IntroDuctIonThe hippocampus and cortex have fascinated neuroscientists for years because of their precise organizations and functional roles in cognition, learning and memory1. As with most structures in the brain, the complexity of these structures makes it challenging to analyze and manipulate easily in vivo. Cell lines derived from central nervous system precursors have limitations because the neurons derived from these lines fail to recapitulate characteristics of central neurons, including the ability to form well-defined axons, dendrites and synapses. Instead, primary cell culture techniques have been successfully developed to study these neurons in vitro.
Primary neuronal cell culturesOne of the most well-established and widely used techniques for the study of hippocampal and cortical pyramidal neurons has been the primary dissociated cell culture system pioneered by Kaech and Banker2 for the culture of embryonic rat neurons. This culture system allows neurons to be cultured in vitro in a far less complex environment than that used in vivo, making them highly accessible to manipulations and observations. The vast majority of cells in these preparations are excitatory pyramidal neurons, with inhibi-tory neurons representing about 18–20% of the population. Several studies, including those from our group3–8, have shown that dis-sociated embryonic neurons maintained in culture undergo dis-tinct stages of differentiation and form well-established synaptic connections9–11. These cells can be grown and maintained in single layers on glass coverslips or plated at higher densities for biochemi-cal studies. Furthermore, coupled with advances in imaging tech-nologies and manipulation of gene expression, these cultures have allowed unprecedented views into the intricate inner workings of neuronal cells. Thus, this technique has been well exploited to dis-sect out various aspects of molecular and cellular mechanisms that underlie neuronal morphogenesis and connectivity12.
The culture of postnatal versus embryonic neurons is advanta-geous for several reasons. It reduces the necessity of killing animals.
It is possible to euthanize one or a few animals from one litter and save the rest for further breeding, which is not possible with embry-onic tissue. Further, the mother need not be killed and can be used for further breeding or other purposes. Moreover, the technique makes it possible to (i) study neurons from genetically engineered mice that are early postnatal lethal; (ii) control expression of genes in a temporally controlled manner using RNAi-mediated knock-downs or tetracycline-regulated expression plasmids in wild-type neurons; or (iii) delete genes in individual neurons using transfec-tion of a vector expressing Cre recombinase in neurons from floxed mice. The ability to delete genes from small subsets of neurons makes it possible to distinguish cell-autonomous versus non-cell-autonomous effects of individual genes and their disease counter-parts on neuronal structure and function13,14.
Culturing postnatal mouse neurons has proved to be more chal-lenging than culturing embryonic mouse or rat neurons. Neurons at earlier developmental stages, as in the case of embryos, are less susceptible to damage during the processing required for their cul-ture. This is partly because their neuronal processes are relatively less well developed in comparison with postnatal animals15,16. In practical terms, the meninges are also more adherent to the tissue in postnatal animals. Thus, postnatal cultures tend to be less robust and have a lower yield of healthy viable cells in comparison with embryonic rat or mouse hippocampal and cortical neurons and have a higher proportion of non-neuronal cells.
Types of primary neuronal culturesTwo types of cultures have been described for culturing primary neurons from rats and mice2. In one method, the neurons are grown sandwiched on an astrocytic feeder layer17. This allows for the neurons not to be in direct contact with the astrocytes, but to be exposed to factors secreted by astrocytes into the medium. The alternate protocol does not involve the use of the feeder layer, and neurons are maintained in serum-free medium (B27) supplemented
Culturing pyramidal neurons from the early postnatal mouse hippocampus and cortexGerard M J Beaudoin III1,3,4, Seung-Hye Lee1,3, Dipika Singh2, Yang Yuan2, Yu-Gie Ng1, Louis F Reichardt1 & Jyothi Arikkath2,3
1Department of Physiology, University of California–San Francisco (UCSF), San Francisco, California, USA. 2Munroe-Meyer Institute, University of Nebraska Medical Center (UNMC), Omaha, Nebraska, USA. 3These authors contributed equally to this work. 4Present addresses: Neurosciences Institute, University of Texas at San Antonio, San Antonio, Texas, USA (G.M.J.B.); Genentech, Inc., South San Francisco, California, USA (S.-H.L.). Correspondence should be addressed to J.A. ([email protected]).
