Characterizing Synaptic Vesicle Proteins Using Synaptosomal Fractions and Cultured Hippocampal Neurons

Materials

• 3- to 4-week old male rat (Wistar or Sprague-Dawley)

• 1× sucrose buffer (see recipe), ice cold

• Percoll gradients (see recipe), ice cold

• Wash buffer (see recipe), ice cold

• Medium L (see recipe)

• 1 M KOH

• 1.0 M sucrose in Medium L (store up to 5 days at 4°C)

• Sucrose gradients (see recipe) in thin-walled ultracentrifuge tubes (ultraclear tubes, 38.5-ml capacity, 25 × 89 mm)

• Protease inhibitors (see recipe)

• 20 mM Tris·Cl, pH 7.4 (APPENDIX 2A)

• Antibodies specific for subcellular fractions: e.g., anti-synaptophysin (Chemicon) or anti-VAMP (StressGen) for synaptic vesicles, anti-Na⁺/K⁺-ATPase (Transduction Laboratories) for plasma membranes, and anti–lactate dehydrogenase (anti-LDH, Abcam) for cytosol

• Rat guillotine

• Surgical tools for isolation of cortex

• 25- and 55-ml Potter-Elvehjem homogenizers (Teflon-glass)

• High-speed centrifuge (e.g., Sorvall RC-5C with SS-34 rotor and adaptors for eight 16 × 100–mm tubes)

• 50-ml (29 × 102–mm) polycarbonate high-speed centrifuge tubes

• Ultracentrifuge capable of 100,000 × g, with:

  • Swinging bucket rotor (e.g., Beckman SW28) and 31-ml, 25 × 89–mm, thick-walled polycarbonate ultracentrifuge tubes

  • Fixed-angle rotor (e.g., Beckman 70.1 Ti or 50 Ti) with 10-ml, 16 × 76–mm thick-walled polycarbonate ultracentrifuge tubes

• 10,000 MWCO dialysis membranes

• Centriprep-10 centrifugal concentrators (Millipore)

• Additional reagents and equipment for rat anesthesia (APPENDIX 4B), dialysis, determining protein concentration (e.g., Simonian and Smith, 2006), SDS-PAGE (e.g., Gallagher, 2006), and immunoblotting (e.g., Gallagher et al., 2008)

NOTE: All isolation steps must be performed at 0° to 4°C, and all solutions, centrifuge tubes, and centrifuge rotors should be precooled below 4°C and kept on ice.

Isolate synaptosomes

• 1

  Anesthetize rat (APPENDIX 4B) and decapitate with a guillotine. Remove brain and place in ice-cold 1× sucrose buffer.

  A total of four to eight rats are recommended to prepare a concentrated synaptosome fraction for biochemical studies.

  EDTA may be omitted when attempting to characterize Ca²⁺-dependent proteins that co-fractionate with synaptic vesicles.

• 2

  Remove cerebellum, brain stem, midbrain, and white matter (Fath et al., 2009; Fig. 2.7.1). Pool four cortices together and place in 9 ml ice-cold 1× sucrose buffer.

• 3

  Transfer to a 25-ml Potter-Elvehjem homogenizer and homogenize the cortex using ten slow up-and-down strokes of the homogenizer.

• 4

  Centrifuge the homogenate 10 min at 1000 × g (3000 rpm in an SS-34 rotor), 4°C.

• 5

  Collect supernatant and carefully load 2 ml supernatant onto each of four Percoll gradients. Centrifuge gradients for exactly 5 min at 32,500 × g (17,250 rpm in an SS-34 rotor), 4°C, using a timer to start counting the time when 17,250 rpm is reached. Collect synaptosomes from the interface at 15%/23% using a Pasteur pipet.

  Alternatively, the supernatant may be loaded onto a discontinuous sucrose gradient (see recipe) consisting of 1.2/1.0/0.8 M sucrose layers and centrifuged for 20 min at 48,200 × g (Cohen et al., 1977). In that case, the synaptosome fraction will be collected from the interface between the 1.2 and 1.0 M sucrose layers.

  Synaptosomes are visible as a cloud of material concentrated at the bottom interface (15%/23% for Percoll or 1.2/1.0 M for sucrose). Use a separate Pasteur pipet to remove upper layers before collecting the synaptosome layer.

• 6

  Transfer the synaptosomes to a 50-ml polycarbonate centrifuge tube and add 30 ml chilled wash buffer. Centrifuge 15 min at 15,000 × g (11,000 rpm in an SS-34 rotor), 4°C.

• 7

  Carefully remove the supernatant and proceed immediately with fractionation (below) or immunogold labeling (see Basic Protocol 2).

  Synaptosomes form a loose pellet. If a more firm pellet is desired, resuspend the pellet in 20 ml cold wash buffer and repeat centrifugation before proceeding with the analysis.

