This protocol is a very detailed version of the microisland technique that was initially prepared for technicians with little culturing experience. This protocol can also be useful to people with more experience, using appropriate skimming. The protocol is a description of the methods used in the Segal lab, and it includes many lab-specific details such as using rats, dissecting out the hippocampus, using particular solutions and arresting cell division with Ara-C. These details will of course be modified for the needs of each lab.
This detailed protocol supplements the more scholarly but less detailed description that appeared in the chapter by Segal et al. in "Culturing Nerve Cells", Banker & Goslin, eds., MIT Press, 2nd edition, 1998.
Many of these solutions are described in the original microisland paper Segal and Furshpan (1990) in J. Neurophysiology or in a later paper Segal (1994) in J. Neurophysiology. Very detailed solution recipes are available on this site.
The procedure detailed here takes one full day, and some additional time on a subsequent day, depending on options chosen. Some steps such as making dishes and preparing some of the solutions are done before this day and a half of culturing.
(This is done in advance, and must be started at least three days before culturing.)
Preparing glass coverslips coated with agarose:
This step is done in the hood. Twelve cover slips (number 1 or 2 thickness) are placed in the bottom of each of several 150 mm plastic petri dishes. Typically 120 coverslips are prepared at once in ten 150 mm dishes. Care must be taken to reject any coverslips with dust on the surface, since these will result in a bad agarose layer.
A 0.2% (weight per volume) solution of agarose (Sigma agarose IIA) is prepared by mixing the agarose powder with water and autoclaving the bottle for at least 10 minutes (with the top loose to prevent explosion or implosion) or boiling the bottle in a beaker with some water to provide a uniform temperature to the bottle. A solution of agarose made previously is also acceptable as long as the agarose solution has not been returned to the bottle after some exposure to dust (such as on a cover-slip). After this, the solution is swirled to make sure it has mixed completely. The bottle is then placed on the top of an inverted 100 mm plastic petri dish to reduce the rate of heat loss to the surroundings, since the agarose solution will gel once it cools down towards room temperature.
Using a 1 ml pipette, agarose solution is placed on each of several coverslips. The drop of agarose should be big enough to cover the central part of the coverslip, so that when the coverslip is placed over the 8 mm well in the 35 mm dish it will cover the entire well, allowing for alignment errors. On the other hand, the drop should not be so large that it reaches the edge of the coverslip, since the hydrated agarose could provide a pathway for fluid to leave the dish.
The object is to leave a thin film of dried agarose. This is done by removing most of the liquid agarose, leaving only a thin film of agarose that will dry on the coverslip. The removal of the agarose is done using the same pipette used to apply agarose. Remove about a third of the agarose at a time, since trying to remove all the agarose in one step usually results in the solution beading up and failing to cover enough of the coverslip. The removed agarose is not re-used since it contains some gelled agarose and dirt particles, which make it difficult to obtain good agarose films. The used agarose is put in the bottom of the 100 ml plastic petri dish and later discarded. Unused agarose solution can be re-used another day, after boiling or autoclaving. The remaining agarose solution is stored in the refrigerator.
The coverslips are allowed to dry overnight in the hood. Faster drying in an oven seems to lead to cracking of the agarose.
Cutting holes in dishes:
This step is started in the hood. It is usually convenient to make about 100 dishes at a time. Small plastic petri dishes (35 mm) are put in large plastic petri dishes (150 mm), with 8 small dishes to a large dish. The small dishes are placed bottom up.
Before bringing the dishes out of the hood to have holes cut in them, the working surfaces outside the hood are cleaned with paper towels and then covered with new aluminum foil. The cutting is done in a semi-sterile way, with hands cleaned with de-ionized water and then 70% ethanol. A vacuum cleaner will be used to suction off stray bits of plastic produced in cutting the holes in dishes, so the suction end of the vacuum should be cleaned with alcohol prior to starting to make the dishes. It is important to avoid having plastic fragments in the dishes because they can remain in the cultures.
Outside the hood, holes (8 mm diameter) are cut in the bottoms of the small plastic petri dishes using a rotating cutting tool. Care is taken to not touch the inside of the dishes at any time. The holes are cut using a moderate speed of the cutting tool (about 60 on our custom-built machine), since a higher speed will result in melting rather than cutting. At least after every large dish, the cutting tool and surrounding area is vacuumed to pick up plastic debris from the grinding. Do not use the vacuum to clean up non-clean debris, since it will contaminate the end of the vacuum.
This is also done outside the hood, on the same aluminum foil surface. Usually the holes are cut in all the dishes before grinding the surfaces of all the dishes. The rough edges of the 8 mm holes are de-burred with a rotating large spherical drill bit (set about 100 on our machine), both on the inside and outside of the dish. The surfaces of each dishes are vacuumed to get off all debris. It is most important that there not be any plastic sticking up from the outer surface of the dish, as this could prevent the glass coverslip from being sealed on properly. Even if it looks as if there is no plastic rim sticking up after cutting the hole, it is important to de-bur this rim to allow for proper attachment of coverslips to the dish in a later step. The inner surface is less important. After every large dish, the grinding tool and surrounding area is vacuumed to pick up plastic debris.
