d, after which the formalin is removed and replaced with 70% ethanol. Tissue may be stored in 70% ethanol for several weeks. Note: Transfer to 70% ethanol is recommended to minimize loss of anti-genicity in tissues owing to prolonged exposure to formalin.

2. Antecedent to processing for paraffin embedding, transfer tissue to fresh 10% phosphate-buffered formalin (two times, 45 min each).

3. Dehydrate tissue by immersion through the following series of solutions (30 min each): 80% ethanol (once), 95% ethanol (two times), 100% ethanol (three times).

4. Immerse tissue in xylene (two 45-min incubations) in order to clear the tissue and allow paraffin infiltration.

5. Immerse tissue four times in hard paraffin (melting point = 56-57°C), which is heated and maintained at 60°C. Embed in paraffin.

3.1.2.2. Processing of Paraffinized Tissue Sections for Immunoperoxidase Staining

1. Cut sagittal sections from the right or left hemispheres for each brain starting from the center (middle) out. Alternatively, coronal sections may be cut. Seven sections 3 |im thick are mounted onto clean subbed glass slides. Number the slides sequentially according to the tissue label (coded). The first and last slides are processed further and stained using hematoxylin and eosin to serve for histological assessment and for orientation of brain sections.

2. Make certain that slides containing tissue sections are dry.

3. Immerse slides in the following solutions for the indicated periods of time. Slides can be inserted into a Coplin jar for each treatment. Move slides gently up and down two to three times after immersing into each solution.

c. Absolute ethanol (2-3 min).

4. Wipe excess PBS from slides by touching edge of slide to a paper towel.

5. Treat tissue sections with endogenous peroxidase inhibitor solution (4 min).

6. Rinse sections in reagent-quality water (5 min).

7. Transfer slides to a humidified chamber and treat with 150-200 |L blocking solution (e.g., SuperBlock) for 1 h at room temperature. If processing immune cells or tissues, supplement blocking buffer with 1% NGS in order to block Fc receptors present on the cell surface of some immune cells.

8. Drain off excess blocking solution.

9. Cover tissue slice with the primary rabbit IgG anti-cannabinoid receptor antibody. It is preferable initially to use at least four dilutions of each affinity-purified antibody (1:10, 1:25, 1:50, 1:100) in blocking buffer. Use a 1:25 dilution of normal rabbit IgG (1 mg/mL) in blocking solution as a negative control. Incubate for 1 h at room temperature in a humidified chamber. Do not let the sections dry out in this or subsequent steps. Ideally, a negative control should consist of preimmune IgG derivative from the same rabbit which was used to produce the anti-cannabinoid receptor antibody. However, when using commercially available anti-cannabinoid receptor antibodies, such preimmune IgG may not be available and normal rabbit IgG can be used as a control.

10. Gently drain the primary antibody solution off the slides. Wash slides (five times, 5 min each) at room temperature in PBS. Use a large volume for the wash (e.g., 50 mL/slide).

11. Cover tissue slice with secondary antibody. Use a 1:32 dilution of goat IgG anti-rabbit IgG-HRPO (Organon Teknika Cappel, Durham, NC) diluted in blocking solution. Incubate for 1 h at room temperature in a humidified chamber at room temperature.

12. Gently drain the secondary antibody solution off the slides by touching the edge of the slide to a paper towel. Wash the slides (five times, 5 min each) at room temperature in PBS.

13. Transfer slides to a Coplin jar and incubate with 0.05 M Tris-HCl buffer, pH 7.6 for 2-3 min.

14. Transfer slides to a humidified chamber and develop with Hanker-Yates reagent in the dark. This can be achieved by placing paper towels over the humidified incubation chamber. Development usually is complete by 15 min. However, intensity of development may vary depending on the primary antibody used and the amount of cannabinoid receptor present in tissue. A general approach is to place a negative control (e.g., tissue slice treated with normal rabbit IgG as the primary antibody) side by side with a positive control (e.g., tissue slice known to contain the cannabinoid receptor at a relatively high concentration). The development reaction is terminated when the positive control qualitatively exhibits a twofold level of staining at the macroscopic level. Note: The intensity of reaction product also can be monitored by placing a positive control slide on an inverted microscope.