  Both fractionation and immunogold labeling require freshly isolated synaptosomes.

Fractionate synaptosomes

The fractionation procedure is illustrated in Figure 2.7.2.

• 8

  Lyse synaptosomes by resuspending the pellet in ice-cold hypoosmotic Medium L. For biochemical studies, combine synaptosomal preparations from six to eight rats in a single fraction and resuspend in a volume of 4 ml Medium L per rat.

• 9

  Save a 0.5-ml aliquot of this synaptosomal suspension for validation of the fractionation procedure after purification (steps 18 to 21).

• 10

  Manually homogenize four times slowly up and down in a 55-ml Potter-Elvehjem homogenizer, then incubate suspension on ice for 15 min.

• 11

  Repeat homogenization once more, then adjust pH to 8.0 with 1 M KOH, and incubate homogenized synaptosomal suspension on ice for another 45 min.

• 12

  Layer 5 ml synaptosomal suspension onto 5 ml of 1.0 M sucrose in Medium L in a 31-ml thick-walled polycarbonate ultracentrifuge tube (using a total of six to eight tubes). Centrifuge 30 min at 96,300 × g (27,000 rpm in an SW28 rotor), 0° to 4°C. Collect the supernatant in fresh 31-ml ultracentrifuge tubes.

  The pellet is a crude synaptosomal mitochondrial fraction. For further biochemical analysis, it can be resuspended in 20 mM Tris·Cl (pH 7.4) containing protease inhibitors and stored up to 1 year at −20°C.

• 13

  Cover the supernatant tubes with Parafilm and gently mix by inverting (do not vortex). Centrifuge 14 hr at 25,000 × g (14,000 rpm in an SW28 rotor), 4°C. Collect the supernatant (the cytosolic protein fraction) and place both it and the pellet on ice.

Analyze protein distribution in fractions

• 18

  Determine protein concentration of each fraction: synaptosomes (step 7), cytosol (step 17a), plasma membranes (step 17b), and synaptic vesicles (step 17b).

• 19

  Analyze aliquots containing equal amounts of protein from each fraction by SDS-PAGE.

  Aliquots containing 5–10 μg total protein should be sufficient to detect most synaptic vesicle proteins.

• 20

  Detect protein(s) of interest by immunoblotting with the appropriate specific antibodies.

• 21

  Determine relative purity of each fraction by sequential immunoblotting with antibodies against well-characterized markers (see Fig. 2.7.3).

Materials

• Unfractionated rat brain synaptosomes (see Basic Protocol 1, step 7)

• Wash buffer (see recipe)

• 0.1 M sodium phosphate buffer, pH 7.4 (see recipe)

• 4% (w/v) paraformaldehyde in 0.1 M sodium phosphate buffer

• PBS (APPENDIX 2A)

• 5% (v/v) normal goat serum/0.1% (w/v) saponin in PBS

• 5% (w/v) nonfat dry milk in PBS

• Primary antibody against synaptic vesicle protein of interest

• Nanogold-conjugated secondary antibody (Nanoprobes)

• 2% (w/v) glutaraldehyde in PBS

• HQ silver enhancement kit (Nanoprobes)

• Filter paper

• Razor blade

• 12-well tissue culture plates

• Additional reagents and equipment for preparing samples for electron microscopy(UNIT 1.2)

NOTE: All steps are performed at room temperature.

Fix synaptosomes

• 1

  Resuspend synaptosome pellet purified from one rat cortex in 1 ml wash buffer, divide into four equal volumes, and microcentrifuge 5 min at maximum speed.

  Use freshly prepared synaptosomes only.

  Synaptosomes from one rat cortex should be divided into a minimum of four samples, as fixation has limited penetration into the pellet.

  When using cultured neuronal cells, omit this step. Instead, remove culture medium from cells, wash cells with 0.1 M sodium phosphate buffer, then proceed to step 2.

• 2

  Discard supernatant and fix pellet by adding 0.5 ml of 4% paraformaldehyde in 0.1 M sodium phosphate buffer and incubating for 30 min.

• 3

  Wash fixed pellet four times, each time for 5 min with 1 ml of 0.1 M sodium phosphate buffer.

• 4

  Place each pellet on filter paper soaked with 0.1 M sodium phosphate buffer and divide into four to six pieces using a razor blade (total 16–24 pieces). Transfer the pieces to separate wells of 12-well tissue culture plates.

Label synaptic vesicle proteins

• 5

  Block nonspecific binding sites and permeabilize synaptosomes by adding 1 ml of 5% normal goat serum/0.1% saponin in PBS to each well and incubating for 1 hr.

  Normal serum should come from the same species as the gold-labeled secondary antibody. The Nanogold antibody used in this protocol is from goat.

