Protocol

Preparation of Rodent Hippocampal Slice Cultures

Department of Biology, University of Iowa, Iowa City, IA 52242, USA.

¹Corresponding author (michael-e-dailey{at}uiowa.edu)

INTRODUCTION

Rodent organotypic hippocampal slice cultures (OHSCs) provide an outstanding preparation of central nervous system tissue for exploring the dynamic structural and physiological features of neuronal and glial cells within their native three-dimensional environments. It is a straightforward matter to obtain tissue slices from neonatal rodents. These slices culture well for periods up to several weeks and are easy to manipulate, allowing for a variety of in vitro experimental models. OHSCs provide good optical and physiological accessibility for studies involving live cell imaging, with high spatial and temporal resolution. This protocol is used to harvest tissues for both immunohistochemical labeling after fixation, and for confocal time-lapse imaging in live tissues labeled by a variety of fluorescent dyes or by biolistic or viral transfection.

RELATED INFORMATION

This protocol is adapted from methods presented in Stoppini et al. (1991). Brief practical guides to imaging cells in live brain slices (Kurpius and Dailey 2005) and maintaining slice cultures in the imaging setup (Dailey et al. 2005) are also available. See our laboratory’s website (http://www.biology.uiowa.edu/daileylab/) for some movies of experiments done using OHSC. Additional information about live cell imaging can also be found on-line (http://microscopyu.com/articles/livecellimaging/index.html).

MATERIALS

Reagents

Acetone (100%)

Dissection medium (Hank’s balanced salt solution [HBSS; Gibco] containing 6 mg/mL glucose)

Ethanol (70%)

Filter culture medium (FCM), prewarmed to 36ºC

Tissue source

This protocol is used to culture slices derived from post-natal day (P) 3 to P12 rat or mouse hippocampi. For long-term cultures (>1 d), slices fare best when they are derived from P5 and P6 animals. As the age of the donor animal increases beyond P7, long-term culture success decreases. This is likely because tissues become more dependent on aerobic-based metabolism as they mature beyond 1 wk of age. Tissues excised from adult animals generally do not fare well for longer than a few hours ex vivo.

Equipment

Beaker (50 mL)

Dishes (35 mm), tissue culture (Falcon or Corning)

Fiber Lite (Model 190, Dolan-Jenner Industries)

Filter paper discs, Whatman #1

Forceps, Adsen-Brown (Fine Science Tools, 11627-12)

Forceps, Dumostar, Dumont #3 (Fine Science Tools, 11293-00)

Forceps, Dumostar, Dumont #5 (Fine Science Tools, 11295-00)

Forceps, Graefe (Fine Science Tools, 11050-10)

Gloves, latex

Hood, tissue culture, laminar-flow

Ice and ice bucket (optional; see Step 5)

Incubator (5% CO_2, 100% humidity, 36ºC)

Inserts (1.0-µm pore size), for tissue culture (Falcon, 353102)

Kimwipes

Microscope, dissection (Leica, WILD M3C)

Paint brushes, size 1 and 2 (Dick Blick Wonder White, 2026)

Parafilm

Pipettes (3 mL), disposable, graduated, sterile

Plates (six well), tissue culture (Falcon, 353502)

Razor blades, double-edge (Persona)

Scalpel, #3

Scissors, iris (Fine Science Tools, 14060-09)

Scissors, spring, Vannas-style (Fine Science Tools, 15002-08)

Scissors, standard surgical (Fine Science Tools, 14000-14)

Spatula, thin-blade

Squirt bottle

Time tape

Tissue chopper, manual (Stoelting Co.)

METHOD

Preparation of Equipment

1. In a laminar flow hood or biosafety cabinet, fill each well of a six-well plate with 1.1 mL of warmed (36ºC) FCM.

2. Place the culture inserts into the wells.

3. To facilitate arrangement of the slices later (see Step 31), wet the top of the insert surface with a few drops of FCM.

4. Remove the excess FCM. Place the plate into a humidified CO₂ culture incubator until use.

5. For each animal, fill three 35-mm dissection dishes with at least 4 mL of dissection medium. Place on ice or in a refrigerator (4ºC).

