Preparation of Organotypic
Hippocampal Slice Cultures
Brain slice cultures are prepared by cutting thin sections
of brain tissue from neonatal animals and culturing
them as intact slices of tissue rather than as
dissociated cells. Like all culture methods, these preparations
offer the advantages of (1) long-term survival;
(2) precise control of the experimental conditions; (3)
excellent accessibility for viral vectors, biolistic transfection,
and other means of gene transfection; (4) survival
of tissue from neonatal-lethal transgenic animals;
and (5) excellent visibility of cells and subcellular
structures for morphological and electrophysiological
studies. Unlike cell cultures, however, these tissue
slices retain many features of their organotypic
organization, as has been described extensively (e.g.,
Zimmer and Gähwiler, 1984; Gähwiler, 1984; Gähwiler et al.
, 1997). This permits the identification of defined
cell groups, the stimulation or lesioning of specific
axonal pathways, and the formation of relatively normally
sized synaptic connections (Debanne et al.
Other advantages of the slice culture technique include
the ability to coculture slices from different brain
regions, thus facilitating the experimental manipulation
of long-distance connections in vitro
Gähwiler et al.
, 1987), and the ability to induce
conventional long-term potentiation (e.g., Debanne et al.
There are two variations of the technique that are in
use currently. In the roller tube technique, pioneered
by Gähwiler (1981), the slices are attached to glass coverslips
and placed in sealed test tubes on a roller drum
in a dry air incubator. In the membrane or interface
technique, pioneered by Stoppini et al.
(1991), the slices
are placed on semipermeable membranes and grown
statically in CO2
incubators. Primary differences
between the two techniques are that the roller tube
cultures generally become thinner than the membrane
cultures, but may be slightly more demanding and
time-consuming to prepare.
This article provides a concise description of the
steps involved in preparing and maintaining hippocampal
slice cultures. More details can be obtained
in Gähwiler et al.
(1998). Of course, many other brain
structures can be cultured readily using these techniques.
It is recommended that beginners start with
the hippocampus, as it is large, easy to dissect, has a
readily visible cell body layer, and has proven to be
robust when cultured with these methods.
Keep all solutions refrigerated until use. Maintain
Roller tube culture medium:
100ml basal medium Eagle (GIBCO, product
50 ml Hank's balanced salt solution (HBSS) with
Earle's salts (GIBCO, product # 24020)
or Hanks' balanced salt solution without
phenol red for fluorescence applications
(GIBCO product # 14025 or Cellgro,
product # MT21-023-CV)
50ml horse serum (GIBCO, heat inactivated previously
at 56°C for 30min in a water bath,
product # 16050)
4 ml 50% glucose solution, and
1 ml 200mM glutamine (from frozen aliquots)
- Membrane culture medium:
100ml MEM with Hank's salts and glutamine
(GIBCO product # 11575)
50ml HBSS as described earlier
50ml horse serum as described earlier
1 ml penicillin/streptomycin solution (Sigma
product # P-4333)
1 g HEPES
1 ml 50% glucose solution
- HBSS + glucose:
500 ml Hanks' balanced salt solution and
6 ml 50% glucose solution
- HBSS + glucose + kynurenate:
50ml HBSS + glucose and
0.5 ml 300 µM kynurenate stock solution
- Chicken plasma: There is variability in the amount
and type of anticoagulants contained in commercial
plasmas. We recommend lyophilized chicken plasma
from Cocalico Biologicals (product # 30-0390L). Reconstitute
to appropriate volume with tissue culture
water. Centrifuge for 18-20 min at 2500 rpm.
- Thrombin: Prepare aliquots at 150 units/ml. Store
frozen. Add 0.75 ml HBSS + glucose to 1 ml aliquot of
thrombin. This dilution can be adjusted to modify the
firmness of the plasma clot (see later).
- Antimitotics: 3mg each of cytosine-β-D-arabinofuranoside,
uridine, and 5-fluoro-2'-deoxyuridine in
100ml HBSS. Aliquot and freeze.
Purchase 12 × 24-mm coverslips of 0/1 thickness.
Place coverslips individually in the bottom of a large
glass dish. Fill the dish with enough 95% ethanol to
cover the coverslips. Soak them overnight. Replace the
95 % ethanol with 100% ethanol and soak overnight. Let
the coverslips dry overnight with the dish covered with
a paper towel. Transfer the coverslips to a petri dish,
wrap in aluminum foil, and bake at 200°C for 4-8 h.
B. Membrane Culture Dishes
Corning Costar Transwell polyester membrane
inserts and multiwell dishes (product # 3460) (also
available with various substrate coatings). Millipore
Millicell membrane inserts can also be used.
Small (ca. 3-cm blades) surgical scissors (1×)
Large (ca. 5-cm blade) surgical scissors (1×)
Scalpel or holder for razor blade shards (1×)
Small (ca. 3 × 20-mm) flat spatulas (6×)
Curved surgical forceps (1×)
Alcohol lamp (1×)
Aclar plastic (Ted Pella, Inc., product # 10501-25) (Cut
into 4 × 4-cm squares.)
