Biolistic Transfection of Neurons

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Science's STKE  26 Sep 2000:
Vol. 2000, Issue 51, pp. pl1
DOI: 10.1126/stke.2000.51.pl1


One method used to study gene function is through the manipulation of gene expression by transfecting cells with DNA constructs designed to overexpress or knock out particular proteins. Unfortunately, transfection of cells and tissues remains a rate-limiting step for molecular studies in many fields, especially neurobiology. Conventional transfection techniques are of limited effectiveness, particularly in intact tissue. This protocol describes an alternative method for transfecting cells, called biolistics. Biolistics is a physical method of transfection in which target tissue is bombarded with DNA-coated gold particles using a "gene gun," produced by Bio-Rad Laboratories. Cells penetrated by gold particles have a high likelihood of becoming transfected. Because biolistic transfection relies only on the physical penetration of a cell's membrane, it is possible to use biolistics to transfect cells that are resistant to transfection by other methods, such as neurons in primary culture and organotypic slice cultures. This protocol provides information on optimizing the biolistic parameters for transfecting neurons in both of these preparations. Once optimized, biolistic transfection is a reliable and efficient method for studying gene function in many cell types, especially postmitotic neurons.


A powerful approach to studying gene function is to manipulate genes of interest by transfecting cells with DNA constructs designed to overexpress or knock out particular proteins. Unfortunately, the introduction of genetic material into cells and tissues remains a rate-limiting step for molecular investigations in several fields, especially neurobiology. Many different methods have been developed to transfect cells: microinjection, electroporation, calcium-phosphate transfection, lipofection, and viral transfection. For many cell types, gene transfer using these methods has become a routine tool for studying gene regulation and function. However, many other cell types, particularly postmitotic neurons, are not easily transfected by these methods.

In the late 1980s, John Sanford, Stephen Johnston, and colleagues developed a device that could potentially allow transfection of such resistant cells (1- 3). These inventors called their method "biolistics"--short for biological ballistics--which describes the process of shooting biological materials into living tissue. Biolistic transfection is accomplished by accelerating DNA-coated gold particles directly into cells, through the cell membrane and into the nucleus. This method was originally designed to circumvent difficulties in transfecting plant cells whose cell walls present a physical barrier to conventional transfection techniques (1). In the 10 years since the method was first reported, biolistics has also proven effective in transfecting a wide variety of animal tissues, as well as both eukaryotic and prokaryotic microbes, mitochondria, and microbial and plant chloroplasts (2- 6).

Biolistics is a physical method of transfection in which micron-sized gold particles are coated with plasmid DNA and then accelerated at high velocity toward target cells or tissue. Cells penetrated by a gold particle have a high likelihood of becoming transfected. For the biolistic device discussed in this protocol (the Helios Gene Gun system; Bio-Rad), high-pressure helium provides the acceleration for the gold particles. Older devices used gunpowder blasts (1) or high-voltage discharge (7) as pressure sources. Because biolistic transfection relies only on the physical penetration of a cell's membrane for successful transfection, it is possible to use biolistics to transfect cells that are resistant to transfection by other means, such as mammalian neurons in primary culture and organotypic slice cultures.

To study the signals that regulate neural development and function, we use both dissociated primary neuronal cultures and organotypic brain slice cultures (8, 9). Conventional techniques for transfecting neurons in these preparations have proven problematic because they are often unsuccessful, unreliable, or too time-consuming. In contrast, biolistic transfection is efficient, reliable, and does not require advanced molecular biological facilities for its application. Here we provide information on optimizing the biolistic parameters for transfecting neurons in both of these preparations. Once optimized, biolistic transfection is a reliable and efficient method for studying gene function in many cell types, including postmitotic neurons.

The procedures for biolistic transfection of dissociated neurons and organotypic slices are divided into four sections. The protocols for preparing dissociated neuron and slice cultures are presented first, followed by the protocol for biolistic transfection, and finally, a protocol for immunostaining slice preparations.


Dissociated Neuron and Astrocyte Cultures

Plastic zip-lock bags


Beaker (1 liter) and Teflon beaker cover

Borax (Na2B4O7-10H2O; Sigma no. B3545;

Boric acid


Cell strainers (70-μm filter; Falcon no. 2350;

Sterile centrifuge tubes, 10 and 50 ml

Clinical centrifuge

Collagen, rat tail (300 to 500 μg/ml; Collaborative Biomedical Products no. 4023B;

Coverslips, 12 mm (Fisher no. 12-545-80;

Culture plates, 24-well (Costar no. 3524;

Culture flasks (50 ml, NUNC no. 136196; 250 ml, Fisher no. 10-126-5;

Earl's balanced salt solution (EBSS; Gibco no. 14015-028;

95% ethanol (EtOH)

Fetal calf serum (FCS; Hyclone no. A-1115-N;

Sterile filter units (Fisher no. 09-740-39A, 0.2-?m filter size;

5-Fluoro-2(-deoxyuridine (Sigma no. F0503;


Hemocytometer (VWR no. 15170-263;

Hepes (1 M solution; Gibco no. 15630-080;

Horse serum (Hyclone no. SH30074.02;

Hydrochloric acid (HCl)



Kynurenate (KYN; Research Biochemicals International no. K-100;

Laminar flow hood with ultraviolet (UV) light

Medium essential medium (MEM; Gibco no. 51200-020;

MEM with 25 mM Hepes (Gibco no. 12360-020;

Sterile 1.5-ml microcentrifuge tubes

Dissection microscope

Inverted tissue culture microscope

N2 supplement (Gibco no. 17502-014;

Nitric acid

Papain (Worthington Biochemicals no. LS03126;

Pasteur pipets (autoclaved)

Penicillin/streptomycin (Pen/Strep; Gibco no. 15070-014;

pH meter

Phenol Red (Gibco no. 15100-019;