Attaching coverslips with Sylgard:
Sylgard 184 (Dow Corning; (800) 248-2481) is mixed using a disposable cup and a 1 ml pipette, since it is difficult to clean anything that had Sylgard solutions on it. Typically, about 10 grams of the solution in the larger bottle is weighed out, followed by one tenth that weight of the solution in the smaller bottle. The two solutions for the Sylgard are mixed thoroughly with the pipette for several minutes. It is important to use the pipette to scrape the walls of the disposable cup to insure that there are no unstirred layers of the Sylgard solutions, since unmixed Sylgard will not solidify properly. Air bubbles will form in the Sylgard, but this is not a problem. After mixing, the Sylgard is then poured into the bottom of a 100 mm petri dish and a new 1 ml pipette is filled to 1 ml with the Sylgard mixture using an automatic pipetter. The disposable cup and first pipette are discarded. The 1 ml pipette is then removed from the pipetter and used to drip small drops of Sylgard onto the dishes.
Several small drops of Sylgard are placed on the outside of the bottom of the 35 mm petri dishes (about 6 is usually good, depending on the size of the drops). The drops should surround the 8 mm hole, but not be so close to the hole as to leak Sylgard into the well, or too close to the outside edge as to not be covered by the cover slip. There must be enough Sylgard to get the coverslip to stick, but not so much that much excess Sylgard leaks into the well after the coverslip is attached.
The coverslips are lifted with a fine forceps and placed agarose side down on the 35 mm petri dishes, centered on the 8 mm holes. This forms a seal that creates an 8 mm well in the petri dishes, with dried agarose covering the bottom of the well inside the dish. It is important to make sure that the coverslips are not resting on the outer ridge of the petri dish; this will prevent a good seal. After the coverslips are placed, it is important to check if they are sealed on properly, with no non-Sylgarded path for fluid to escape the well, and little or no Sylgard in the well. If there is not enough Sylgard, it is sometimes necessary to add another drop to the edge of the coverslip. If this doesn't correct the problem, the dish should be discarded.
The finished dishes are placed in a warming oven for 24 - 48 hours at 40 - 50 ºC. Above 50 ºC the plastic dishes will begin to melt. The temperature within drying ovens can be uneven, so the dishes should be placed away from the heat source. It is important to check that the Sylgard has cured fully, with no stickiness left. It also seems important to open the door of the drying oven several times during the period that the dishes are in the oven to allow the xylene in Sylgard to escape. It also seems to be important to leave the dishes out of the oven for a day before using them. They can be left in the hood or in a drawer.
Spraying collagen on culture dishes is done in the hood. For each culturing, typically 24 dishes are used, 6 small dishes to each big dish. Aluminum foil is placed in the hood to provide a protective surface on which the collagen can be sprayed. This foil will later be discarded, making it unnecessary to try to clean collagen off the bottom of the hood. Six 35 mm petri dishes are put off to the one edge of the foil. The bottoms of these dishes will be used to hold the tops of dishes being sprayed; the tops are discarded.
Collagen (Vitrogen 100, about 3 mg/ml, Collagen Corporation, Palo Alto CA (800) 227-8933) is used undiluted. About 2.5 ml is pipetted into the atomizer sprayer (Thomas Scientific #2753-L10) taking care not to break the small glass tube inside the sprayer. The sprayer is then placed on the platform made from petri dishes that keeps the sprayer horizontal and prevents its tip from touching the hood bottom.
Six 35 mm culture dishes are then placed near the center of the foil, in two rows of three, and their tops are placed on the six dish bottoms near the edge of the foil. Collagen is then sprayed using the sprayer, taking care not to spray collagen onto parts of the hood not covered with aluminum foil. The size and number of collagen dots will depend on the force of spraying, the horizontal distance to the dishes (~ 20 cm), the height (~10 cm) and the number of sprayings (15 -50). The size and number of dots can be monitored between sprays by placing a sample dish under a microscope. The dots should be about 100 - 200 µm diameter, and should be numerous, but not so numerous as to begin to overlap to form a mass culture.
As the sprayed collagen is drying, it is a good time to label the dishes. The dishes are labeled 1 - 24 on the plastic of the bottom of each dish and then on the top of each dish.
After spraying, the remaining collagen is discarded by pouring it into the garbage and spraying out the small amount of collagen that remains in the glass tube of the sprayer. The sprayer is then washed with about 2 ml of 0.3 M HCl, using a 2 ml pipette and putting about 0.7 ml at a time into the spraying chamber, taking care not to break the small glass tube inside the sprayer, and discarding and spraying the HCl as was done for the collagen. The sprayer is then washed twice with 70% ethanol from the wash bottle in a similar manner. The aluminum foil is discarded, and if culturing is to be done the same day, three of the dish bottoms are saved in the hood to be used during the culturing process.