• 6

  Dilute the desired primary antibody as appropriate in 5% normal goat serum/0.1% saponin/PBS. Add 0.5 ml diluted primary antibody to each well and incubate for 1 hr.

  Antibody may be incubated for a longer time period if done at a lower temperature (e.g., 4°C). The dilution and incubation time will need to be determined empirically for each antibody used.

• 7

  Wash four times, each time for 5 min with 1 ml of 5% nonfat dry milk in PBS.

• 8

  Dilute Nanogold-labeled secondary antibody 1:400 in 5% nonfat dry milk in PBS. Add 400 μl diluted secondary antibody to each well and incubate for 1 hr.

• 9

  Wash as in step 7.

• 10

  Post-fix by adding 1 ml of 2% glutaraldehyde in PBS to each well and incubating for 30 min.

• 11

  Wash four times, each time for 5 min with 1 ml PBS.

• 12

  Wash four times, each time for 5 min with distilled water.

• 13

  Add 1 ml of silver enhancement solution to each well and incubate for 4 min.

• 14

  Immediately wash four to six times for a total of 10 min, each time with 1 ml water.

• 15

  Wash three times, each time for 5 min with 1 ml of 0.1 M sodium phosphate buffer.

Process samples for EM

• 16

  Process for electron microscopy as described in UNIT 1.2, starting with osmium tetrox-ide treatment, but use 1 ml of 0.2% (instead of 1%) osmium tetroxide in 0.1 M sodium phosphate buffer.

  Figure 2.7.4 shows immunogold electron micrographs of synaptosomes isolated from adult rat brain.

Materials

• Hippocampal neurons cultured on coverslips (see Support Protocols 1 and 2)

• Phosphate-buffered saline (PBS; APPENDIX 2a), pH 7.4

• Fixative solution: 4% (w/v) paraformaldehyde/4% (w/v) sucrose in PBS

• Blocking/permeabilization buffer: 10% (v/v) normal goat serum/0.2% (v/v) Triton X-100 or saponin in PBS (prepare fresh)

• Primary antibody against protein of interest

• Antibody dilution buffer: 10% (v/v) normal goat serum/0.05% (v/v) Triton X-100 or saponin in PBS (prepare fresh)

• Fluorophore-conjugated secondary antibody (e.g., TRITC, FITC, Cy2, Cy3, or Cy5; store protected from light)

• Anti-fading mounting medium (e.g., Fluoroshield, Accurate Chemical & Scientific Corp.)

• 12-well tissue culture plates

• Microscope slides

• Forceps

• Fluorescence microscope with 40× and/or 63× oil-immersion objectives

NOTE: Fluorophores are light-sensitive reagents. All solutions and stained samples should be protected from light, e.g., using aluminum foil.

NOTE: All steps are performed at room temperature.

Fix cells

• 1

  Transfer each coverslip containing cultured hippocampal neurons to 1 ml PBS in an individual well of a 12-well tissue culture dish.

• 2

  Aspirate PBS, add 1 ml fixative solution to each well, and incubate 10–15 min.

  Alternatively, cells can be fixed with ice-cold methanol for 10 min at −20°C.

• 3

  Remove fixative and wash coverslips three times, each time for 5 min with 1 ml PBS, pH 7.4.

• 4

  Remove PBS, add 1 ml blocking/permeabilization buffer to each well, and incubate for 1 hr to block nonspecific sites and permeabilize the cells.

  Triton X-100 is commonly used as detergent in this step; however, saponin is the preferred alternative for some proteins, especially plasma membrane proteins.

Immunolabel synaptic vesicle proteins

• 5

  Dilute primary antibody as appropriate in antibody dilution buffer. Add 400 μl diluted primary antibody to each well and incubate 2–3 hr.

  Incubation may be extended to overnight at 4°C. Use the same detergent in the dilution buffer as used in the blocking/permeabilization buffer. The optimal antibody dilution and incubation time will need to be determined empirically for each antibody used.

• 6

  Remove antibody and wash coverslips as in step 3.

• 7

  In a microcentrifuge tube, dilute fluorophore-conjugated secondary antibody as appropriate in a volume of 400 μl antibody dilution buffer per coverslip, and then microcentrifuge 5 min at maximum speed to remove fluorophore aggregates.

  Commercial preparations are typically diluted at 1:100 to 1:500.

• 8

  Add 400 μl diluted secondary antibody to each coverslip and incubate for 1–2 hr.

• 9

  Remove antibody and wash coverslips as in step 3. After last wash, add 1 ml PBS to each coverslip.

Mount samples

• 10

  Remove PBS from the coverslips, add 1 ml water, and leave for 10 min before mounting.

• 11

  Label microscope slides and place 1 drop of mounting medium on each slide.

• 12

  Using forceps, pick up one coverslip and gently blot off excess water by touching a corner to the edge of a Kimwipe. Carefully place the coverslip cell-side-down onto the drop of mounting medium on a slide.