6. Working under a fume hood, thoroughly and carefully wipe down a double-edged razor blade with acetone-soaked Kimwipes to remove any traces of oil:
Protect your hands with latex gloves during this procedure.

i. Fold two Kimwipe sheets together several times to make a thick pad. Saturate the Kimwipes with acetone from a squirt bottle.

ii. Hold the noncutting edges of the blade between the thumb and forefinger of one hand. Use short strokes of the folded Kimwipe pad to wipe the blade from center to edge.
Be very careful not to wipe in a direction along the length of the blade as this may lead to injury.

iii. Repeat this procedure, rotating and flipping the blade over, as necessary, to clean both edges and both sides of the blade.

iv. Put the cleaned blade into a 50-mL beaker filled with 70% ethanol.

7. Add the paint brushes to the ethanol, checking that the bristles are thoroughly saturated.
Repeated ethanol exposure can cause drying and cracking of the brush handles. To prolong the life of the brushes, wrap the handles with Parafilm (Fig. 1i , arrow).

View larger version (108K): [in this window] [in a new window]

Figure 1. Illustration of the collection of hippocampal tissues. With some practice, the following procedure takes as little as 15 min from decapitation of the animal to placing the cultures in the incubator. Brains are collected and prepared (a-c), and hippocampi are removed (d-f). Once the tissue chopper is ready (g), the hippocampi are chopped (h), transferred to fresh dissection medium (i), and slices are separated (j). Ideal slices (k) have a compact and clearly visible dentate gyrus granule cell body layer (D) and pyramidal neuron cell body layers (CA3 and CA1). Slices are then placed on tissue culture inserts (l) for short-term or long-term culture in a CO2 incubator.

8. Prepare the tissue chopper:

i. Place the tissue chopper and the beaker containing the razor blade and paint brushes in the sterile tissue culture hood.

ii. Remove the cleaned razor blade from ethanol. Allow the blade to air-dry.

iii. Spray down the tissue chopper with 70% ethanol. Allow the chopper to air-dry in the tissue culture hood.

iv. Place two strips of filter paper (one on top of another) onto the stage of the tissue chopper. Secure the edges of the filter paper to the stage with time tape.

v. Attach the dried blade to the tissue chopper. Check that the cutting edge of the blade is contacting the filter paper evenly when the arm is lowered.

vi. Moisten the filter paper with dissection medium, making sure that there are no wrinkles in it.
Be sure that no part of the blade is exposed along the top of the arm at any time during the procedure (Fig. 1g). If the blade is not tightly fastened to the holder, it can shift during the cutting, thus exposing the top of the blade. This can lead to injury. Make sure to remove the blade carefully after the procedure is complete.

Collection of Hippocampi

9. Swiftly decapitate the animal with surgical scissors. Place the head into a dish of cold dissection medium.

10. Using iris scissors, make a midline incision to remove the skin from the top of the skull (Fig. 1a).

11. Expose the brain by cutting the skull shallowly up the midline from the foramen magnum to the level of the eye sockets.
Keep the scissor-tips pointed up to avoid damaging the underlying brain tissue.

12. Make lateral incisions toward each eye (Fig. 1a, dashed lines).
The thicker skulls of older animals may also need to be cut at the base of the brain, above the cerebellum.

13. Reflect the skull flaps back using the Adsen-Brown forceps (Fig. 1b).

14. Gently slide the spatula under the brain. Flip the brain out of the skull into the second dish of cold dissection medium.

15. Using a scalpel, cut the brain in half using a midline incision. Orient the hemispheres medial-side up.
The next portion of the dissection is best performed on the stage of a dissection scope (100X-400X total magnification) equipped with both trans- and epi-illumination.

16. To hold each hemisphere in place during the next stage of dissection, pierce through the rostral end of the brain (Fig. 1c, red dots) with one of the Dumont forceps.