The dishes sprayed with collagen can be used the same day or stored for at least several days at room temperature before use.
On the day of culturing, dishes are placed in a UV hood with the inside of the tops and bottoms of both the small and large dishes facing towards the UV lights. The dishes are handled in a sterile way using the dish tongs kept in the UV hood. The tongs are used to put the small dishes in place before UV exposure, but the large dishes can be placed by hand. The dishes and tongs are then exposed to UV for 60 - 120 minutes (using the timer), and then tongs are used to remove both the large dishes and small dishes, in such a way as to avoid touching the insides of the dishes with your hands. The dishes are then placed in the hood. As noted by Stephen Rayport, it appears that the UV exposure is critical for the dots of collagen to remain attached to the agarose, in addition to the anti-microbial effect.
The glass rings are kept in a covered Pyrex storage dish in 70% ethanol. Rinse the rings with distilled water three times, and drain as much water as possible so the rings will dry in the autoclave. Autoclave for about 20 minutes on the GRAVITY setting, and place the covered Pyrex dish in the hood. Also autoclave the magnetic stir bar.
Preparing enzyme and inhibitor solutions:
A water bath is heated to 37 ºC. Take out one 5 ml tube of kynurenate / magnesium solution (sodium kynurenate 10 mM, magnesium chloride 100 mM) from the freezer and warm it up in the water bath. It is important to make sure that all the material is in solution in the kynurenate / magnesium tube; vortexing is usually necessary.
Get out a 50 ml flask and label it "DMKyMg" (for dissection medium with kynurenate and magnesium). Forty five ml of dissection medium is poured into this flask. With a 10 ml pipette, the 5 ml of kynurenate / magnesium solution is added to dissection medium in the flask and the solution is mixed by passing fluid in and out of the pipette several times.
Using the same 10 ml pipette, 10 ml of DMKyMg is placed in a 13 ml tube which is labeled as "DMKyMg" and placed in the refrigerator. Using the same 10 ml pipette, and taking care not to touch the sides of the tube since this pipette will be re-used, 9.8 ml of DMKyMg are placed into one of the 13 ml tubes labeled as "E RAW", which contains cysteine (4.5 mg) that will be used to activate the enzyme. Using the same 10 ml pipette, 9.8 ml of DMKyMg is also placed in one of the 13 ml tubes labeled as "i raw", which contains the trypsin inhibitor (100 mg).
Using a 1 ml pipette, 3 drops of 0.3 M NaOH are dropped into the "E RAW" tube, being careful not to touch the sides since this pipette will be re-used. The cap is placed back on the tube and the solution is mixed by inverting it. If the solution looks yellow, add one drop at a time until it looks like pH 7.4 or pH 7.5 (it is OK for the solution to be a bit more alkaline than pH 7.4 since the papain latex that is added later is slightly acidic). It may be necessary to add half a drop by touching the inside of the tube, but this touching is not a problem as long as the tube containing cysteine is done before the tube containing trypsin inhibitor.
If you overshoot neutral pH, you can add 0.3 M HCl.
Next, add 4 drops of 0.3 M NaOH into the "i raw" tube. The cap is placed back on the tube and the solution is mixed by inverting it. If the solution looks yellow, add one drop at a time until it looks like pH 7.4. It may be necessary to add half a drop by touching the inside of the tube.
Add the correct volume of papain latex that contains 100 U of enzyme to the "E RAW" tube (papain is from Worthington Biochemical, Freehold NJ, (800) 445-9603 ). It is very important to vortex the papain latex suspension very well before taking some out, since it is a suspension, not a solution, and it settles out. The appropriate volume for 100 U is usually somewhere between 50 µl and 0.2 ml of the suspension; the volume needed is marked on the tube. The papain latex is kept refrigerated and the top of the tube is covered with parafilm to keep the small volume of enzyme suspension from drying out. Mix the "E RAW" tube by turning it over once or twice. The addition of the papain latex suspension makes the tube a bit cloudy; this will clear in 20 minutes unless the room is very cool. Leave the "E RAW" and "i raw" tubes out of the way in the hood, to be filtered later.
Preparing growth medium:
Prepare 140 ml of growth medium (170 if AraC is used instead of irradiation later) in a 250 ml flask according to the recipe for "5% - high prog". One of the ingredient is a solution of additives that contains glutamine, which breaks down at warm temperatures to glutamate. Since glutamate is toxic to neurons, it is important to keep solutions containing glutamine as cool as possible. Consequently, the additives should be brought to room temperature in a beaker containing unheated de-ionized water, but the frozen 9 ml serum tube and modified stable vitamin mix tube can be warmed in the 37 ºC water bath.
Filtering growth medium:
All of the growth medium is filtered using a 0.2 µm nylon filter stand assembly that is pre-washed with autoclaved water. Initially 100 ml of the 140 ml of growth medium is filtered (or 130 of the 170 ml), verifying on the scale of the flask that 40 ml remain in the flask. The 100 ml of growth medium is then poured into a new 250 ml flask labeled "5% - high prog". Take care not to contaminate the filter stand because it will be re-used.