• 13

  Gently blot mounted coverslip with paper towel. Allow to air dry.

  Fixed, immunostained, and mounted samples can be stored in the dark for 6 months to 1 year at 4°C.

  Samples are ready to be viewed under a fluorescence microscope with a 40× or 63× oil-immersion objective. Figure 2.7.5 shows an immunofluorescence photomicrograph of cultured hippocampal neurons.

Materials

• Xylenes

• 100% (v/v) ethanol

• Polyornithine (Sigma P4957)

• Ultrapure water

• Phosphate-buffered saline (PBS; APPENDIX 2A)

• Fibronectin (Sigma F1141-5MG)

• P0 mouse pups or P2-3 rat pups

• Hank’s balanced salt solution (HBSS; see recipe)

• HBSS containing 20% fetal bovine serum (FBS), ice cold, prepare fresh

• 50 mg/ml trypsin (type XI, Sigma, T1005) in H₂O (store 10-mg aliquots at −20°C)

• 20 mg/ml DNase (type IV, Sigma, D5025) in H₂O (store 1-mg aliquots at −20°C)

• Digestion buffer (see recipe), ice cold

• Dissociation buffer: 150 mg MgSO₄·7H₂O in 50 ml HBSS (final 12 mM), ice cold, prepare fresh

• Plating medium (see recipe), 37°C

• Feeding medium 1 (see recipe) with 3 μM and 6 μM Ara-C, 37°C

• 12- or 25-mm-diameter round coverslips

• Sonicator

• Gauze pads

• Glass Petri dishes

• 250°F oven

• Dissecting tools:

  • Spring scissors, straight and angled

  • Forceps, straight and curved

  • Angled dissecting bone chisel

  • Scalpel

  • Straight surgical scissors

• 35- and 60-mm tissue culture dishes

• Dissecting microscope with fiber-optic lights

• 9-in. borosilicate glass pipets (Pasteur pipets)

• Plastic transfer pipets

• 15-ml polypropylene (not polycarbonate) conical centrifuge tubes

• 0.22-μm filter

• Fire-polished, sterile, siliconized/plugged Pasteur pipets, 0.5- and 0.75-mm bore size

Clean and coat coverslips

• 1

  Wash 12- or 25-mm round coverslips by sonicating in xylenes for ~30 min.

  CAUTION: Perform this step in a chemical fume hood, and dispose of xylenes (including ethanol washes below) in appropriate waste containers.

• 2

  Remove coverslips from xylenes and rinse two to three times with 100% ethanol to remove residual xylenes.

• 3

  Sonicate coverslips for 1 hr in 100% ethanol, rinse in ethanol, and repeat for a total of 3 times.

• 4

  Rinse at least 5 times with distilled, deionized water.

• 5

  Separate coverslips manually and allow them to air-dry on clean gauze pads.

• 6

  Transfer coverslips to a glass Petri dish and bake at 250°F for at least 2 hr, preferably overnight.

• 7

  Dilute polyornithine 1:6 in sterile ultrapure water. Incubate coverslips in polyornithine in individual 35-mm dishes for 2–6 days in a sterile container shielded from light (an incubator works well).

• 8

  One day before dissections, remove the polyornithine solution and rinse coverslips three times with PBS.

• 9

  Dilute fibronectin 1:1000 in ultrapure water. Incubate coverslips in fibronectin solution in 35-mm dishes for 6–24 hr.

• 10

  Just before plating cells, rinse coverslips twice with ultrapure water.

Dissect tissue

• 11

  Decapitate a rat or mouse pup using sharp scissors. Holding the head tightly with one hand, carefully cut through the scalp with spring scissors from the point of decapitation towards the orbits. Use angled spring scissors to cut through the skull and expose the brain.

• 12

  Using curved forceps, carefully remove the brain and place in a 60-mm tissue culture dish containing 8 ml ice-cold HBSS/20% FBS.

• 13

  Using a dissecting chisel, hemi-dissect the brain and remove the brainstem and thalamus to expose the hippocampus. Under a dissecting microscope, remove the hippocampus, leaving some of the subiculum attached.

  Tug and pull as little as possible. Always cut, don’t crush the tissue, and be careful not to stab the tissue at any point.

• 14

  Using the back end of a Pasteur pipet or a cut plastic transfer pipet for the gentlest handling, transfer hippocampi to a second 60-mm dish containing 8 ml ice-cold HBSS/20% FBS.

• 15

  Using straight forceps, peel off all adhering membranes and blood vessels. Cut away the fimbriae with a scalpel or scissors, and transfer the cleaned hippocampi to a third dish containing 8 ml ice-cold HBSS/20% FBS.