Pipetman and tips

Poly-D-lysine (PLYS; Collaborative Biomedical Products no. 40210;

High molecular weight poly-D-lysine (HMW PLYS; Sigma no. P-2636;

Razor blades


Soldering iron

Stripettes (sterile pipets; Falcon;


Surgical instruments:

Fine scissors (Fine Science Tools no. 14063-09;

Two pairs of fine forceps (Fine Science Tools no. 11251-35;

Blunt forceps (Fine Science Tools no. 11006-12;

Hippocampal-dissecting tool (Fine Science Tools no. 10099-15;

Ultrafine dissecting scissors (Fine Science Tools no. 15200-00;

Trypsin-EDTA (Gibco no. 25300-047;

Uridine (Sigma no. U6381;

Water bath at 37°C

Slice Cultures

Agarose (Gibco no. 15510-019;

Basal Medium Eagle (BME; Gibco no. 21015-037;

Hank's balanced salt solution (HBSS; Gibco no. 24020-117;

Horse serum (Hyclone no. SH30074.02, heat-inactivated;

Hepes (1 M solution; Gibco no. 15630-080;

Penicillin-streptomycin (Gibco no. 15070-063;

Dissecting microscope (wash with 95% EtOH to sterilize)

Squirt bottles filled with 70 and 95% EtOH

One glass petri dish (painted black to ease visualization of opaque brain slices and autoclaved to sterilize)

Beaker, 250 ml (autoclave to sterilize)


Reagents for Hepes-buffered artificial cerebrospinal fluid (ACSF):
Hepes (Gibco no. 11344-033;






Laminar flow hood with UV light

Paint brushes, size 000, rinsed in EtOH

Pasteur pipets (glass, autoclaved); cut thin tips off and place pipet bulb on cut side so slices can be sucked into large-bore end

pH meter

Slicer (10) (SD Instruments; phone no. 541-479-8697); sterilize winders by rinsing with 95% EtOH and exposing to UV for 10 min each side

Sterile tissue culture inserts, 0.4-μm pore size (Millicell CM Inserts no. PICM 03050 or Falcon no. 3090;

Sterile tissue culture plates (Falcon no. 1146;

Sterile disposable tissue culture filter units (0.2-μm filter size; Nalgene no. 156-4020;

Note: Nalgene filters are optimal as they have minimal detergent in their filter; in my hands, Corning filters invariably cause unhealthy slices, probably due to a higher detergent level in their filters

Surgical instruments: (autoclave to sterilize):

Razor blades

Fine scissors

Blunt forceps

Fine forceps

Water bath at 37°C

Biolistic Transfection

9-V batteries

Bio-Rad Helios Gene Gun system (Bio-Rad no. 165-2431; (includes gene gun, tubing prep station, optimization kit, and high-pressure helium regulator)


15-ml centrifuge tubes

Plasmid DNA (25 μg per preparation)

100% EtOH (fresh bottle)


Gold particles, 1.6 μm in diameter (Bio-Rad;

Compressed helium [grade 4.5 (99.995%) or higher], placed beside the laminar flow hood

Laminar flow hood

1.5-ml microcentrifuge tubes

Minicentrifuge (VWR no. 20668-212;

Compressed nitrogen [grade 4.8 (99.998%) or higher] with a regulator that registers a maximum of 30 psi; placed near and connected to the tubing prep station

Nylon mesh (Small Parts no. CMN-90-D;

Pipetman and pipet tips

Polyvinylpyrollidone (PVP; Bio-Rad;

Qiaprep spin miniprep kit (Qiagen no. 27104;

Quantum plasmid midiprep kit (Bio-Rad no. 732-6120;

Razor blades


Spermidine (Sigma no. S-0266;

Syringes (10 cc)


Gold-coat tubing (Bio-Rad;

Ultrasonic cleaner (Branson 1210;


Visualizing Transfected Cells by Immunostaining


Bovine serum albumin (Sigma no. A9418;


Chromium potassium sulfate

Dry ice

95% EtOH

100% EtOH

300-bloom gelatin (Sigma no. G2500;


Combination hot plate and stir plate

Krystalon (Fisher no. 23750008;




Normal goat serum (Gibco no. 16210-072;


Phosphate buffer (PB) (0.1 M)

Phosphate-buffered saline (PBS)


Slide racks

Sodium phosphate monobasic (NaHPO4 · H2O)

Sodium phosphate dibasic (NaH2PO4 · H2O)

Subbed slides


Triton X-100 (Sigma no. T9284;