The remaining 40 ml of non-filtered growth medium is then poured into the top of the filter stand without washing the filter a second time. The remaining 2 ml of serum from the 9 ml serum tube is put into the non-filtered 40 ml of growth medium using a 2 ml pipette, and the mixture is filtered and poured into a new 50 ml flask labeled "10% - high prog".
Putting growth medium in tubes and carbon dioxide exposure:
Using a new 10 ml pipette, 4.5 ml of the "5% - high prog" growth medium is put into one 13 ml tube, and 8.4 ml is placed into another 13 ml tube. Both tubes are labeled "5% - ".
Both flasks and both tubes of growth medium are then exposed to carbon dioxide gas (filtered) and put in the refrigerator to keep the glutamine from breaking down. The growth medium may become orange-yellow from this exposure, but this will balance out carbon dioxide loss later in the procedure.
Washing previous cultures:
If AraC had been used in a previous culturing on cultures to which more cells are to be added, the AraC is washed off before new cells are added. For each set of dishes, fluid is removed from the culture dish using vacuum and using the same 2 ml pipette, which is discarded. All vacuuming operations should be done from the outer part of the dish, never letting the central well dry out. Then 1 ml of "5% - high prog" is added to each dish, outside the well and the cultures are returned to the incubator. After this washing, add a 10 mm glass ring to each dish, centered over the well. WARNING: Any time dishes with growth medium are being moved, be very careful not to tilt the dishes. This can wet the top corners of the petri dish with growth medium, and lead to growth of fungus coming from the less sterile outside. Be very careful about this since it is probably one of the most common causes of fungal contamination.
Filtering enzyme and inhibitor solutions:
The enzyme solution should now be completely clear, so it should be filtered into a new 13 ml tube labeled ENZYME. The solutions are filtered using a 20 ml syringe and a small 0.2 µm filter that is pre-washed with 10 ml of autoclaved water.
The inhibitor solution should be filtered using another 20 ml syringe and another pre-washed small 0.2 µm filter, into a new 13 ml tube labeled INHIBITOR.
Both tubes are placed in the refrigerator.
Two 60 ml dishes are put in the dissecting area, and the remaining dissection medium with kynurenate and magnesium "DMKyMg" is poured into the two dishes. Two of the dish bottoms left over from the collagen spraying can be placed on either side of the dissecting microscope to hold the fine forceps and dissecting scissors, and one dish bottom is left in the hood to hold the heavy forceps.
If dialysis membranes are used: Use the heavy forceps to count out 24 rubber and glass rings and place them in a 1 liter beaker. If the rings are kept in Pyrex dishes with exactly 24 of each type of rings, it will not be necessary to count them using the forceps. Wash the rings with distilled water at least three times, until there is no longer any foaming. Be careful not to let the rings fall out into the sink when you are pouring out the water. Fill the beaker to 800 ml with distilled water, and place it on the burner pad above the Bunsen burner and light the burner. Be careful to never let the water boil away completely, since this would destroy the rings. Wash the Pyrex dish three times with distilled water after the rings are removed, and leave it to dry on the shelf with the top on.
The tips of the heavy forceps, fine forceps and dissecting scissors are then sprayed over the sink with 70% alcohol from the wash bottle. The instruments are then flamed by passing them briefly through the Bunsen burner flame, being sure to let the soon-to-be-flaming alcohol drip away from you, and they are placed on the 35 mm dishes.
The anesthetizing area is prepared by getting out the dissecting dish, a small plastic bag for animal remains, a 1 ml syringe with a 30 gauge needle filled with chloral hydrate (20% w/v), and a 50 ml beaker two-thirds full with 70% ethanol for dipping rats.
Get 5 newborn rat pups. Take out the first pup and inject it intraperitoneally with 0.2 ml chloral hydrate. Leave the needle in, since the chloral hydrate would leak out otherwise. Put the pup down on the counter since your movements would otherwise move the needle around and make the pup uncomfortable. When the pup no longer moves when touched, hold it by the tail and dip it into the 50 ml beaker with 70% ethanol and then pin it out on the dissecting dish, placing the pins at shallow angles so they do not get in your way during the dissection. Then inject the next rat with only 0.1 ml chloral hydrate, since the anesthetic will have a longer time to take effect before you dissect that pup.
Dissecting out the hemispheres:
Under the dissecting microscope, cut the scalp in the midline and then remove the scalp as far forward as the nose and as far laterally as the ears. Then make a small cut in the skull laterally on each side where the great veins are, and then cut the skull along the midline from the mid-cerebellum to anterior to the olfactory bulbs. Then cut laterally on each side and remove the delineated portion of the skull, fully exposing the cerebral hemispheres. Then cut deep in the midline to sever the corpus callosum, and cut the olfactory bulbs to separate their anterior and posterior halves. Then remove each cerebral hemisphere by cutting deeply with the scissors, paying special attention to making some cuts laterally enough to make sure that the temporal lobe comes free easily. Place each cerebral hemisphere in one of the 60 mm dishes with "DMKyMg" as soon as it is free.