Culture tubes (Nunclon Flat sided TC tubes, 110× 16mm)
Petri dishes (60 × 15 and 35 × 10mm)
E. Roller Drum and Drive Unit
Available from Bellco Glass (product # 7736-10164
and -20351). Set tilt angle to ca. 12° and rotation to ca.
F. Tissue Chopper
McIlwain Mechanical Tissue Chopper (Brinkmann
Instr., product # 023401002).
A. Prior to Dissection
Day before Culturing
B. Tissue Dissection
- Set out three 150-ml glass beakers with distilled
water, 70% ethanol, and 95% ethanol. Turn on hood
and light alcohol lamp.
- Sterilize instruments by dunking in 70% ethanol
and then in 95% ethanol. Large scissors and spatulas
should be flamed in the alcohol lamp after
removing from 95% ethanol.
- Fill culture tubes with 750µl of medium. Seal and
- Sterilize one aclar sheet for each animal by dunking
in 70% ethanol and then in 95% ethanol. Allow to
dry on sterile gauze pads in the hood.
- Break a double-edged razor blade in half, wipe with
95% ethanol, and insert into the tissue chopper. Swab the stage and mounted blade with 95%
- Set the micrometer on the chopper for the desired
slice thickness (start with 400µm). Set blade force
(start at the "9 o'clock" position).
- Thaw and prepare thrombin and chicken plasma.
- Fill one 60 × 15-mm petri dish with HBSS + glucose solution to cover the bottom of the dish. These
dishes are used for the dissected hippocampi. One
dish is needed for each animal to be dissected. Store
in a refrigerator.
- Fill one 35 × 10-mm petri dish with HBSS + glucose + kynurenate to cover the bottom of the dish. These
dishes are for the cut slices. One dish is needed for
each animal to be dissected. Store in a refrigerator.
- Mount an aclar sheet on the chopper stage.
- Place a 60mm × 15-mm petri dish containing
chilled HBSS + glucose in the hood.
- It is recommended that you start with rat or
mouse pups that are 5-7 days old. Younger animals
can also be used, but the dissection will be more challenging.
Animals older than 10 days rarely survive
more than a few days in vitro.
- Place one pup in a closed beaker with a small
piece of dry ice for anesthesia.
- When anesthetized, hold animal gently by head,
rinse neck area with 70% ethanol, and decapitate with
- Hold head right side up with the nose pointing
away. Insert tip of small scissors into the foramen
magnum toward the nose with the flat of the blades in
the horizontal plane. Cut by moving primarily the
blade on the outside of the skull. Repeat on the other
side. Discard bottom of head.
- Hold top of head nose down over the petri dish.
Using a spatula dipped in HBSS, push the brain stem,
cerebellum, and midbrain down gently, leaving the
cortex and hippocampus inside the dorsal skull. While
sliding the wet spatula along the sides of the cortex,
slide the cortex and hippocampus gently into the petri
- Under a dissecting microscope, position the
brain dorsal side down. Use one spatula in your left
hand to hold the brain in place by impaling the anterior
brain. Use the razor blade chip to free one hippocampus
at a time (Fig. 1). First, cut the lateral end of
the hippocampus and then cut posterior to the hippocampal
fissure, using the prominent blood vessel as
a guide. Repeat for the other hippocampus, making
one cut along the midline and another between septal
nuclei and the hippocampus.
- Using a spatula dipped in HBSS, gently lift and
roll the hippocampus away from the cortex in the
rostral to caudal direction so that it rests with area CA1
up. It is important that this be done without bending
- Transfer the hippocampus to the aclar sheet on
the chopper stage with CA1 up. The long axis of the
hippocampus should be perpendicular to the length of
the razor blade. Use a spatula to wipe any excess HBSS
away from the hippocampus on the aclar. Failure to
do so will result in the hippocampus being lifted by
the blade while chopping. Repeat for the other
- Chop slices.
- Remove the aclar sheet from the chopper and,
holding one corner, trim the aclar sheet into a "tongue"
shape around the sliced hippocampi. Insert the aclar
tongue into the small petri dish with HBSS + glucose + kynurenate and slide the slices off.
- Under a dissecting microscope, separate the
slices using two sterile spatulas. It is important that
this be done gently, without bending the slices.
- With proficiency, repeat steps 1-12 with a
- If desired, X-irradiate the dishes containing the
slices (1500 rad over ca. 1.5 min).
|FIGURE 1 Dissection of the hippocampus. The diagram shows a
view of the brain after peeling away the brain stem, midbrain, and
thalamus and removing it from the skull as described in the text. It
is pictured laying on its dorsal (i.e., cortical) surface and is viewed
from the ventral aspect. The hippocampus is dissected free by
making a series of sequential cuts with a razor blade shard as indicated
by the numbered dashed lines. After cutting, a spatula is
placed under the hippocampus and it is rolled caudally so that it
becomes free of the cortex and rests on its
C. Mounting Slices for Roller Drum Cultures
D. Mounting Slices in Interface Culture Wells
- Place one 20-µl drop of chicken plasma in the
center of each coverslip in one 60-mm dish.