• 16

  Unroll the dentate gyrus. Use the subiculum to pin down the tissue while unrolling and cutting off the dentate gyrus with scissors. Then, cut away the hippocampi from the remaining subiculum and transfer the hippocampi to a fourth 60-mm dish containing 8 ml ice-cold HBSS/20% FBS.

• 17

  Cut hippocampi into approximately ten thin slices and place on ice. Repeat steps 11–17 until all dissections are complete.

  Each dissociated hippocampus yields ~5 × 10⁵ cells. It is important to complete the dissection for one animal before proceeding to the next, and to keep the tissue pieces on ice.

• 18

  Under sterile conditions, use a plastic pipet to transfer the sliced hippocampal tissue to a 15-ml polypropylene conical tube. Let the tissue settle and discard the supernatant.

• 19

  Wash once with 10 ml HBSS/20% FBS, allow tissue to settle to the bottom of the tube, and remove the supernatant. Repeat wash three more times with 10 ml HBSS alone.

Digest tissue

• 20

  Add 200 μl of 50 mg/ml trypsin (10 mg) and 50 μl of 20 mg/ml DNase (1 mg) to 2 ml digestion buffer and filter-sterilize the solution directly onto the last pellet (step 19) using a 0.22-μm filter. Incubate for 5 min at 37°C, occasionally shaking gently.

• 21

  Discard solution and stop digestion by adding ~10 ml HBSS/20% FBS.

• 22

  Wash once with 10 ml HBSS/20% FBS and once with 10 ml HBSS alone, as above.

Dissociate cells

• 23

  Add 50 μl of 20 mg/ml DNase (1 mg) to 2 ml dissociation buffer and filter-sterilize onto the last pellet as above.

• 24

  Mechanically dissociate cells by gently triturating with two fire-polished, sterile, siliconized/plugged Pasteur pipets with successively smaller bore sizes (0.75 and 0.5 mm). Avoid bubbles and always pipet onto the wall of the tube. Allow clumps of undissociated tissue to settle for 2 min, then transfer the supernatant to another tube.

  This step separates suspended cells from tissue particles.

  CAUTION: Tips that are too small will lyse the cells; tips that are too large will not dissociate cell aggregates.

• 25

  Add 3 ml HBSS/20% FBS to the cell suspension (5 ml total), and centrifuge for 10 min at 200 × g, 4°C.

• 26

  Discard supernatant and resuspend pellet in prewarmed (37°C) plating medium. Determine cell concentration using a hemacytometer.

Plate and maintain cells

• 27

  Place drops of suspension containing ~1 × 10⁵ cells individually onto 12-mm polyornithine/fibronectin-coated coverslips in separate 35-mm dishes. Incubate 1–2 hr to allow the cells to adhere.

  The volume of suspension should be between 50 μl (to avoid evaporation) and 200 μl.

  The first short cell processes should be visible within this time.

• 28

  Add 2 ml prewarmed plating medium to each dish and incubate for 48 hr.

• 29

  Change plating medium to selection medium by replacing 1 ml old medium with 1 ml fresh pre-equilibrated feeding medium 1 containing 6 μM Ara-C.

  For the first change of medium, the concentration of Ara-C is doubled, because it will be diluted by half in the plating medium. All subsequent feedings are done with medium containing 3 μM Ara-C.

• 30

  Feed cells twice a week by replacing one third or one half of old medium with fresh pre-equilibrated feeding medium 1.

  Cells can be maintained in culture for up to 3 weeks. Always keep neurons covered with medium, as they are extremely sensitive and will die if exposed even briefly.

Materials

• 18- or 19-day pregnant rat or mouse

• 70% (v/v) ethanol

• Dissection buffer (see recipe), ice-cold

• Papain solution (see recipe)

• 1/1 solution (see recipe)

• 10/10 solution (see recipe)

• Plating medium 2 (see recipe), 37°C

• Feeding medium 2 (see recipe), 37°C

• Dissecting tools:

  • Straight wide-tipped forceps

  • Straight surgical scissors

  • Straight fine scissors

  • Forceps, curved and straight

  • Spring scissors, straight and angled

  • Dumont forceps (#5/45, 45° angled tip)

• Dissecting microscope and fiber-optic lights

• 35- and 100-mm tissue culture dishes

• 15-ml polypropylene (not polycarbonate) conical centrifuge tubes

• 5-ml pipet

• Polyornithine/fibronectin-coated 12- or 25-mm-diameter round coverslips (see Support Protocol 1, steps 1–10)

• Additional reagents and equipment for rodent anesthesia and euthanasia (APPENDIX 4B & 4H) and counting cells with a hemacytometer (APPENDIX 3B)

Isolate embryos

• 1

  Anesthetize pregnant rat or mouse with CO₂ or by another approved method, then euthanize by decapitation.

• 2

  Sterilize abdomen with 70% ethanol.