Note: All solutions should be made in a sterile hood, unless indicated otherwise.
Recipe 1: PLYS
Prepare a 1 mg/ml solution in sterile distilled H2O.
Recipe 2: Collagen stock
Prepare a 300 to 500 μg/ml solution of rat tail collagen in sterile distilled H2O.
Recipe 3: Collagen/PLYS mixture
PLYS stock (Recipe 1) 160 μl
Collagen stock (Recipe 2) 800 μl
Sterile distilled H2O 8 ml
Mix and filter with a sterile filter unit.
Recipe 4: D-Glucose solution
Prepare a 1 M solution in MEM and filter with a sterile filter unit.
Recipe 5: Astrocyte medium
MEM 175 ml
FCS 20 ml
D-Glucose solution (Recipe 4) 4 ml
Pen/Strep 1 ml
N2 supplement 1 ml
Phenol Red 200 μl
Mix medium in a 250-ml sterile culture flask and filter with a sterile filter unit. Store medium in a 5% CO2/37°C incubator loosely capped.
Recipe 6: Neuronal medium
MEM with 25 mM Hepes 175 ml
Horse serum 20 ml
D-Glucose solution (Recipe 4) 4 ml
N2 supplement 2 ml
Pen/Strep 1 ml
Mix medium in a 250-ml sterile culture flask and filter with a sterile filter unit. Store medium in a 5% CO2/37°C incubator loosely capped.
Recipe 7: KYN solution
Dilute to 100 mM in DMSO.
Recipe 8: Dissection solution
Hepes (1 M solution) 5 ml (10 mM final concentration)
EBSS (sterile bottle) 500 ml
For preparation of cortical neurons, add:
KYN (100 mM, Recipe 7) 1 mM final concentration
If all solutions are sterile, it is not necessary to filter with a sterile filter unit.
Recipe 9: Dissociation solution
Dissection solution (Recipe 8) 5 ml
Papain enzyme 100 μl
For preparation of cortical neurons, add
KYN (100 mM, Recipe 7) 1 mM final concentration
Mix in a sterile 10-ml centrifuge tube. Warm in a water bath until the papain has dissolved. Filter with an in-line sterile syringe filter before adding tissue.
Note: Dissociation solution must be made fresh before each neuron preparation.
Recipe 10: FUDR stock
5-Fluoro-2′-deoxyuridine 25 mg (8.1 mM final concentration)
Uridine 625 mg (20.4 mM final concentration)
MEM 12.5 ml
Mix well, filter with a sterile filter, and store in a -20°C frost-free freezer in 500-μl aliquots.
Recipe 11: Borate buffer [0.1 M (pH 8.4)]
Borax 1.9 g
Boric acid 1.24 g
Distilled H2O 400 ml
Adjust pH to 8.5 and filter through a 0.2-μm sterile filter unit.
Recipe 12: HMW PLYS stock (1 mg/ml)
Borate buffer (Recipe 11) 5 ml
Mix well. Aliquot 1 ml per tube into sterile microcentrifuge tubes. Store in -20°C frost-free freezer.
Note: 5 mg is the total amount of HMW PLYS in the container if purchased from Sigma (no. P-5899).
Recipe 13: Culture medium for slice cultures
BME 50%
HBSS 25%
Horse serum 25%
D-Glucose 33 mM final concentration
Hepes (1 M solution) 10 mM final concentration
Penicillin-streptomycin 100 U/ml
Filter with a sterile filter unit. Maintain 100 ml of medium at 2°C for slicing and 400 ml in a 5% CO2/37°C incubator for feeding cultures.
Recipe 14: Hepes artificial cerebrospinal fluid (ACSF)
NaCl 140 mM
KCl 5 mM
MgCl2 1 mM
D-Glucose 24 mM
Hepes (powder) 10 mM
CaCl2 1 mM
Adjust pH to 7.2 with NaOH, filter with a 0.2-μm sterile filter unit, and store at 2° to 4°C.
Note: For rodent cultures, replace NaCl with equimolar sucrose.
Recipe 15: 1.85% agarose
Weigh enough agarose to make a 1.85% solution (weight to volume). Add to water. Dissolve by heating agarose solution in a microwave. Pour into a large petri dish, allow to solidify, cover, and store at 2° to 4°C indefinitely. Sterilize with UV before use.
Recipe 16: 1 M CaCl2 solution
Prepare a 1 M solution in distilled H2O.
This solution does not need to be sterile.
Recipe 17: PVP solutions
Stock: 20 mg PVP in 1 ml of fresh 100% EtOH
Dilute solution: 10 μl of PVP stock into 8 ml of fresh 100% EtOH (final concentration 0.05 mg/ml)
Note: Optimally, this must be made fresh each day from a newly opened bottle of EtOH. Because this can get expensive, aliquotting a fresh bottle of EtOH quickly and freezing the aliquots in a -20°C freezer also works well.
Recipe 18: Spermidine solution
Prepare a 0.05 M solution in distilled H2O.
Recipe 19: 0.1 M PB
Na2HPO4 10.24 g
NaH2PO4 3.4 g
Dissolve in 1 liter of distilled H2O.
pH to 7.4
Recipe 20: 10× PBS
NaH2PO4 H2O 32.1 g
Na2HPO4 206.5 g
NaCl 1318 g
Dissolve in 15 liters of distilled H2O.
Note: To make a working solution, dilute the stock buffer 1:10 with distilled H2O.
Recipe 21: 5% CaCl2
Dissolve 5 g of CaCl2 in 100 ml of distilled H2O.
Recipe 22: Immunocytochemistry (ICC) fix
Heat 80 ml of 0.1 M PB (Recipe 19) to 60°C.
Add 2.5 g of paraformaldehyde. Mix for 1 min.
Add 4 g of sucrose. Mix for 5 min.
Add NaOH (4 N) dropwise until solution is clear.
Adjust volume to 100 ml with 0.1 M PB.
Add drops of a 5% CaCl2 (Recipe 21) solution until cloudy.
Filter with a 0.2-μm filter.
Store protected from light at 4°C for up to 14 days.
Recipe 23: Blocking solution
Normal goat serum 10%
Bovine serum albumin 2%
Triton X-100 0.25%
Dissolve in 0.1 M PB (Recipe 19).
Recipe 24: 1% HCl in 70% EtOH
95% EtOH 700 ml
Distilled water 250 ml
Concentrated HCl 9.5 ml
Recipe 25: Subbing solution
300-bloom gelatin 15 g (1.5%)
Chromium potassium sulfate 0.5 g (0.05%)
Distilled H2O 1 liter
Heat the water in a microwave for 3 min on high power. Add gelatin slowly, stirring until dissolved. Add chromium potassium sulfate and filter through a 0.2-μm filter.
Note: For 1-by-3-inch slides, make 750 ml of subbing solution using 11.25 g of gelatin and 0.375 g of chromium potassium sulfate.
Recipe 26: 30% sucrose
Prepare a 30% solution in 0.1 M PB (Recipe 19).


Conventional Dissociated Neuron Cultures

There are numerous methods for preparing primary neuronal cultures. The methods for culturing nerve cells described here are modified from the protocols in the second edition of Culturing Nerve Cells by Banker and Goslin (11). This method produces a layer of neurons plated over a monolayer of astrocytes and is modified from Bekkers and Stevens (12). Often, these cultures are healthier and thus easier to transfect with biolistics than are the "Banker" cultures described below.