When both hemispheres are removed, take the dissecting dish out of the hood and put the remains of the rat into the plastic bag. Then dip in ethanol the rat that was anesthetized, and anesthetize another with only 0.15 ml chloral hydrate. Repeat this until you have finished all the pups you are dissecting.
Dissecting out the hippocampuses:
Place the dish with the cerebral hemispheres under the microscope, lower the microscope to focus on the hemispheres, and increase the magnification. For each hemisphere, dissect out the hippocampus and place it into the other 60 mm dish with "DMKyMg". When removing the hippocampus, it is helpful to orient oneself by noting the position of the cut olfactory bulb. After each hippocampus is removed, place the rest of that hemisphere in the 35 mm dish used to hold the fine forceps.
When the dissection is complete, put the scissors and fine forceps on the lab bench to be washed later. Put the 35 mm dish on the counter to be discarded with the animal remains, and discard the 35 mm dish that supported the scissors.
Take the ENZYME tube out of the refrigerator and put it in the water bath.
Label a 13 ml tube "MAGNET", and put the magnetic stir bar into this tube by pouring the magnet in a sterile fashion from the tube in which it was autoclaved.
Put a 5 ml pipette into the pipetter, and take up ~4 ml of the "DMKyMg" solution from the dish that the hippocampuses are in, so as to make the inside of the pipette less sticky for brain tissue. Then expel this fluid into the dish and gently take up all the hippocampuses into the pipette, being careful here (and in all future steps) not to wet the cotton plug of the pipette. Expel the hippocampuses into the MAGNET tube, making sure that no hippocampus is sticking to the walls of the pipette. If any do stick, taking up and expelling more fluid with some bubbles will dislodge the tissue.
Take the ENZYME tube out of the water bath, discard its top, and rest a 2 ml pipette with the rubber bulb in the tube. Then remove the fluid from the MAGNET tube using the 5 ml pipette in the pipetter, being careful not to take up any hippocampuses. Add to the MAGNET tube about half of the 10 ml of ENZYME solution, and put on the top of the MAGNET tube and place the tube in the water bath. Turn on the magnetic stirrer, and make sure that the magnetic stir bar is rotating, slowly enough to avoid damaging the tissue, but quickly enough to be sure it keeps rotating. Set the timer for 20 minutes.
This waiting period is a good time to wash off the fine forceps and scissors. The scissors should be opened up wide by disconnecting the catch at the end, and the instruments should initially be cleaned with a wet kimwipe, with a motion going from the base to the tip. The instruments are then sprayed over the sink with 70% ethanol, then flamed. Several seconds after the flame goes out, they will be cool enough to put their tip covers back on and put them back where they are stored.
If dialysis membranes are used: The distilled water in which the rings are being boiled is then poured out, being careful that no rings fall out, and 600 ml of distilled water is put back in.
After the first 20 minute incubation is over, remove the fluid from the MAGNET tube, being careful not to suck up any hippocampuses, and add the rest of the ENZYME solution that has been sitting at room temperature in the hood. Place the closed MAGNET tube back in the water bath for another 20 minutes of stirring, again setting the timer and checking that the magnet is turning.
Discard the 5 ml pipette, move the 2 ml pipette used to dispense the ENZYME solution into the pipetter. This rotation of tubes for the pipetter insures that the tubes used for previous solutions get removed, so as not to expose the tube with the hippocampuses to solutions that should have been washed out already.
Discard the empty ENZYME tube.
Place the INHIBITOR and "DMKyMg" tubes from the refrigerator into the water bath to warm up.
Washing with DMKyMg:
After the second 20 minutes are over, remove the MAGNET tube and the "DMKyMg" tube from the water bath. Discard the top of the "DMKyMg" tube, and put a new 2 ml pipette on the rubber bulb into the tube.
Remove the fluid containing the enzyme from the MAGNET tube using the pipette in the pipetter, being careful not to wet the cotton plug in the pipette and not to suck up any hippocampuses. Add about 3 ml of DMKyMg to the MAGNET tube, in two pipette loads of about 1.5 ml, aiming the solution down the walls of the tube to wash away any remaining ENZYME solution. Remove the fluid with the pipette in the pipetter as before, and repeat this two more times until the DMKyMg tube is empty.
Then discard the pipette in the pipetter, put the pipette from the DMKyMg tube in the pipetter, discard the DMKyMg tube.
Washing with Inhibitor:
Remove the INHIBITOR tube from the water bath, put a 2 ml pipette in with the rubber bulb. Then remove the solution from the MAGNET tube and place about a third of the 10 ml INHIBITOR solution into the MAGNET tube, and place the MAGNET tube in the water bath and set the timer for 5 minutes.