• 3

  Gripping the skin with straight wide-tipped forceps, cut the skin and muscle along the midline with surgical scissors.

• 4

  Make two cuts to the sides and pull out both horns of the uterus. Cut the uterus and transfer to a 100-mm tissue culture dish.

• 5

  Open the uterus with straight fine scissors. Remove the embryos and decapitate immediately with sharp scissors. Place heads in a 100-mm dish containing 15 ml cold dissection buffer.

Isolate hippocampi

• 6

  Using curved forceps, hold one head firmly by inserting the tips of the forceps into the orbits. Using curved spring scissors, carefully cut through the skin and skull from the point of decapitation towards the orbits.

• 7

  Holding the head with one pair of Dumont forceps, use another pair of Dumont forceps to peel back the skin and skull, exposing the brain. Work caudally from beneath the olfactory bulbs.

• 8

  Using curved forceps, carefully remove the intact brain and place in a clean 35-mm dish containing dissection buffer.

• 9

  Under a dissecting microscope with fiber-optic lights, separate the hemispheres of the brain by moving Dumont forceps through the midline.

• 10

  One hemisphere at a time, remove the septum, thalamus, and hypothalamus from the cortex to expose the hippocampus, which will appear curved along the medial edge of the cortex.

• 11

  Remove the olfactory bulb and striatum to further expose the hippocampus for easy removal.

• 12

  Carefully remove the meninges and choroid plexus.

  The meninges form a thin layer covering the surface of the brain. The choroid plexus attaches to the meninges at the edge of the hippocampus. Both of these layers are highly vascularized, which facilitates locating and removing them.

• 13

  Using sharp Dumont forceps, cut or separate the hippocampus from the rest of the brain and place it in a 35-mm dish containing dissection buffer.

• 14

  Repeat steps 10–13 with the other hemisphere, then repeat the entire procedure with the remaining heads. Place all hippocampi in the same 35-mm dish containing 3 ml dissection buffer.

  It should take less than 1 hr from the time the animal is sacrificed to the time all hippocampi are completely dissected. After ~1 hr, cell viability decreases dramatically, and the brain tissue becomes more difficult to work with.

Dissociate cells

• 15

  In a laminar-flow hood, remove the dissection buffer from the dish and mince tissue into fine pieces using spring scissors.

• 16

  Using a 1000-μl pipet tip, aspirate the minced tissue and place into a 15-ml tube containing 5 ml papain solution. Incubate in a 37°C incubator for 45 min with gentle agitation every 10 min.

• 17

  Triturate tissue gently with a 5-ml pipet until pieces are broken up and solution appears cloudy.

• 18

  Centrifuge at 200 × g for 5 min. Gently remove supernatant by aspiration.

• 19

  Resuspend pellet in 5 ml of 1/1 solution. Using a 5-ml pipet, gently pipet up and down until the tissue is totally dissociated (15–20 times). Do this slowly to avoid bubble formation and facilitate maximum trituration.

• 20

  Allow suspension to settle for 10 min.

  After 10 min, some tissue chunks will have fallen to the bottom of the tube.

• 21

  Using a 1000-μl pipet tip, gently remove the supernatant and layer it on 10 ml of 10/10 solution, rotating the 10/10 solution tube as the supernatant is layered.

  The 10/10 solution column traps dead cells and debris. This ‘cushion’ should be ~8 to 10 ml in volume.

Plate and maintain cells

• 22

  Centrifuge suspension at 200 × g for 10 min. Discard the supernatant and resuspend the pellet in 2–6 ml plating medium 2. Determine cell density using a hemacytometer.

• 23

  Just before plating cells, remove water from polyornithine/fibronectin-coated cover-slips. Add 2 ml plating medium 2 and plate 4–8 × 10⁵ cells to each coverslip in a 35-mm dish.

• 24

  On the second day (after 24–36 hr), remove half of the plating medium and replace with fresh pre-equilibrated feeding medium 2.

• 25

  Twice a week, replace half of the medium with fresh pre-equilibrated feeding medium 2.

  Cells can be maintained in culture for up to 3 weeks. Always keep the neurons covered with medium, as they are extremely sensitive to air and will die if exposed even briefly.

1/1 solution

• 500 μl ovomucoid inhibitor (Worthington, 3182)

• 0.5 vial (0.25 mg) DNase (Worthington, LK003172)

• Add HBSS (no phenol red, Invitrogen, 14025-092) up to 5 ml

• Filter-sterilize using a 0.22-μm filter

• Prepare fresh

• Before use, dissolve 1 vial ovomucoid inhibitor in 32 ml HBSS.