17. With the other hand, remove the brain stem and cerebellum (if still attached) with the Graefe forceps.

18. Gently remove the midbrain (Fig. 1c, dotted black circle).
The hippocampus is delineated by a large vessel running along its length (Fig. 1d, arrow). Extreme care must be taken to avoid damaging the hippocampus while removing the overlying midbrain tissues.

19. Cut the tissue at each end of the hippocampus with the spring scissors (Fig. 1e, arrows).

20. Flip the hippocampus over and cut it free of the hemisphere (Fig. 1f, arrow).

Preparation of Hippocampal Slices

21. Cut the end of a disposable plastic transfer pipette so that the hippocampi will fit inside.

22. Using the cut pipette, transfer the hippocampi individually to the stage of the tissue chopper.
More than one hippocampus can be cut at once. To provide good visibility, focus a fiber optic light onto the stage of the tissue chopper.

23. Orient the hippocampi so that the side with the major blood vessel is facing down.
Because there is a natural curve to the hippocampus, make sure that the central portion is oriented perpendicular to the blade. Don’t try to completely straighten them out; the tips don’t yield usable slices.

24. Cut transversely through the hippocampus at an interval of 300-400 µm (Fig. 1h).
See Troubleshooting.

25. Roll a wetted paint brush underneath each sliced hippocampus, transferring the tissue from the filter paper to the brush. Gently lift it off the chopper (Fig. 1i).
See Troubleshooting.

26. Place the slices into the third dish of cold dissection medium (Fig. 1j).

27. Using #3 or #5 forceps, gently separate the hippocampal slices (see Movie 1 ).

View as movie: (61K): [in QuickTime]

Movie 1. Post-natal day 5 rat hippocampal slices are separated with two pairs of Dumont #5 forceps. The hippocampus is oriented vessel-side down, while the slices are pulled apart at the subiculum. Any damaged tissue is cut from the rest of the slice using the spring scissors before the slices are positioned on inserts (Fig. 1l).

See Troubleshooting.

28. Isolate the best slices for culture and gather them into groups of four to six.
Select slices in which the CA3 and CA1 pyramidal neuronal cell body layers and dentate gyrus granule cell body layers are both compact and clearly visible (Fig. 1k). These slices have the least amount of neuronal cell damage and will maintain the best morphology over an extended culture period. Exclude those with any visible damage. A P6 rat typically yields an average of eight excellent slices per hippocampus, while a P6 mouse yields an average of six excellent slices.

29. Transfer the six-well tissue culture plate from the incubator to the tissue culture hood.
Turn off the tissue culture hood blower so as not to dry out the tissue slices.

30. Using a plastic transfer pipette with a cut end (I.D. ~3-4 mm), carefully transfer the slices with a minimal amount of dissection medium to a culture insert. Remove excess medium from the insert’s top surface.

31. Using a wetted paint brush, gently position the slices 2-3 mm apart near the center of the membrane insert (Fig. 1l).
Be careful not to stab the slices with the ends of the paint brush bristles.

32. After placing all tissue slices on membranes, immediately return the culture plate to the humidified CO₂ incubator.
After 24 h, the slices will begin to adhere to the membranes. The slices will flatten and spread out somewhat over a period of several days.

33. For ideal culture health, change the media every 2-3 d.
Removal of all of the original media in the culture wells is not possible. Thus, only 1 mL of FCM should be added at each subsequent feeding so that the slices do not get "flooded." Exchange media under a sterile tissue culture hood. As the cultures can dry out if the lid is off for an extended period of time, shut off the blower for the short time that media is actually being exchanged to minimize this risk. Care should be taken to use sterile technique and to keep the sash as low as possible to minimize contamination. See Troubleshooting.

TROUBLESHOOTING

Problem: Slices stick to the chopper blade.

[Step 24]

Solution: Assuming the razor blade has been properly cleaned with acetone, then it is likely that dissection medium dries on the blade in between animals, leaving the blade sticky. Wiping the blade with a paint brush dipped in clean HBSS right before use should stop the slices from sticking.

Problem: Sliced hippocampi do not come off the chopper intact.