During this and the next two five minute stirrings, there is time to put 10 mm glass rings on the wells of the new culture dishes. Remove the top from the Pyrex dish that has these autoclaved rings. Start with the last of the dishes of cultures (usually labeled 19-24 on the dishes). The heavy forceps should still be sterile, so remove the top from the 150 mm dish and remove the top from one 35 mm dish at a time and place a ring centered over the 8 mm well.
If dialysis membranes are used: You should replace the water in the beaker with the rings, adding 800 ml this time.
After the first two 5 minute incubations with INHIBITOR, remove the fluid from the MAGNET tube using the pipette in the pipetter, and add approximately 3 ml more of the solution from the INHIBITOR tube.
After the last third of the fluid from this INHIBITOR tube is added, discard the pipette on the pipetter, replace it with the pipette from the INHIBITOR tube, discard the inhibitor tube and put both "5% - " tubes from the refrigerator into the water bath.
After this third 5 minute stirring in INHIBITOR, take the MAGNET tube and the tube with only 4.5 ml of "5% - " from the water bath into the hood. Put a 2 ml pipette into the "5% - ", pipette out the fluid from the MAGNET tube using the automatic pipetter, and add 1.5 ml of "5% - " to the MAGNET tube. Repeat this washing twice, using the remaining part of the 4.5 ml of this "5% - " tube. Then rotate the pipettes as before, and discard the empty "5% - " tube.
Turn off the magnetic stirrer.
Take the tube with 8.4 ml of "5% - " from the water bath into the hood. Put a 2 ml pipette with a rubber bulb in the "5% - " tube. Remove the solution from the MAGNET tube with the automatic pipetter, and replace it with about a third of the 8.4 ml of "5% - ". Then use the pipette from the "5% - " tube to do the dissociation.
First, run some 10% carbon dioxide over the MAGNET tube and hold the pipette just inside the mouth of the tube near the gas flow, pressing on the bulb several times to fill it with carbon dioxide. This is to ensure that the gas bubbled into the MAGNET tube during the three triturations does not make the fluid alkaline. When this is done, leave the pipette in the fluid in the tube.
Expel about 0.5 ml of air from the pipette, and take up some of the fluid in the tube, allowing hippocampuses to come up into the pipette. Be careful not to expel too much air, since this will make the solution more alkaline and increase the risk of taking up too much liquid and letting the cotton plug in the pipette get wet. Press on the bulb to expel almost all of the fluid, allowing a bubble of air to come out only rarely so as not to create too much suction, and then take up more fluid. It is difficult to convey the amount of force that should be used, however, at the end of the triturations the fluid should be a little cloudy and about two thirds of the tissue should remain.
Remove the pipette and place it back in the "5% - " tube, and allow the sediment to settle in the MAGNET tube. Prepare a new 13 ml tube labeled "CELLS". After the sediment has fallen in the MAGNET tube, place a new 2 ml pipette in the pipetter and use it to remove as much cell suspension as possible from the MAGNET tube, without taking any of the sediment, and add the cell suspension to the CELLS tube.
Add half of the remaining "5% - " to the MAGNET tube, and triturate as before, with the goal of getting half of the remaining tissue to go into suspension. Again, put the suspension into the CELLS tube, and add the remaining "5% - " and triturate as before. Again, let any sediment settle (there should be very little now), and put the supernatant into the CELLS tube.
Put the MAGNET tube aside outside the hood. Later you will clean the magnet using de-ionized water and put the magnet back into its tube to be autoclaved.
Make sure the carbon dioxide gets turned off.
Plate cells onto the new cultures:
Note the time that plating is started, since you will begin washing cells off two hours later.
Put a new 1 ml pipette into the pipetter to use for plating cells into the wells of the dishes. Take up 1 ml at a time from the CELLS tube, mixing the cell suspension beforehand with the automatic pipetter. Put three drops into each well, touching the inside of the glass ring at the beginning to prevent the added fluid from beading up into a drop that doesn't touch the glass ring, which would result in an uneven distribution of cells plated, with most cells at the center of the drop. Putting the cells into the glass rings accurately without moving the rings is a delicate operation, and it is best to stabilize the automatic pipetter using a finger from your other hand, while holding the dish top face down with that same hand.
Do this until all 24 new cultures have cells plated onto the dishes.
Add cells onto the old cultures:
For each dish of cultures from the previous culturing, add cells. WARNING: Any time dishes with growth medium are being moved, be very careful not to tilt the dishes. This can wet the top corners of the petri dish with growth medium, and lead to growth of fungus coming from the less sterile outside. Be very careful about this since it is probably one of the most common causes of fungal contamination.
If not already done so (in the case of AraC treated cultures), add 10 mm glass rings to each of the 6 dishes in the 150 mm dish. Then take up 1 ml of the CELLS solution at a time, mixing the cell suspension as before. Drop 1-3 drops of the suspension into each well (with the number chosen depending on experience with cell survival and desired number of neurons per microisland), trying not to touch the glass ring or the culture. Drops are used instead of a measured volume to protect against the possibility of transferring contamination from one dish to another by immersing the pipette.