15. Wash the slides in 0.05 M Tris-HCl buffer, pH 7.6, for 2-3 min.

16. Rinse the slides briefly in distilled water.

17. Immerse the slides sequentially in the following solutions for the indicated periods of time:

d. Absolute ethanol (2-3 min).

18. Mount in Permount Organic Medium (Fisher Scientific, Pittsburgh, PA). Apply a large cover slip (22 x 50 mm) using a few drops (25-20 | L) of Permount. Allow Permount to harden on a slide warmer.

19. Examine under the light microscope and photograph using Kodak Elite Chrome 160T Tungsten 35-mm film (Eastman-Kodak Company, Rochester, NY). Alternatively, images may be obtained using a digital camera system such as the Spot RT Slider Digital Camera (Diagnostic Instruments Inc., Sterling Heights, MI). Figure 1 shows results obtained for identification of the CB1 cannabinoid receptor in rat brain fixed in formalin and embedded in paraffin.

3.1.3. Procedure for Processing of Cells Maintained in Culture for Demonstration of Intracellular Cannabinoid Receptors

1. Seeding of cells and cell culture:

a. For adherent cells, grow cells on sterile coverslips which have been placed (four cover slips/well) in 6-well tissue culture dishes (Falcon No. 3046, Becton Dickinson Labware, Franklin Lakes, NJ). Allow cells to reach approx 90% confluency. Alternatively, place coverslips in a 60-mm Petri dish. Add a 100-|L volume of cell suspension (1 x 106 cells/mL in culture medium) to each cover slip so as to form a bubble confined to the cover slip. Place cover on Petri dish and place in a humidified CO2 incubator maintained at 37°C for 2 h or until cells attach to the cover slip. Attachment of cells can be monitored by screening the Petri dish with its contained cover slips on an inverted microscope.

b. For nonadherent cells, apply 10-20 |L of cell suspension (1 x 106 cells/mL) to each coverslip which has been precoated with poly-L-lysine. Allow to sit 10 min, then proceed with fixation. Alternatively, cells in suspension can be attached to cover slips using a cytocentrifuge according to the manufacturer's instructions. A StatSpin Cytofuge 2 Cytocentrifuge (StatSpin Technologies, Norwood, MA) works well. Typically, one adds 100 | L of a cell suspension

Fig. 1. Light micrographs depicting the distribution of CB1 cannabinoid receptor immunoperoxidase staining in rat brain. (A) Immunoreactive labeling in the cerebellum is concentrated in the molecular (m) layer while minimal labeling is identified in the granular (g) layer. Immunoreactive product is absent also from the white matter (w). The arrow denotes the Purkinje cell layer. (B) Localization of immunoreactive labeling in the Purkinje cell layer. Intense labeling is localized within areas (arrow) that form arborizations into the molecular layer (m). Cells with morphology consistent with that of basket cells show intense immunoreactivity for the CB1. (C) Immunoreactive labeling for the CB1 receptor is localized as intense clusters (arrows) within processes forming parallel arrays extending through the molecular layer. (D) Localization of immunoreactive product for the CB1 receptor within the cytoplasm of individual cells (arrow) within the amygdala. Scale bars: A, 100 |im; B, 50 |im; C, 25 |im; D, 10 |im.

(1 x 105 - 1 x 106 cells/mL) to each cell concentrator unit for spinning down onto a glass slide.

2. Remove cover slips from tissue culture well and drain off excess medium by touching edge of cover slip to a paper towel.

3. Rinse gently (three times) in PBS (room temperature) and air dry for 30 min.

4. Fix cover slips in absolute acetone for 5 min.

5. Air dry cover slip cultures for 30 min.

6. Store cover slips in a Petri dish in a desiccator (with desiccant) at 41° C long term (e.g., up to 2 yr) or leave in Petri dish at room temperature short term (e.g., up to 6 mo).