[Step 25]

Solution: Be sure the filter paper is thoroughly wetted with dissection medium, and that the paint brushes are clean and wet. If you are having trouble transferring slices to the paint brush, place one brush in front of the hippocampus and the other behind it, rolling the second one underneath the hippocampus as you gently lift up.

Problem: Hippocampal slices do not come apart after being sliced.

[Step 27]

Solution: Make sure that the cutting edge of the blade is completely flush with the surface of the filter paper during chopping. An angled blade leaves the hippocampus only partially cut. Older animals have tougher meninges, so be sure that they are completely sliced through by checking that the paper has been penetrated by the blade (Fig. 1h).

Problem: Slices die in culture.

[Step 33]

Solution: Several factors can contribute to poor health or death of slice cultures. To minimize problems, make sure the FCM is warmed, gassed, and at proper pH (7.2-7.4) before feeding slices. We typically maintain the FCM at 4ºC for no more than 1 mo. Slice cultures can die if the stock solutions of the media components are used for longer than 1 mo. In preparation for feeding, FCM should be maintained in a warmed, gassed incubator for no more than 72 h. Moreover, slices can die if they dry out during preparation or feeding. Make certain that the tissue culture hood blower is off whenever the cover of the six-well plate is off and the cultures are exposed to air.

DISCUSSION

Healthy OHSCs typically have smooth edges, do not have a sandy or grainy appearance to their surface, and maintain their tight transparent neuronal cell body layers. If they are overfed and become flooded, they will become opaque. After 1 wk, cultures initially cut at 400 µm normally thin down to about 150-200-µm thick. Underfed cultures thin out much sooner and become nearly invisible.

As an in vitro tissue preparation, OHSCs provide distinct experimental advantages over other in vitro and in vivo preparations. Foremost, they present a central nervous system tissue environment that retains a more native complement of neurons and glia, with excellent accessibility for high-resolution optical imaging of cell structures or electrophysiological studies. OHSCs are amenable to various labeling techniques, including bath application, injection, or ballistic delivery of fluorescent structural or physiological indicators, as well as biolistic or viral transfection to express cDNAs or siRNAs in a subset of cells. Slices can be cultured for short (<1 d) or long durations (weeks to months). Over the course of the first week in culture, slices from neonatal tissues support studies of developmental events, including axonal and dendritic growth and synapse formation. More mature slice cultures (>10 d in vitro) can be used to study phenomena such as synaptic plasticity or cell death. Slice cultures can be prepared from transgenic, knockout, or GFP-reporter animals. Pharmacological agents can be applied easily for studies on the long-term effects of such compounds on development and pathology. Numerous slices can be harvested from a single animal and manipulated independently, so that control and experimental conditions can be studied in parallel.

There are, however, limitations to the procedure. Long-term OHSCs require donor tissue from a fairly narrow developmental age window (typically P4-P7). More mature tissues (>P7) generally are not suitable for long-term culture. In addition, the tissue isolation and slicing procedure induces death in some neurons. The procedure also induces glial cell activation and, within a week, the formation of an astrogliotic scar that encases the healthier tissues in the center of the slice. Also, neuronal injury and disruption of afferent and efferent connections may induce axonal and dendritic remodeling and reorganization of synapses among the remaining neurons.

ACKNOWLEDGMENTS

We thank former Dailey lab members, especially Drs. Glen Marrs and Raheel Ahmed, who helped develop, teach, and refine the methods.

REFERENCES

Dailey, M., Marrs, G.S., and Kurpius, D. 2005. Maintaining live cells and tissue slices in the imaging setup. In Imaging in neuroscience and development: A laboratory manual (eds. R. Yuste and A. Konnerth), pp. 1–8. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York.

Kurpius, D. and Dailey, M. 2005. A practical guide to imaging microglia in live brain slices and slice cultures. In Imaging in neuroscience and development: A laboratory manual (eds. R. Yuste and A. Konnerth), pp. 425–428. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York.

Stoppini, L., Buchs, P.-A., and Muller, D. 1991. A simple method for organotypic cultures of nervous tissue. J. Neurosci. Methods 37: 173–182.[Medline]