Do this for all 24 of the old cultures.
Wait two hours from the beginning of plating before starting to wash off the cultures to remove cells and debris that didn't adhere.
In the meantime, if you are using dialysis membranes you can add assemble the membranes (see that section).
Warm up the medium:
Take the flasks with "5% - high prog" and "10% - high prog" out of the refrigerator and warm them up in the water bath. Leave them in only long enough to warm a bit above room temperature, since growth medium has glutamine. If you are using dialysis membranes, this warming was started as you finished boiling the membranes. When you are finished warming the medium, turn off the heater in the water bath.
Wash each 150 mm dish of new cultures:
Take out four 2 ml pipettes, one for each 150 mm dish, and put a 10 ml pipette in the pipetter. Turn on the vacuum line. For each 150 mm dish, remove the rings from each of the six dishes using the heavy forceps, and put the rings back in their Pyrex dish. Don't drop the rings from high up, since this causes them to break. Fill the 10 ml pipette with 6 ml of the 5% solution, and start dripping 1 ml of 5% solution directly onto the well of each 35 mm dish, at approximately two drops per second. This step is one of the most important parts of the culturing, since it determines how many cells are left after the culturing. When the dish is positioned to catch the light correctly, you can see the debris being removed by the falling drops. Keep dropping fluid, if necessary from higher up, until this macroscopic debris is gone, since it will produce bridging between microcultures. Once the debris is gone, expel the remaining fluid into the dish away from the well. Following this procedure will ensure that there is little debris between microcultures, without washing so much that cells on microcultures are washed away as well.
Move the next dish to wash into view by turning the large 150 mm dish, to ensure that the dish is in the correct spot where you can see the debris best. This spot can be marked with black tape, which increases contrast. When the debris is gone from all six 35 mm dishes, vacuum the solution from all six dishes. All vacuuming operations should be done from the outer part of the dish, never letting the central well dry out. Then fill the same 10 ml pipette with 9 ml of the 10% solution, and add 1.5 ml to each of the six dishes, usually adding the fluid away from the wells, and only adding directly onto the wells if debris remains.
When all new cultures are fully washed, transfer the remaining few ml of the 10% solution into the 5% flask.
Wash each 150 mm dish of old cultures:
For each 150 mm dish, remove the rings as before. Fill the 10 ml pipette with 6 ml of the 5% solution, and drip 1 ml of the 5% solution onto the well of each 35 mm dish as before. Vacuum as before, but wash each dish with 1.5 ml of the 5% solution.
If irradiation is being used to stop cell division, the old cultures are left in the incubator until the next day when they are irradiated (see Irradiating), and then membranes are placed on the cultures (see "Assembling the Membranes" and "Placing Membranes on the Cultures").
The new cultures should be left undisturbed until 4 days later when they are irradiated or treated with AraC, and then left in the incubator until the next plating when these will be the old cultures and will have glial microcultures. If AraC is used, it is also applied to the new cultures on day 4, at which time the medium is changed to one with 10 mM magnesium and 1 mM kynurenate.
Some people assemble dialysis membranes attached to glass rings with rubber gaskets as a way of protecting cultures during feeding. Assembly of these membrane / ring combinations is done in a sterile fashion on aluminum foil on the counter.
Preparing the area:
Clean off the counter with damp paper towels, then put down some clean aluminum foil. Take the heavy forceps from the hood and spray 70% alcohol on them with the wash bottle at the sink to clean them off, then flame the forceps and put them down on the aluminum foil. Turn off the Bunsen burner.
Fill the two 250 ml flasks in the hood with distilled water, and put one back in the hood and put the other on the aluminum foil.
Put one new 35 ml dish and one new 60 mm dish on the foil. Discard the bottom of the 35 mm dish. Put the bottom of the 60 mm dish upside-down and take out the Teflon pusher from its plastic container and put it on the 60 mm dish bottom. (Try to keep the same orientation for the Teflon pusher, since one end touches the membranes and the other end touches your hands).
Get the package of membranes out from the refrigerator, and a new package as well if that will be needed. (Membranes are Spectra/Por molecular porous dialysis membranes, # 132498, 50 membranes per package, 33 mm diameter, MW cutoff 12-14,000; currently ordered from Fisher even though they are not listed in their catalog).
Fill the Pyrex dish that held the rings half way with distilled water, and put it onto the foil. Pour some distilled water into the top of the 60 mm dish. Pour most of the water out of the beaker with the rings, leaving enough to keep some of the rings wet.