Published online 30 August 2012; doi:10.1038/nprot.2012.099
the ability to culture and maintain postnatal mouse hippocampal and cortical neurons is highly advantageous, particularly for studies on genetically engineered mouse models. Here we present a protocol to isolate and culture pyramidal neurons from the early postnatal (p0-p1) mouse hippocampus and cortex. these low-density dissociated cultures are grown on poly-l-lysine–coated glass substrates without feeder layers. cultured neurons survive well, develop extensive axonal and dendritic arbors, express neuronal and synaptic markers, and form functional synaptic connections. Further, they are highly amenable to low- and high-efficiency transfection and time-lapse imaging. this optimized cell culture technique can be used to culture and maintain neurons for a variety of applications including immunocytochemistry, biochemical studies, shrna-mediated knockdown and live imaging studies. the preparation of the glass substrate must begin 5 d before the culture. the dissection and plating out of neurons takes 3–4 h and neurons can be maintained in culture for up to 4 weeks.
with cofactors necessary for neuronal growth and maintenance18. Both techniques have been widely used successfully. The genera-tion of the astrocyte feeder layer can be cumbersome, but it can also yield high-quality primary neurons and is advantageous for studying neuron-glia interactions19. Some laboratories have also pioneered the use of the NS21 supplement as an alternative to the use of B27 supplement for both rat20 and embryonic mouse cultures21 and postnatal mouse cultures22.
Optimized protocol for cultureHere we describe optimized protocols for the culture (PROCEDURE) and transfection (Box 1) of postnatal mouse hip-pocampal and cortical neurons. An overview of the procedure is shown in Figure 1. These protocols are based on the widely used, non-feeder layer–based, rat embryonic culture procedures with modifications that improve the health, viability and yield of post-natal mouse neurons4,7,8. These cultures can be prepared by pooling
Box 1 | Transfection of neurons ● tIMInG 1 hNeurons cultured as described in the main PROCEDURE are easily amenable to transient transfections. For all transfections, we prepare the DNA following the manufacturer’s instructions. We have had success with using the EndoFree DNA preparation kit from Qiagen. For low-efficiency transfections, such as those required for single-neuron imaging studies, we routinely take advantage of a lipid-based technique. With this technique, transfection efficiencies vary from greater than 10% in neurons transfected before DIV 7 to 0.5–5% in neurons transfected at DIV 8–14. We routinely use transfections for overexpression and shRNA-mediated knockdown. For shRNA- mediated knockdown, we have had success with the pSUPER.gfp+neo (Oligoengine) vectors.
For high-efficiency transfections, such as those required for biochemical studies, we take advantage of an electroporation-based technique, commonly called nucleofection. The obvious disadvantage of the electroporation-based technique that we have described is that the neurons can only be subjected to the treatment before plating and establishment of neurites. However, new technologies available from Lonza Biosciences claim to support the nucleofection of established neurons in a 96-well format. Coating of DNA onto magnetic beads also permits transfection of plated neurons26. Such transfections might be very useful for library screens and high- efficiency transfections of mature neurons. More recently, we have successfully used recombinant lentiviruses to infect rat neurons in culture, and these could be adapted for use in mouse neurons8. In addition, it might be possible to use the nucleofection technique to introduce plasmids in inducible vectors (e.g., the tetracycline-regulated expression vectors) at plating and induce or suppress the expression of the gene of interest at much later stages in culture, thus allowing temporal control of gene expression in large cell populations. Clearly, advances in such technologies will allow further exploitation of this neuronal cell culture technique to assess various aspects of neuronal physiology, structure and function.
We present here a low-efficiency transfection protocol using Lipofectamine. For high-efficiency transfections, we have successfully overexpressed and knocked down protein levels using the nucleofection technique (Lonza Biosciences). We routinely use low-efficiency transfections for imaging studies. This allows the delineation of entire arbors from individual neurons. High-efficiency transfections are suitable for biochemical studies.
low-efficiency transfection of neurons for imaging1. Adjust the volume of the culture medium in each well of a 12-well plate containing growing neurons to 1 ml.2. Add 1 µl of Lipofectamine to an Eppendorf tube containing 25 µl of Neurobasal medium. Incubate for 5 min.3. In a separate tube, add 0.1 to 1 µg of DNA for transfection to 25 µl of Neurobasal medium. We prefer using a high concentration of DNA for transfections (1–3 mg µl − 1).4. Combine solutions from Steps 2 and 3 with gentle pipetting. crItIcal step It is important that the solutions be combined with gentle mixing so as to not disrupt the lipid-DNA complexes.5. Incubate in the laminar flow hood for 20–30 min at room temperature.6. Add the mix to each well and swirl gently.7. Return cells to the incubator.8. After 24 h, replace half of the medium in each well with fresh maintenance medium at 37 °C. Continue to maintain the neurons as described in PROCEDURE Step 42.