Note: All of the following procedures should be performed in a sterile hood, unless indicated otherwise.

Preparation of Clean Coverslips

This procedure can be performed under nonsterile conditions.

1. Place coverslips in a 1-liter beaker.

2. Add nitric acid to cover the coverslips, and place a Teflon beaker cover over the beaker.

3. Sonicate for 1 hour, swirling occasionally.

4. Pour nitric acid into a large bucket filled with H2O and dispose of it.

5. Wash coverslips three times with distilled H2O.

6. Add HCl to cover the coverslips, and place a Teflon beaker cover over the beaker.

7. Sonicate for 1 hour, swirling occasionally.

8. Wash 10 times with distilled H2O.

9. Rinse two times with 95% EtOH.

10. Store coverslips in 95% EtOH.

Preparation of Astrocyte Cultures

In order to prepare neuron cultures, you must have a stock of astrocytes to seed the coverslips. So the first step is to prepare astrocyte cultures.

1. Dissect and dissociate cells from the brain region of interest (as described below).

2. Plate approximately 400,000 cells into a 25-cm2 (50-ml) tissue culture flask in about 5 ml of astrocyte medium (Recipe 5).

3. Allow the astrocytes to grow nearly to confluence (approximately 10 days of growth).

4. Place the flask into a plastic bag and place onto a laboratory shaker (nonsterile) at room temperature. Shake at a medium speed (setting no. 7 on a Labline shaker) overnight. This will kill any neurons in the flask.

5. Observe the cells under a microscope to confirm that all process-bearing cells have shaken off the astrocytes.

6. Exchange the medium with fresh astrocyte medium (Recipe 5).

7. Maintain these astrocyte cultures in a 5% CO2/37°C incubator.

Note: These cultures can be maintained for months. Exchange the medium when it starts to turn yellow. The cells will only survive two or three passages, though.

Preparation of a Glial Monolayer on Coverslips

1. Using sterile forceps, place cleaned coverslips into 24-well culture plates at an angle against the side of each well to dry.

2. Shake the plate so the coverslips are flat on the bottom of each well after EtOH has evaporated.

3. Sterilize under UV light for 20 min.

4. Coat the coverslips with the collagen/PLYS mixture (see Recipe 3) by adding 70 μl to each coverslip (it should form a bubble over the coverslip).

Note: This solution should flow over the coverslips smoothly. If the solution beads, the coverslips are not sufficiently clean and should be prepared again (or the cultures will be unhealthy and patchy).

5. Gently aspirate the solution off the coverslips into a pipet and discard. This will leave a thin film of collagen/PLYS coating on the coverslips.

6. Air dry in the hood.

Note: Coated coverslips can be stored for a few weeks at room temperature.

7. Aspirate medium from an astrocyte flask.

8. Add 2.5 ml of trypsin-EDTA.

9. Incubate flask for 5 min in a 5% CO2/37°C incubator.

10. Remove from the incubator and add 2.5 ml of astrocyte medium.

11. Triturate five times with a 5-ml pipet.

12. Transfer to a sterile 15-ml centrifuge tube.

13. Centrifuge on low speed (1500 rpm in a clinical centrifuge) for 5 min.

14. Aspirate the solution and resuspend the cells in 3 ml of astrocyte medium (Recipe 5).

15. Count the cells on a hemocytometer.

16. Plate cells onto collagen/PLYS-coated coverslips at a density of 10,000 cells per coverslip in 500 μl of astrocyte medium (Recipe 5) per well of a 24-well plate.

17. Maintain the cells in a 5% CO2/37°C incubator until astrocytes have formed a confluent monolayer on the coverslips. Once a monolayer is formed, the cultures are ready for neurons to be plated.

Note: Astrocytes can be replated into flasks to form another monolayer; they remain healthy for only two or three passages.

Preparation of Dissociated Neurons

The following procedure is specifically for cortical cultures. This same basic protocol will also work for making neuron cultures from other brain regions, although adding kynurenate seems to selectively improve the health of cortical neurons.

1. Prepare the dissection solution (Recipe 8) and dissociation solution (Recipe 9) by adding KYN (Recipe 7) to optimize the health of cortical neurons.

2. Place all equipment in a hood and wipe with 95% EtOH to sterilize.

3. Place freshly prepared dissociation solution in a 37°C water bath.

4. Add 10 ml of the dissection solution to two medium petri dishes (10 ml per dish).

5. Anesthetize a newborn rat (or mouse) pup.

Note: Although hippocampal neuron cultures are most healthy when prepared from newborn rodents, cortical neurons can be prepared from pups up to at least postnatal day 8.

6. Wipe the animal's head with a Kimwipe soaked with 95% EtOH.

7. Decapitate with a razor blade and immediately place head on a Kimwipe in a hood.

8. Remove the brain (using the angled scissors, blunt forceps, and hippocampal-dissecting tool).

9. Place the brain in a petri dish containing dissection solution (Recipe 8).

10. Place the petri dish onto a dissection microscope.

11. Using fine forceps, carefully remove the pia and dissect out the desired region of the cortex with the ultrafine scissors.

12. Using the hippocampal-dissecting tool, place the block of cortex into the second petri dish.

13. Cut the blocks of brain into small pieces.

14. Sterile filter the dissociation solution (Recipe 9).

15. Add the tissue pieces to the filtered dissociation solution and incubate in a 5% CO2/37°C incubator for 20 min.

16. Carefully aspirate the dissociation solution, and rinse the tissue pieces with 5 ml of neuronal medium (Recipe 6).

17. Allow the pieces of tissue to settle on the bottom of the tube.

18. Aspirate the medium.

19. Add exactly 2 ml of neuronal medium (Recipe 6).

20. Gently triturate the tissue with a 5-ml pipet until there are no more large pieces visible (7 to 10 cycles). Avoid creating bubbles.