Assembling each membrane:
Take one membrane from its package with the forceps, and place it in the water. Take a rubber gasket with the forceps and place it on the floating membrane. When the membrane becomes wet, put the membrane on top of the rubber gasket in the water, centered on the ring. Then take one glass ring with the forceps, and put the ring on the membrane, again centered. Take the top of the 35 mm dish and press it down on the rings and membrane, to force the glass ring down into the rubber ring, forming a tight seal around the membrane. It may take some tilting of the plastic dish to get the glass ring to go down into the rubber ring. Then lift the ring assembly, using the forceps to hold onto the rubber ring, and put it, with the membrane side up, on top of the 35 mm dish top. Then use the Teflon pusher to push the rubber ring 1-2 mm down, securing the rubber ring and the membrane on the glass ring. Do this carefully to avoid pushing too far down. Take the ring assembly with the forceps and place it in the Pyrex dish, open side up.
Boiling the Ring Assemblies:
When all rings are assembled, the top of the Pyrex dish is placed open side down in the hood and the bottom of the Pyrex dish is placed on the Bunsen burner. (If the top is left on during boiling it can be knocked off and broken by the boiling, and fluid also bubbles back and forth touching the less sterile outer surface of the petri dish. The dish is boiled for about 15 minutes, taking care not to let the water boil off completely, since this would destroy the rings and membranes.
Clean up the area where the membranes were assembled. Return the 250 ml flask to the hood, and place the heavy forceps back into the hood after alcohol flaming them.
When the membranes have finished the 15 minutes of boiling, the Pyrex dish is transferred to the hood using the insulated gloves. The vacuum is turned on, and the cotton plug is sucked out of a 5 ml pipette. The pipette is then attached to the vacuum, and the water is vacuumed out of the Pyrex dish. Water is added from one of the 250 ml flasks, and vacuumed again using the same 5 ml pipette. Then water is added from the other 250 ml flask, and the top is put on the Pyrex dish and secured with autoclave tape.
If washing off of cultures is to be done in a few minutes, the growth medium solutions are placed in the water bath now.
The Pyrex dish with the membranes is then autoclaved, set for about 30 minutes on the liquid setting. Don't interrupt the cell washing to remove the membranes from the autoclave. They do not need to be taken out immediately, but if they stay for many hours, all the water will boil off and the rings will begin to stick to each other (set a timer). Put the dish with membranes into the hood when they are done.
It is often helpful to look at the dishes soon after culturing. It is good to look at a few of the new dishes and a few of the added dishes after the first day of culturing. Another good time to look is just before irradiating, since any contamination introduced will be sterilized. Look for reasonable numbers of glial cells in the new dishes and reasonable numbers of neurons in the added cultures. In all cultures, if there is too much bridging between microcultures it is possible that too little washing was done on the first (or second) plating.
This ends the first day of the culturing. Shutdown checklist:
If membranes are used, the following procedures are done:
Prepare and arrange materials:
Make up 100 ml of "5% 1 KyMg low prog" growth medium in a 250 ml flask (see recipes). Give this some carbon dioxide.
Turn on the vacuum in the hood. In the hood there should be the Pyrex dish with the autoclaved membranes. Take the autoclave tape off this dish. Take out four 150 mm dishes and four 100 mm dishes. Put one 100 mm dish inside each 150 mm dish. Take out a 10 ml pipette and four 2 ml pipettes. Have the flame-sterilized heavy forceps on a 35 mm dish in the hood.
Incubate membranes with growth medium:
For each 100 mm dish, put 5 ml of growth medium in the dish. With the large forceps, lift one membrane assembly by its glass ring, pour out as much water as possible, and place it in the 100 mm dish, membrane side down. Be careful not to try to lift a ring assembly that is upside-down, since you might puncture the membrane. Transfer membrane assemblies until there are six in the 100 mm dish. Vacuum off the small amount of water in each membrane assembly using a 2 ml pipette, using the same pipette for the group of six membrane assemblies. Then take up 9 ml of growth medium and add 1.5 ml to each membrane assembly. Place the 150 mm dish with its contents into the incubator. When subsequent 150 mm dishes are added, keep track of the order in which they were added, because they will be removed in the same order.
When all four 150 mm dishes are in the incubator, warm up the growth medium in the 250 ml flask for about 5 minutes and take out eight 2 ml pipettes.
Transfer membranes to the dishes:
Remove the first 150 mm dish of membranes from the incubator, together with one of the 150 mm dishes of old cultures that were irradiated earlier that day. For each 150 mm dish, vacuum the medium off the old cultures with a new 2 ml pipette. (All vacuuming operations should be done from the outer part of the dish, never letting the well dry out completely). Then add 1 ml of growth medium to each culture, and put the membrane assemblies into the dishes, being careful not to get bubbles under the membranes. Then vacuum the medium that is outside the ring assemblies with another 2 ml pipette. If fluid has leaked out of the membranes, vacuum less fluid to compensate. This is very important because it makes sure that all cultures start out with the same amount of medium.
Copyright © 2002 Dr. Michael M. Segal, Department of Neurosurgery, Brigham & Women's Hospital, Boston MA 02115, USA. Individual copies may be made of page for use in educational institutions. Please send suggestions, comments and questions about the protocol by e-mail to microislands at-sign segal.org