10/10 solution

• 100 mg trypsin inhibitor (Sigma, T9253)

• 100 mg bovine serum albumin (Sigma, A9647)

• Add HBSS (no phenol red, Invitrogen, 14025-092) up to 10 ml

• Mix well and incubate at 37°C for 10 min to dissolve

• Filter-sterilize using a 0.22-μm filter

• Prepare fresh

Basic medium

• 500 ml modified Eagle medium (MEM, Invitrogen, 51200-038)

• 2.5 g glucose

• 100 mg NaHCO₃

• 50 mg transferrin (Calbiochem, 616420)

• Adjust pH to 7.4

• Filter-sterilize using a 0.22-μm filter

• Store at 4°C

Digestion buffer

• 137 mM NaCl (400 mg/50 ml)

• 5 mM KCl (18 mg/50 ml)

• 7 mM Na₂HPO₄ (50 mg/50 ml)

• 25 mM HEPES (150 mg/50 ml)

• Adjust pH to 7.2

• Filter-sterilize using a 0.22-μm filter

• Store at 4°C

Dissection buffer

• 500 ml HBSS (Invitrogen, 14170-112)

• 5 ml 1 M HEPES (Invitrogen, 15360-080)

• 5 ml 100× penicillin/streptomycin (Invitrogen, 15140-122)

• Store at 4°C

Feeding medium 1

• 97 ml basic medium (see recipe)

• 1 ml 200 mM glutamine (Sigma, G7513, stored at −20°C)

• 2 ml 50× B-27 supplement (Invitrogen, 17504-044)

• 3 μM cytosine arabinoside (Ara-C, Sigma, C6645)

• Filter-sterilize using a 0.22-μm filter

• Store up to 1 week at 4°C

Feeding medium 2

• 9.75 ml neurobasal medium (Invitrogen, 21103-049)

• 200 μl 50× B-27 supplement (Invitrogen, 17504-044)

• 50 μl GlutaMAX-1 (Invitrogen, 25030-061)

• Filter-sterilize using a 0.22-μm filter

• Prepare fresh

Hank’s balanced salt solution (HBSS)

• HBSS without calcium, magnesium, or sodium bicarbonate (Powder, Sigma, H2387)

• 350 mg/liter NaHCO₃ (4 mM)

• 1.3 g/liter HEPES (5 mM)

• Adjust pH to 7.3–7.4

• Filter-sterilize using a 0.22-μm filter

• Seal with Parafilm to help maintain consistent pH

• Store up to 2 months at 4°C

Medium L, pH 8.0

• 0.174 g K₂HPO₄ (1 mM final)

• 2 ml 0.5 M EDTA (0.1 mM final)

• 1 liter H₂O

• Store up to several months at 4°C

Papain solution

• 1 vial (100 units) papain (Worthington, 3176-PAP2)

• 0.5 vial (0.25 mg) DNase (Worthington, LK003172)

• Add HBSS (no phenol red, Invitrogen, 14025-092) up to 5 ml

• Filter-sterilize using a 0.22-μm filter

• Prepare fresh

Percoll gradients

• 23% Percoll:

• 12 ml 100% Percoll and 14 ml H₂O in 26 ml 2× sucrose buffer (see recipe)

• 15% Percoll:

• 30 ml 23% Percoll in 16 ml 1× sucrose buffer

• 10% Percoll:

• 20 ml 15% Percoll in 10 ml 1× sucrose buffer

• 3% Percoll:

• 6 ml 10% Percoll in 14 ml 1× sucrose buffer

• Adjust solutions to pH 7.4 and store up to 5 days at 4°C.

• To create a discontinuous gradient, carefully layer 2 ml of each Percoll solution in an 11-ml (16 × 100–mm) polycarbonate high-speed centrifuge tube, starting with 23% at the bottom. Prepare gradients on the day of or the day before the experiment, and keep at 4°C until use.

Plating medium 1

• 89 ml basic medium (see recipe)

• 10 ml fetal bovine serum (FBS, heat-inactivated at 57°C for 30 min, stored at 4°C)

• 1 ml 200 mM glutamine (Sigma, G7513, stored at −20°C)

• 200 μl 12.5 mg/ml insulin (stock prepared from powdered insulin [Sigma, I5500] in 10 mM HCl and stored at −20°C)

• Prepare fresh on day of use

• Filter-sterilize using a 0.22-μm filter

Plating medium 2

• 9.23 ml neurobasal medium (Invitrogen, 21103-049)

• 0.5 ml FBS (heat-inactivated 30 min at 55°C)

• 200 μl 50× B-27 supplement (Invitrogen, 17504-044)

• 25 μl 12.5 mg/ml insulin (stock prepared from powdered insulin [Sigma, I5500] in 10 mM HCl and stored at −20°C)

• 25 μl GlutaMAX-1 (Invitrogen, 25030-061)

• 20 μl G5 supplement (Invitrogen, 17503-012)

• Prepare fresh on day of use

• Filter-sterilize using a 0.22-μm filter, then pre-equilibrate in a 37°C, 5% CO₂ incubator

Protease inhibitors

• Aprotinin: Prepare 10 ml of an aqueous solution containing 20 mg aprotinin (2 mg/ml). Store in aliquots up to 1 year at 4°C. Where indicated, add immediately before use at a 1:1000 dilution (2 μg/ml final).