High-efficiency transfection:1. Prepare Nucleofector working solution by adding supplementary solution to the main reaction buffer obtained from the manufacturer.2. Pellet 1 to 2 million cells obtained after PROCEDURE Step 39 gently (500g for 5 min) in plating medium.3. Aspirate the medium carefully and resuspend the cells in 100 µl of the mouse Nucleofector solution. crItIcal step It is important to be very quick from this point onward so as to minimize the time that the neurons are in the Nucleofector solution.4. Add 4 µg of plasmid DNA and gently mix to resuspend the pellet.5. Quickly transfer the cell-DNA mix to the cuvette and pulse it in the Nucleofector using the recommended program (O-005).6. Add a small volume of maintenance medium to the cuvette and transfer the cell suspension to the dish containing maintenance medium, and then mix gently. These cells can be plated on Petri dishes for biochemical studies or Petri dishes containing coated coverslips for imaging studies. A substantial number of the surviving neurons should be transfected. We routinely obtain 60–80% transfection efficiencies with this technique.7. Incubate in a cell culture incubator at 37 °C.8. Change the medium and maintain cells as described in PROCEDURE Step 42. ? trouBlesHootInG
tissue from several animals or by using individual animals to generate separate cultures. Individual mouse cultures allow the comparison of control and mutant neurons from animals of the same litter. This culturing protocol is relatively straightforward and does not use an astroglial feeder cell layer. The serum-free main-tenance medium for neurons is not conducive to the growth and survival of astrocytes18, and thus the surviving cell population is predominantly neuronal. However, one can adapt this protocol to include a feeder layer, if useful, for example, to study the effects of secreted astrocyte factors on neurons. Hippocampi are dissected, after which neurons are trypsinized, dissociated and plated and maintained in medium containing the B27 supplement. By using this protocol, postnatal neurons can be maintained well in culture for up to about 4 weeks. During this period, they show the immunohistochemical characteristics of neurons, develop well- differentiated axons and dendrites, establish spines and form func-tional synaptic connections. They are also highly amenable to both low-efficiency and high-efficiency transfections. We demonstrate the application of these cultures to shRNA-mediated knockdowns (Box 1), immunohistochemistry (Box 2) and time-lapse imaging (Box 3). Other potential applications of these cultures include in situ hybridization, nucleic acid and proteomic analyses, neuro-toxicology and electrophysiology4,23.
Limitations. Although the neurons can be used for a variety of applications, as discussed above, the culture system does have some disadvantages. Neurons prepared using this technique have a higher proportion of non-neuronal cell types than cultures prepared
from embryos because of the advanced developmental stage of the animals. To avoid glial overgrowth, it is possible to use inhibitors, such as araC, to inhibit the growth of dividing non-neuronal cells in long-term cultures. In addition, it is very difficult to maintain these cultures beyond 28 days in vitro (DIV); hence, studies must be limited to this time period. Also, primary cultured neurons, either embryonic or postnatal, are highly sensitive even to subtle changes in
Preparation of coverslips
Coating with poly-L-lysineWashing and incubation in medium
Dissection of the hippocampus/cortex
Cell dissociation and plating
Maintenance of neurons
Figure 1 | Overview of the technique. Coverslips are treated, prepared and coated with poly-l-lysine. After washing, they are incubated in medium. Hippocampi or cortices are dissected; cells are trypsinized and dissociated, and plated on coverslips. The neurons are maintained in culture for up to 28 d.
Box 2 | Immunostaining and mounting of mouse neurons cultured on coverslips ● tIMInG 24–36 h1. Warm the paraformaldehyde/sucrose mixture to 37 °C. crItIcal step It is important to ensure that the solution is warmed to 37 °C to preserve neuronal morphology.2. Gently aspirate the medium from wells containing coverslips on which neurons are plated.3. Add 1 ml of the warmed paraformaldehyde/sucrose mixture quickly, but carefully, to the well to ensure that neurites are not damaged.4. Incubate on bench at room temperature for 10 min.5. Aspirate the paraformaldehyde/sucrose mixture and add 1 ml of 0.1% (vol/vol) Triton X-100 solution in PBS to the well. Incubate on bench for 10 min at room temperature.6. Wash the coverslips once gently with PBS and add 1 ml of 5% (wt/vol) bovine serum albumin or 10% serum in PBS. Incubate on bench for an hour at room temperature.7. Add primary antibody diluted in 1% (wt/vol) BSA or 1% (wt/vol) serum in PBS to the coverslip and incubate overnight in a humidified chamber at 4 °C.8. Wash the coverslips three times in PBS.9. Add secondary fluorescent-conjugated antibody diluted in 1% (wt/vol) BSA/PBS to the coverslips and incubate on bench at room temperature for an hour. crItIcal step From this point onward, it is necessary to protect the coverslips from light.10. Wash the coverslips three times with PBS.11. On a clean slide, add a drop of ‘antifade’ mounting solution. Gently hold the edge of the coverslip with forceps and tap the edge carefully on a Kimwipe to drain excess liquid.12. Carefully invert the coverslip, cell side down, onto the ‘antifade’ mounting medium on the slide.13. Gently remove excess mounting medium from the sides of the coverslip. crItIcal step Be very gentle and careful and ensure that the coverslip does not move on the slide. This can result in considerable damage to neuronal morphology. (continued)
reagents or environment. We have provided a troubleshooting guide to help identify and overcome some of these culture problems.