7. Prewet cover slips by immersion in PBS (5 min).

8. Treat cover slips with endogenous peroxidase inhibitor solution (4 min).

9. Rinse cover slips in reagent-quality water (5 min).

10. Treat cover slips with 50 |L blocking solution (e.g., SuperBlock) for 1 h at room temperature. Note: Supplement blocking solution with 1% NGS if processing immune tissues or cells in order to block for Fc receptors that are present on surfaces of some immune cells.

11. Drain off excess blocking solution by touching edge of cover slip to a paper towel.

12. Cover cover slips with the primary rabbit anti-cannabinoid receptor antibody. It is preferable initially to use at least four dilutions of each affinity purified antibody (1:10, 1:25, 1:50, 1:100) in blocking buffer. Use a 1:25 dilution of normal rabbit IgG (1 mg/mL) in blocking solution as a negative control. Incubate for 1 h at room temperature in a humidified chamber. Do not allow the sections to dry out in this or subsequent steps.

13. Gently drain the primary antibody solution off the cover slips by touching the edges to a paper towel. Wash cover slips (five times, 5 min each) at room temperature in PBS. Use a large volume for the wash (e.g., 50 mL).

14. Add 25 |L secondary antibody goat IgG anti-rabbit IgG (heavy and light chains)-HRPO to cover slips. Use a 1:32 dilution of goat anti-rabbit HRPO (1

mg/mL) diluted in blocking solution. Incubate for 1 h at room temperature in a humidified chamber at room temperature.

15. Gently drain the secondary antibody solution off the cover slips by touching edge to a paper towel. Wash the cover slips (five times, 5 min each) at room temperature in PBS.

16. Immerse cover slips in 0.05 M Tris-HCl buffer, pH 7.6 for 2-3 min.

17. Develop with Hanker-Yates reagent. Development usually is complete by 15 min. However, intensity of development may vary depending on the primary antibody used and the amount of cannabinoid receptor present in tissue. A general approach is to place a negative control (e.g., cells on a cover slip treated with normal rabbit IgG as the primary antibody) side by side with a positive control (e.g., cells trans-fected with a cannabinoid receptor expression construct). The development reaction is terminated when the positive control qualitatively exhibits a twofold level of staining at the macroscopic level. Alternatively, the staining reaction can be monitored under an inverted cell culture microscope.

18. Wash cover slips in 0.05 M Tris-HCl buffer, pH 7.6, for 2-3 min.

19. Rinse cover slips briefly in distilled water.

20. Remove excess water by touching edge of cover slip to a paper towel. Proceed to step 21 for mounting in aqueous mounting medium or to step 22 for mounting in inorganic medium. Note: Mounting with AquaMount requires a small amount of residual water to allow medium to harden. Do not air dry for Permount or Aquamount. If mounting in Permount, must go from water to alcohols to xylene without interruption.

21. Aqueous medium mounting: Mount (25 ||L) with Aquamount under a 12-mm circular cover slip if cells have been spun down onto a glass slide. If cells are adherent to the cover slip, add 25 | L directly onto a glass slide and place the cover slip cell-side down onto the droplet.

22. Organic medium mounting: Dehydrate the cells on the cover slips by 5-min washes with graded concentrations of ethanol (70%, 95%, and two changes of absolute ethanol). Transfer to 100% xylene (two changes, 10 min each). If cells have been spun down onto a glass slide, apply a cover slip using a few drops (e.g., 25 | L) of permanent mounting medium suitable for light microscopy such as Permount (Fisher Scientific, Pittsburgh, PA). If cells are adherent to the cover slip, add 25 |L directly onto a glass slide and place the cover slip cell side down onto the droplet.

3.1.4. Procedure for Processing of Cells Maintained in Culture for Demonstration of Cell Surface Cannabinoid Receptors

1. Prepare cell cultures on cover slips and harvest for immunoperoxidase staining as described in steps 1-3 in Subheading 3.1.3. Note: Trypsinization of cell mono-layers should not be considered since this process may remove cannabinoid extracellular domains from cell surfaces.

2. Fix in 2% formalin in PBS for 20 min.

3. Rinse cover slips in four to five changes of PBS for 10 min each. Do not allow cells on cover slips to dry out at any point.

4. Proceed with steps 10-22 as described in Subheading 3.1.3.

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