21. Pass the dissociated cells through a cell strainer to remove any large clumps of tissue.

22. Count the cells with a hemocytometer. Count all bright cells and average two fields to obtain a final density.

Note: The healthiest neuron preparations have a large proportion of cells with apical dendrites still attached (they look like tails).

23. Plate the cells onto coverslips with established astrocyte monolayers at desired density in 500 μl of neuronal medium per well of a 24-well plate (Recipe 6). Low-density cultures range from 5000 to 8000 cells per coverslip.

24. Add 3 μl of FUDR stock (Recipe 10) to the cultures 5 days after plating neurons to avoid proliferation of oligodendrocytes and macrophages.

Note: Adding FUDR before 5 days will damage the neurons, but waiting longer than 5 days will allow these other glial cells to proliferate.

Preparation of Banker Cultures

This protocol is adapted from chapter 13 in Culturing Nerve Cells (11) and produces a pure population of neurons on coverslips grown in astrocyte-conditioned medium. Most of the general methods are the same as for conventional cultures, with a few notable exceptions. This culture system is based on a "sandwich" configuration, in which astrocytes grow directly in the wells of culture plates while neurons grow on coverslips elevated above the astrocytes (11). In this manner, the neurons are nourished by astrocyte-conditioned medium but do not directly contact the glial cells.

Note: All of these procedures should be performed in a sterile hood, unless otherwise indicated.

Preparation of Modified Culture Plates

1. Melt three small indentations with a soldering iron into the bottom of each well of the plastic, sterile, tissue culture plates in order to elevate the neuron-containing coverslips over the bottom of the culture well.

Note: Be sure not to melt a hole through the bottom of the well, which could cause media leakage and contamination.

2. Sterilize the plates under UV light for 30 min, and store the plates indefinitely.

Preparation of Coverslips Coated with HMW PLYS

1. Using sterile forceps, place cleaned coverslips (procedure described above) into a separate set of 24-well culture plates at an angle against the side of each well to dry.

2. Shake the plate so that the coverslips are flat on the bottom of each well after EtOH has evaporated.

3. Sterilize under UV light for 30 min.

4. Coat sterilized coverslips with HMW PLYS (see Recipe 12) by adding 56.5 μl of stock per 12-mm coverslip in a sterile hood.

5. Allow the solution to air dry completely (usually overnight).

6. Rinse the coated coverslips twice for 2 hours with sterile distilled H2O.

7. Use these coverslips immediately or store in neuronal medium in a 5% CO2/37°C incubator for several days.

Preparation of Astrocytes in the Modified Culture Plates

1. Plate astrocytes in astrocyte medium (see Recipe 5) at 500,000 cells per well (in 500 μl of medium) directly into the modified culture plates.

2. Maintain in a 5% CO2/37°C incubator until cells are confluent.

3. Replace the astrocyte medium in the astrocyte plates with neuronal medium.

4. Incubate for at least 1 day to condition the medium before making neuron cultures.

Preparation of Neurons for the Banker Culture

1. Dissect and dissociate neurons as described above for conventional cultures.

2. Plate the neurons at a desired density (approximately 8000 cells per 12-mm coverslip for low densities) in neuronal medium (Recipe 6), 500 μl per well.

3. Maintain the cells on the coverslips in a 5% CO2/37°C incubator for 4 hours.

4. Remove the coverslips with sterile forceps and place cells side-down onto the melted plastic risers in the astrocyte plates (into the conditioned neuronal medium).

5. Add 3 μl of FUDR (Recipe 10) to the cultures 5 days after plating neurons to inhibit the proliferation of astrocytes on the coverslips.

Preparation of Slice Cultures

Organotypic slice preparations are a powerful experimental system, because they preserve the three-dimensional architecture and local environment of neurons to a far greater extent than do dissociated cell cultures, yet still allow easy access for experimental manipulations and observations (13). There are several methods for preparing brain slices, such as using a tissue slicer, a tissue chopper, or a vibratome. In my experience, the method that produces the healthiest slices for culturing is the tissue slicer, a device designed by Lawrence C. Katz (10) and distributed by SD Instruments (Fig. 1A). Although the reasons why this method produces healthier slices remain unknown, it may be because it is extremely fast; the time from brain removal to placing slices in the incubator is only a few minutes.

Fig. 1.

Tissue slicer. A very efficient and reliable method for obtaining healthy organotypic slice cultures is to use the tissue slicer designed by Katz (10) and distributed by SD Instruments. (A) The slicer works by propelling thin gold wires, spaced 400 μm apart, into a block of tissue. In this case, the tissue block was a piece of visual cortex and the slices were cut in a coronal orientation. (B) The tissue is placed onto a small slab of agarose for protection. (C) After the spring is unloaded, sending the wires through the tissue, individual slices are carefully separated with a paintbrush. This picture shows a sliced piece of tissue above the wires and some slices that were separated from the tissue below the winder.

There are also multiple methods for maintaining slices in culture: interface (13) and roller (14) cultures. Because roller cultures shrink slices to a monolayer over time, we use interface cultures to preserve the three-dimensionality of the tissue slice. To prepare this culture type, slices are placed on culture inserts so that the top surface of the slice is exposed to the incubator atmosphere while the lower surface contacts the culture medium (8, 9). In this configuration, slices remain healthy for weeks and continue to differentiate, forming layer-specific connections (15, 16).

Preparation of Slicer

1. Prepare Hepes ACSF (Recipe 14) and place on ice.

2. Prepare slice culture medium (Recipe 13) and place on ice.

3. Prepare agarose (Recipe 15) and place on ice.

4. Make winders according to the procedure included with the apparatus.

5. Autoclave surgical instruments, glassware, and pipets.

6. Place all equipment (microscope, glassware, surgical instruments, culture supplies, brushes, and slicer) in a hood; wash everything with 70% EtOH and expose to UV for 20 min.