Experimental designFor routine studies requiring imaging fixed cells, we plate neurons on 18-mm coverslips in 12-well dishes. Alternatively, four or five coverslips can also be placed in 100-mm Petri dishes for
such studies. When using coverslips, they need to be prepared and coated with poly-l-lysine, as described in Steps 1–11 of the PROCEDURE. For time-lapse imaging, we plate neurons on glass-bottom dishes. For biochemical studies, we plate neurons on tissue culture–treated Petri dishes coated with poly-l-lysine. When doing time-lapse imaging or biochemical studies, start the PROCEDURE at Step 7, as there is no need to prepare coverslips.
Box 3 | Live imaging and FM4-64 labeling to examine synaptic function FM dyes are widely used in cell biology to examine various aspects of exocytosis and endocytosis. FM4-64 is a lipophilic styryl dye that reversely binds to the outer leaflet of the lipid bilayer, but is not membrane permeable. At neuronal synapses, recycling of synaptic vesicles can be induced by depolarization through electrical stimulation or incubation in high K + buffers. During the subsequent endo-cytosis, the dye becomes trapped in endocytic vesicles and can be used as a marker of a functional synapse. Advasep-7 is used as a FM dye scavenger to reduce the background FM dye signal. An alternate protocol (see modifications to make at Steps 2 and 6) takes advantage of FM4-64FX to label recycling vesicles. Cells that take up this dye can be fixed and immunostained without loss of FM4-64FX fluorescence. This offers the advantage of being able to store coverslips for later imaging or co-labeling with antibodies.
FM4-64 labeling of recycling vesicles in live neurons:1. Incubate live cultured neurons in Tyrode’s solution in the cell culture incubator for 10 min.2. Aspirate the solution carefully and incubate with the high-KCl Tyrode’s solution containing 10 mM FM4-64 for 1 min in the cell culture incubator. If you wish to fix cells later, use 10 mM FM4-64X in place of FM4-64. crItIcal step From Step 3 until imaging, use Tyrode’s with CNQX and AP5. CNQX is an AMPA/kainate receptor antagonist and AP-5 is an NMDA receptor antagonist. These inhibitors are used to prevent additional firing to keep FM4-64 in recycled vesicles.3. Aspirate the solution and add fresh Tyrode’s solution; repeat twice.4. Add Tyrode’s solution containing Advasep-7 for 1 min at 37 °C.5. Aspirate the solution and replace it with Tyrode’s solution.6. Wash three times with fresh Tyrode’s solution and incubate in fresh Tyrode’s solution for 10 min in a cell culture incubator. If you wish to fix the cells, fix them using 4% (wt/vol) paraformaldehyde in HBSS without phenol red for 10 min on ice and then wash them twice with HBSS. If desired, immunofluorescence staining and mounting can be performed on fixed cells as described in Box 2.7. Image by using an inverted confocal imaging system after placing the cells in a CO2 and temperature-controlled chamber. FM4-64 can be excited at 510–570 nm (peak at 538 nm) and emission can be collected at 560–750 nm (peak at 654 nm) in a lipid bilayer membrane environment. FM4-64 excitation/emission spectra do not overlap with GFP or Alexa Fluor 488 excitation/emission spectra, and thus can be used for co-labeling with GFP or Alexa Fluor 488.8. Recycling vesicles can be imaged as punctae at synaptic terminals (Fig. 5d).
Box 2 | (continued)14. Allow the coverslip to dry in the dark for an hour.15. Gently seal the sides of the coverslip with clear nail polish. Let the nail polish dry completely before imaging.16. The slides with the coverslips are now ready for imaging. pause poInt The slides can also be stored at − 20 or − 80 °C for several months without substantial loss of fluorescence.