- Under the dissecting scope, choose a healthy
slice and transfer to the plasma droplet with a spatula.
Healthy slices have a clear, well-defined continuous
cell body layer and no obvious signs of damage.
Repeat for all coverslips in dish.
- Spread the plasma around and around the entire
surface of the first coverslip around the slice (~3s).
Add a 20-µl drop of thrombin to the coverslip and mix
thoroughly with the plasma over the entire coverslip
(~3s). Position the slice in the center of the coverslip
and wipe excess plasma/thrombin off with a spatula.
Repeat with the other coverslips and then for the
second dish. After 5 min, the clot should have the consistency
of a fairly liquid gelatin and should retain an
indentation produced with a spatula.
- Lift coverslips from dishes and slide into tissue
culture tubes so that the bottom surface of the coverslip
rests on the flat surface of the tube. Cap the tube
tightly and gently tap the coverslip down to the
bottom of the tube, if necessary. If slices fall off coverslips,
increase the concentration of the thrombin to
produce a firmer clot.
- Place tubes in incubator.
The tissue dissection procedures are identical for
the roller tube and membrane culture methods. Procedures
for the latter preparations are different primarily
in the substrate upon which the slices are placed. For
membrane cultures, transfer slices with a wide-bore,
fire-polished glass Pasteur pipette into the wells. To
facilitate removal of slices from wells, small membrane
pieces (e.g., Whatman Nucleopore membranes
# 112107) can be cut and placed at the bottom of the
insert before plating the slices.
1. Add antimitotics after the first 4 or 5 days in culture.
2. Add 20µl to each tube.
3. Exchange culture medium after 24 h.
F. Feeding the Cultures
1. Feed the cultures once per week.
2. Pour off medium into a beaker in the hood. About
250 µl will remain in tube.
3. Replace with 500µl fresh medium.
G. Assessing the Health of the Cultures
- Cultures should remain adhered firmly to the
plasma and coverslip. If they fall off despite a firm
clot at the time of culturing, suspect either insufficiently
cleaned coverslips or a problem with the
- After 24-48 h in vitro, the slices will appear more
opaque as the macrophages and other cells proliferate;
remove damaged tissue. The cell body layer
should remain relatively clear and visible. Glial cells
migrating out of the edges of the slice should be
- After 14 days in vitro, most of the macrophages
should have disappeared and the cell body layer
should be continuous and more transparent than the
dendritic layers. Our studies indicate that the cultures
are not completely mature before this time.
- In our experience, unhealthy cultures can usually
be attributed to mishandling of the tissue during dissection
and mounting. If you have had good cultures
previously and an entire batch goes bad, suspect a bad
ingredient common to all of the cultures. If some cultures
are good and some are bad, suspect a problem of
dissection (it is helpful to note which slices came from
Debanne, D., Gähwiler, B. H., and Thompson, S. M. (1994). Asynchronous
pre- and postsynaptic activity induces associative longterm
depression in area CA1 of the rat hippocampus in vitro
. Proc. Natl. Acad. Sci. USA 91
Debanne, D., Guerineau, N. C., Gähwiler, B. H., and Thompson, S.
M. (1995). Physiology and pharmacology of unitary synaptic
connections between pairs of cells in areas CA3 and CA1 of rat
hippocampal slice cultures. J. Neurophysiol
Gähwiler, B. H. (1981). Organotypic monolayer cultures of nervous
tissue. J. Neurosci. Methods
Gähwiler, B. H. (1984). Development of the hippocampus in
vitro: Cell types, synapses, and receptors. Neuroscience 11
Gähwiler, B. H., Capogna, M., Debanne, D., McKinney, R. A., and
Thompson, S. M. (1997). Organotypic slice cultures: A technique
has come of age. Trends Neurosci
Gähwiler, B. H., Enz, A., and Hefti, E (1987). Nerve growth factor
promotes development of the rat septo-hippocampal cholinergic
projection in vitro
. Neurosci. Lett
Gähwiler, B. H., Thompson, S. M., McKinney, R. A., Debanne, D.,
and Robertson, R. T. (1998). Organotypic slice cultures of neural
tissue. In "Culturing Nerve Cells"
(G. Banker and K. Goslin, eds.),
2nd Ed., pp. 461-498. MIT Press, Cambridge, MA.
Stoppini, L., Buchs, P.-A., and Muller, D. (1991). A simple method
for organotypic cultures of nervous tissue. J. Neurosci. Methods 37
Zimmer, J., and G/ihwiler, B. H. (1984). Cellular and connective
organization of slice cultures of the rat hippocampus and fascia
dentate. J. Comp. Neurol