7. Cut a slab of agarose to fit the slicer, place on removable slicer component, and place both on ice.

8. Place winder into slicing apparatus.

Preparation of Slice Cultures

Note: Wear gloves, wipe hands with 70% EtOH often, and wear a face mask. The slices will invariably be healthier if all of the solutions are ice-cold. This also makes the tissue more solid and easier to handle.

1. Anesthetize the animal.

2. Rinse the animal's head with 70% EtOH and decapitate.

3. Transfer the head into the hood and place on paper towel.

4. Remove the brain and place in a petri dish filled with ice-cold Hepes ACSF (Recipe 14).

5. Using the dissecting microscope, peel the pia from the desired brain region using fine forceps and remove the desired block of tissue.

6. Place the tissue block onto the agarose and insert into the slicer (Fig. 1B).

7. Hit slicer spring, sending winder into tissue and cutting 400-μm-thick slices.

8. Unscrew winder from slicer and remove the tissue and winder.

9. Carefully place the tissue and winder into a clean petri dish with fresh, ice-cold Hepes ACSF (Recipe 14) (Fig. 1C).

10. Gently separate the slices with a paintbrush, touching the brain tissue as little as possible.

11. Place culture inserts into the culture plates.

12. Carefully suck the slices into the large-bore end of a Pasteur pipet and gently drop them onto an insert.

13. Using a paintbrush, gently separate and flatten slices so they can't grow together.

14. Using the thin-bore side of a Pasteur pipet, remove as much Hepes ACSF as possible from the membrane without touching the slices.

15. Place 1 ml of culture medium (Recipe 13) under each insert; the medium should wet the insert without floating the slices. Remove any bubbles from beneath the insert.

16. Maintain slices in a 5% CO2/37°C incubator.

17. Change the medium every 3 days.

Biolistic Transfection

Biolistic transfection is a relatively simple, flexible, and effective method for transfecting neurons and glia in mammalian brain slices, resulting in high levels of expression of reporter proteins in transfected cells. In general, biolistics consists of three steps: coating gold particles with DNA, preparing the cartridges, and accelerating the particles into the neurons or tissue.

The details of performing this procedure are described in the Helios Gene Gun System Instruction Manual from Bio-Rad Laboratories. I use several modifications to these basic procedures that enhance the reliability of transfection of both dissociated neurons and organotypic slice cultures (8, 9).

Supercoiled plasmid DNA constructs can be purified with commercial plasmid isolation kits, such as Qiagen spin miniprep kits or Bio-Rad midiprep kits. DNA amplified in this manner is usually pure enough to facilitate reliable transfections. However, if transfection is unsuccessful, it is possible that the DNA might need to be cleaned of impurities.

Preparation of DNA-Coated Gold Microprojectiles

Plasmid DNA is precipitated from solution in the presence of gold particles and the polycation sperimidine by the addition of CaCl2. The particles are then washed extensively with EtOH to remove all water and then are resuspended in PVP solution (Recipe 17) to enhance the ability of the gold to adhere to the inside of the tubing.

For cortical and hippocampal neurons, 1.6-μm gold particles transfect the greatest number of healthy neurons. It is possible that cells with smaller diameters will be optimally transfected with smaller gold particles, which are also available from Bio-Rad.

1. Make a fresh set of PVP solutions (Recipe 17).

2. Coat the gold particles with spermidine by mixing 0.0125 g of gold (particles 1.6 μm in diameter) and 100 μl of spermidine solution (Recipe 18) in a 1.5 ml-microcentrifuge tube.

3. Vortex and sonicate for 5 s each.

Note: This corresponds to half of the Bio-Rad-recommended concentration of gold particles. The full amount of gold causes too much tissue damage in slices and destroys the glia in dissociated cultures. The concentration recommended here is optimal in my experience.

4. Add 25 μg of DNA to the tube and vortex for 5 s.

5. Add 100 μl of 5% CaCl2 (Recipe 21) dropwise.

6. Allow the gold and DNA to precipitate for 10 min, flicking tube periodically to mix the contents.

7. Spin for 15 s at maximum rpm in a minicentrifuge to pellet the gold.

8. Remove the supernatant and discard it.

9. Wash the gold particles three times with 1 ml of fresh 100% EtOH. Spin for 5 s at maximum rpm between each wash.

10. Resuspend the gold particles in 200 μl of dilute PVP solution (Recipe 17) and transfer to a 15-ml centrifuge tube.

11. Rinse all of the gold particles out of the microcentrifuge tube with 800 μl of dilute PVP solution (Recipe 17) and transfer to a 15-ml centrifuge tube.

12. Add 2 ml of dilute PVP solution (Recipe 17) to a final volume of 3 ml of gold particles in PVP solution. This solution will be called the DNA/PVP/EtOH mixture.

Preparation of Cartridges

This step is clearly outlined in the Bio-Rad Helios Gene Gun Instruction Manual.

1. Purge a 30-inch length of gold-coat tubing (provided in the Bio-Rad optimization kit) with nitrogen for 30 min. Insert the tubing into the tubing prep station and push into the tubing support cylinder and through the O-ring. Turn on the nitrogen and adjust the flow to 0.35 liters per minute (LPM).

2. After 30 min, turn off the nitrogen with the flowmeter on the prep station.

3. Attach a syringe to one end of the tubing (using the adapter tubing provided by Bio-Rad).

4. Vortex the gold suspension.

5. Suck 3 ml of the DNA/PVP/EtOH mixture into the open end of the purged gold-coat tubing, using suction from the syringe. Fill the tubing steadily until the DNA/PVP/EtOH mixture is 2 inches from the syringe.

6. Keeping the syringe attached and the tubing horizontal, insert the loaded tubing into the tubing prep station until the end is through the O-ring.