• Leupeptin: Prepare 10 ml of an aqueous solution containing 10 mg leupeptin (1 mg/ml). Store in aliquots up to 1 year at −20°C. Where indicated, add immediately before use at a 1:1000 dilution (1 μg/ml final).

• Phenylmethylsulfonyl fluoride (PMSF): Prepare 10 ml of 100% isopropanol containing 0.348 g PMSF (200 mM). Store in aliquots at −20°C (stable at least 9 months). Where indicated, add immediately before use at a 1:200 dilution (1 mM final).

Sodium phosphate buffer, pH 7.4, 0.1 M

• 11.4 g NaH₂PO₄·H₂O

• 85.79 g Na₂HPO₄·7H₂O

• 2 liters H₂O (final 0.2 M sodium phosphate)

• Store up to several months at 4°C

• Dilute 1:1 (final 0.1 M) immediately before use

Sucrose buffer, pH 7.4, 1×

• 2 × sucrose buffer:

• 21.9 g sucrose (0.64 M)

• 2 ml 0.1 M EDTA (2 mM)

• 240 mg HEPES (10 mM)

• 100 ml H₂O

• Filter-sterilize using a 0.22-μm filter

• Store up to 2 weeks at 4°C

• 1 × sucrose buffer:

• Dilute 2× solution 1:1 with H₂O

• Adjust pH to 7.4 with NaOH

• Store up to 5 days at 4°C

Sucrose gradients

• 7 ml 1.2 M sucrose

• 6 ml 1.0 M sucrose

• 6 ml 0.8 M sucrose

• 6 ml 0.6 M sucrose

• 6 ml 0.4 M sucrose

• Prepare sucrose solutions individually in Medium L (see recipe) and store up to 5 days at 4°C. On day of experiment, layer solutions on top of each other in a thin-walled, 38.5-ml, 25 × 89–mm ultracentrifuge tube, starting with 1.2 M at the bottom. Keep at 4°C until use.

Wash buffer

• 6.142 ml 2 M NaCl (122 mM final)

• 500 μl 1 M KCl (5 mM final)

• 460 μl 0.25 M NaH₂PO₄ (1.15 mM final)

• 4 ml 0.5 M PIPES (20 mM final)

• 0.1 g D(+)-glucose (1 mg/ml final)

• 100 ml H₂O

• Adjust to pH 6.8

• Prepare fresh on day of use and keep at 4°C

Background Information

The development of FM styryl dyes and the establishment of techniques permitting the staining and destaining of recycling synaptic vesicles at nerve terminals has advanced our knowledge of the synaptic vesicle cycle (Betz et al., 1992, 1996; Cousin and Robinson, 1999; also see UNIT 2.6). However, a number of challenges still remain in our elucidation of the molecular mechanisms underlying the synaptic vesicle cycle, particularly the modulation of these mechanisms by synaptic vesicle proteins. Thus, application of a multidisciplinary approach to visualizing and identifying the proteins on synaptic vesicles is critical to the field. Classical methods such as differential and density centrifugation and electron microscopy are still very useful. When used in combination with immunostaining techniques as described herein, they provide biochemical and morphological tools to determine the relative intracellular localization of particular proteins in intact synapses (Ilardi et al., 1999; Su et al., 2001; Tian et al., 2003; Leenders et al., 2008).

Electron microscopy of pre-embedding immunogold-labeled material allows direct, high-resolution localization of specific proteins in subcellular compartments, such as synaptic vesicle proteins in synapses. Although extra care should be taken with the silver enhancement to preserve this high resolution, this method is preferred over the im-munoperoxidase method, because it is easier to perform and more sensitive. However, it is not appropriate if ultrastructural studies require higher precision, as the diaminoben-zidine (DAB) reaction product can spread beyond the antigen-localization site (Cuello, 1993).

Critical Parameters and Troubleshooting

Anticipated Results

Typical results for each Basic Protocol are shown in Figures 2.7.3 to 2.7.5. In Figure 2.7.3, an immunoblot of protein markers for each subcellular fraction indicates the purity of the synaptic vesicle fraction, with both synaptophysin and VAMP2 as markers for the synaptic vesicle proteins. Both immunogold EM staining of intact synapses from synaptosomal preparations (Fig. 2.7.4) and immunofluorescent staining of cultured hippocampal neurons (Fig. 2.7.5) show specific staining of the synaptic vesicle membrane protein synaptophysin. The same monoclonal antibody against synaptophysin was used in all three experiments.

Time Considerations