7. Allow the gold to settle for 5 min. Keep the syringe attached.

8. Slowly and steadily suck out the solution, leaving the settled gold undisturbed. Detach the syringe and discard the solution.

Note: The Bio-Rad manual suggests that the solution be removed with a peristaltic pump. This is complicated and unnecessary as long as the solution can be removed with the syringe without disturbing the gold.

9. Immediately turn the tubing 180° using the switch on the prep station, allowing the gold to coat the tubing for 5 s.

10. Turn the switch on the prep station to "on" to rotate the tubing for 30 s.

11. Open the valve on the flowmeter to allow 0.35 LPM nitrogen to dry the gold onto the inside of the tubing while it rotates for 5 min.

12. Turn the motor off. Remove the tubing.

13. Cut off any pieces of tubing that are not uniformly coated with gold. Use the tubing cutter (provided with the system by Bio-Rad) to cut the tubing into 0.5-inch cartridges.

14. Store the cartridges in a cartridge storage vial (supplied by Bio-Rad) with a desiccator pellet, label, and wrap with parafilm. These cartridges can be stored for up to a year at4°C.

Blasting Cells or Slices with DNA-Coated Gold

1. Cut a piece of 2-by-2-inch nylon mesh and attach it with tape to the barrel liner, covering all of the exposed parts. Sterilize this mesh and the cartridge holders by exposure to UV light for 30 min.

Note: The remaining steps should be performed in a sterile laminar flow hood.

2. Insert a 9-V battery and an empty cartridge holder into the Helios Gene Gun.

3. Attach the Gene Gun to the helium tank with the special helium hose provided by Bio-Rad. Fire a few shots at 100 psi to pressurize the helium hose and the reservoirs in the gun.

4. Load the cartridges containing the DNA-coated gold into the cartridge holders (provided by Bio-Rad) and load the holder into the Gene Gun.

5. Attach the barrel liner with the nylon mesh to the Gene Gun.

6a. Primary neuron cultures:

1. Using sterile forceps, place a coverslip containing neurons into a small sterile petri dish.

2. Immediately place the gun exactly perpendicular and at a distance of 1 cm above the cells.

3. Shoot the DNA-coated gold into the culture using a pressure of 50 psi for dissociated neurons.

4. Immediately replace the cells in their original medium and return them to the incubator.

5. Continue until all desired cultures are transfected.

Note: The pressure of the blast and distance from the cells should be optimized for each new culture type. These parameters are optimal for cortical and hippocampal cultures.

6b. Slice cultures:

1. Using sterile forceps, place a filter insert containing slice cultures onto a slab of agarose [presterilized by UV; see instructions for preparation of slice cultures for directions on agarose preparation (Recipe 15)].

2. Immediately place the gun above the slices at an angle exactly perpendicular to the cells and bring the end of the barrel flush with the top of the filter insert.

3. Shoot the DNA-coated gold into the slices at a pressure of 100 psi.

4. Immediately replace the inserts in their original medium and return them to the incubator.

5. Continue until all desired slices are transfected.

Note: The pressure of the blast and distance from the slices should be optimized for each new tissue type. These parameters are optimal for cortical and hippocampal slices.

7. Close the helium tank, release the pressure from the Gene Gun, and detach the Gene Gun from the helium tank.

8. Remove the cartridge holder and discard the cartridges.

9. Check for successful transfection by visualizing the gold in the neurons or slices using an inverted tissue culture microscope. The gold particles should be evenly dispersed in the cultures, with no central zone of dead cells, or should be evenly distributed in the slice to a depth of 300 μm.

Visualizing Transfected Reporters by Immunostaining

Cells transfected with biolistics are easily identified by the presence of a gold particle in their nucleus and by the reporter gene product. In many early experiments, the reporter gene we used was β-galactosidase (β-Gal), which can be visualized with X-Gal histochemistry or with better resolution using immunostaining with an antibody to β-Gal [(5 Prime-3 Prime) see Figs. 2 and 3 (8, 9)]. More recently, the use of the green fluorescent protein (GFP) as a reporter has become widespread ( This gene can also be fused to various proteins to function as a fluorescent tag on those proteins (Fig. 4). Because GFP is naturally fluorescent, it labels living neurons and can be visualized without requiring immunostaining or histochemical procedures. A general method for performing immunostaining of slice cultures is described here.

Fig. 2.

An organotypic slice transfected using biolistics. This is a coronal slice of the visual cortex made from a ferret at postnatal day 14. The slice was transfected with a lac-Z expression construct (19) using biolistics immediately after slicing. β-Gal was allowed to fill transfected neurons for 36 hours before staining with X-Gal histochemistry. This picture illustrates the large number of neurons (in the hundreds) that can be successfully transfected in a single slice with biolistics. Extensive controls were performed to ensure that this method of transfection did not adversely affect the health of the neurons (8, 9). Scale bar, 125 μm.

Fig. 3.

Neurons transfected using biolistics. These are pyramidal neurons from slices of ferret visual cortex (postnatal day 14) that were transfected with a lac-Z expression construct (19). (A) All of the fine processes of this layer 5 pyramidal neuron are completely filled with β-Gal, including the axon (the thin process projecting out of the base of the cell soma) that can be followed for millimeters across the slice. (B) Immunostaining transfected neurons with an antibody to β-Gal (5 Prime-3 Prime) increases the resolution of staining of these neurons dramatically over X-Gal histochemistry as shown by this layer 4 pyramidal neuron. Scale bars, 15 μm. Adapted and reproduced with permission from (8).

Fig. 4.

A dissociated neuron transfected with a GFP-fusion construct using biolistics. These are dissociated hippocampal neurons from a newborn rat pup, cultured in a conventional preparation for 12 days in vitro. The neurons were transfected with a VAMP-pEGFP construct generously provided by R. Scheller (Stanford University) and then imaged with a laser-scanning confocal microscope 24 hours after transfection. (A) The gene product--GFP fused to the synaptic vesicle protein VAMP--is distributed throughout the cell soma and some proximal dendrites. The gold particle in the nucleus that transfected this neuron is indicated by an arrow. (B) A fluorescence image of the axonal arborization of this neuron illustrates the punctate nature of the localization of VAMP-GFP. This is a synaptic distribution, because over 90% of these punctae stained and destained with the dye FM4-64. (C) By superimposing the fluorescence image in (B) with the corresponding transmitted light image, one can see that the synapses made by the transfected neuron are located along the dendrites and cell bodies of nontransfected neurons in this field.

Preparation of Subbed Slides

1. Wash slides in hot soapy water.

2. Rinse slides in several changes of hot water.

3. Rinse slides in distilled water.

4. Air dry.

5. Soak slides in 1% HCl in 70% EtOH (Recipe 23) for 5 min.

6. Rinse slides in 100% EtOH.

7. Rinse slides in acetone.

8. Air dry.

9. Dip slides several times into the subbing solution (Recipe 25).

10. Dry overnight in a 37°C oven.

Fixation and Staining

1. Fix slices in ICC fix (Recipe 22) for 1.5 hours at room temperature.

Note: Remove the culture medium and add fix under the insert first, then gently drop fix onto the slices until the tissue is submerged.

2. Remove the fix and replace with 30% sucrose (Recipe 26). Incubate at room temperature for 2 hours.

3. Freeze-thaw the samples into PBS (Recipe 20). Cut slices out of the filter using a sharp scalpel blade by cutting a small square around the slice. Be careful not to touch the slice itself. Place filter with slices attached onto a slide and place directly onto dry ice with the slice side up. Once frozen, remove the slide and allow the slice to thaw. Be sure to place the slices in PBS before they start to dry.

Note: Keeping the slice attached to the filter makes the immunostaining procedure easier and enhances the tissue integrity during staining. This step promotes penetration of the tissue by the antibodies.

4. Remove PBS and replace with blocking solution (Recipe 23). Incubate for 2 hours at room temperature.

5. Remove blocker and replace with primary antibody diluted in blocking solution (Recipe 23) at 4°C overnight.

6. Wash at room temperature in blocking solution four times for 10 min each wash.

7. Remove blocking solution and replace with secondary antibody diluted in blocking solution (Recipe 23). Incubate at room temperature for 4 hours.

8. Wash three times with PBS (Recipe 20) for 15 min each wash.

9. Dry the slices (filter side down, tissue side up) onto subbed slides.

10. Dip slides into xylenes for 3 min in a fume hood.

11. Place a coverslip over the slices using Krystalon as mounting medium.

Related Techniques

Although there is no method as reliable as biolistics for transfecting neurons to date, there are several alternative methods that have distinct advantages over biolistic transfection. Depending on the application, one potential disadvantage of biolistics is that the transfection efficiency is quite low, which is expected for a random physical method of transfection. Both calcium phosphate and viral transfection techniques support much greater transfection efficiencies. Transfection using calcium phosphate is easy to perform and yields large numbers of transfected neurons (17). However, many laboratories report that this method is capricious: It will work beautifully for weeks but then will suddenly stop working for months. Viral transfection also provides high transfection efficiencies and is particularly advantageous for in vivo transfection of groups of neurons (18). Unfortunately, there are conflicting reports of neurotoxicity concerning all of the currently available viruses.

Notes And Remarks

A number of factors affect the success of biolistic transfection in a given tissue. These parameters must be optimized for each cell and tissue type, and controls must be designed to ensure that the bombardment process does not cause cell or tissue damage. For organotypic brain slices and dissociated cultures, the most important parameter is neuronal health. If the slices or cells are unhealthy for any reason, levels of transfection will be reduced. However, once the specific parameters of the biolistic transfection are optimized for a specific tissue type, the method itself is quite reliable and reproducible and does not require advanced molecular biological facilities for its application. More often than not, if no transfected cells are obtained, it is the health of the cultures that is the problem.

Transfection rates obtained with biolistics vary according to the density of cells plated (for dissociated cultures), the amount of gold used in each blast, the efficiency of the expression construct, and the exact blasting pressure. The two most important parameters that must be optimized for each tissue and cell type are the number of gold particles and the helium pressure. For each cell type, there is a threshold gold concentration that will provide the greatest number of transfected cells without tissue damage. Increasing the amount of gold above that threshold can cause extensive tissue damage in brain slices and massive cell death in dissociated neuron cultures. Similarly, there is an optimal helium pressure for each tissue type that will provide the greatest penetration of the slices without extensive tissue damage. Pressures above that threshold produce a helium shock wave that is very destructive to neurons in slices and will literally blow neurons off of coverslips in dissociated cultures. It is important to note that, even in cases in which a large amount of cell death is observed, a few neurons are often transfected. Thus, the simple observation of transfected cells is not sufficient to ensure the health of the tissue or cells. The nylon mesh described above minimizes this shock wave, but the helium pressure must still be optimized and careful controls performed to ensure that biolistics is not damaging the transfected cells.

Biolistic transfection of neurons in brain slices and dissociated cultures has several advantages. Because plasmid DNA is used for gene transfer, biolistics is considerably less labor-intensive than viral techniques and avoids the neurotoxicity associated with many viruses. Also, in contrast to current transgenic techniques, biolistics allows flexibility in the time and location of gene manipulation: The age of the animal and the region of the brain from which slices and cultures are prepared determines when genes are manipulated. Most important, biolistics allows the transfection of individual neurons in an otherwise normal background tissue. Finally, biolistic transfection can also be used in combination with transgenic approaches to rescue the phenotype of individual cells in knockout mice, thereby clarifying the effects of gene deletion. Because experimental studies of genes depend on the ability to transfect cells with DNA constructs designed to test their function, biolistic transfection has made it possible to answer many questions regarding gene function in previously intractable preparations, such as postmitotic neurons.


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