For cultureP0 mouse pups (8–10 pups, either sex; we routinely use C57BL/6 mice) ! cautIon All experiments must be conducted in accordance with the relevant institutional and governmental guidelines and regulations.Medium components, as described in Table 1Cell culture plates (12 well)Culture dishes (60 mm)Sterile microcentrifuge tubes for individual mouse cultureTime-lapse dishes (Mattek, cat. no. P3SG-0-14-C)Petri dishes, bacteriological grade, 100 mm
For transfection with Lipofectamine and nucleofection only (Box 1)Lipofectamine 2000 (Invitrogen, cat. no. 11668-027)EndoFree kit for preparation of DNA (Qiagen, cat. no. 12362)Neurobasal medium (Invitrogen, cat. no. 21103-049)Mouse Nucleofector kit (Lonza Biosciences, cat. nos. VAPG-1001, VPG-1001, VVPG-1001, includes cuvettes)Maintenance medium
For immunostaining only (Box 2)Fixing solution (4% (wt/vol) paraformadehyde/4% (wt/vol) sucrose in 1× PBS, see Reagent Setup for details)Triton X-100Primary antibodies. We have used the following antibodies for immuno-fluorescence: mouse gephyrin-specific (1:2,000; Synaptic Systems), rabbit anti-vGAT (1:1,000; Synaptic Systems), guinea pig vGluT1-specific (1:2,000; Millipore Bioscience Research Reagents), mouse psd95-specific (1:250; clone 6G6-1C9, Thermo Scientific, cat. no. MA1-045), mouse MAP2-specific (1:2,000, clone HM-2, Sigma-Aldrich cat. no. M4403), rabbit Tau-specific (1:1,000, Abcam) and γ-aminobutyric acid (1:4,000, Sigma-Aldrich). We have used the following antibodies for western blotting: mouse β-tubulin–specific (Clone E7, University of Iowa Developmental Studies hybridoma bank) and mouse δ-catenin–specific (BD transduction Labs, cat. no. 611537)DAPI (Sigma-Aldrich, cat. no. D-9564)Serum (species in which the secondary antibody is raised) or BSAFluorophore-conjugated secondary antibodies (We have used Invitrogen’s Alexa Fluor–conjugated secondary antibodies.)PBSProLong Gold antifade (Invitrogen, cat. no. P36934)Glass slides (Fisher Scientific, cat. no. 12-544-3)Nail polish
For FM4-64 labeling of recycling vesicles only (Box 3)FM4-64 dye (Invitrogen, cat. no. T-13320) or FM4-64FX dye (Invitrogen, cat. no. F34653)
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Tyrode’s solution (124 mM NaCl, 3 mM KCl, 2 mM CaCl2, 1 mM MgCl
2,
10 mM HEPES (pH 7.3) and 5 mM d-glucose; see REAGENT SETUP)Tyrode’s solution with CNQX (Sigma-Aldrich, cat. no. C127) and AP5 (dl-2-amino-5-phosphonovaleric acid; Sigma-Aldrich, cat. no. A5282) (see REAGENT SETUP)High-KCl Tyrode’s solution (72 mM NaCl, 50 mM KCl, 2 mM CaCl
2,
1 mM MgCl2, 10 mM HEPES (pH 7.3) and 5 mM d-glucose; see
REAGENT SETUP)Paraformaldehyde (4% (wt/vol)) in Hank’s buffered saline solution (HBSS) without phenol red ! cautIon This is a hazardous material and a possible carcinogen; use it in a chemical safety hood.Sodium hydroxideDMSO
EQUIPMENTTwo Dumont-style (no. 5) forceps—to handle coverslipsOne small scissor, one Dumont-style forceps—to remove brainTwo Dumont-style (no. 5) forceps, one fine scissor—to dissect hippocampusDry sterilizer (Germinator 500; Roboz, cat. no. DS-401, optional)Dissecting microscope with illuminationHemocytometerLight microscopeFluorescent/confocal microscope37 °C water bathFire-polished glass pipette (orifice size 1.5–2 mm)Laminar flow cell culture hoodHigh-temperature dry oven (225–270 °C).Bunsen burnerCell culture incubator—5% CO
2, 95% humidity
Lonza Nucleofector II (Lonza Biosciences, cat. no. AAD-1001S)CO
2 and temperature-controlled chamber mounted on microscope for live
Poly-l-lysine solution Dissolve 0.5 mg ml − 1 of poly-l-lysine in borate buffer; the solution may be stored at − 20 °C for 2–3 weeks.Trypsin Dissolve 2.5% (wt/vol) trypsin in sterile double-distilled water; aliquot and store at − 20 °C. The solution is stable for 2–3 months.DNase Dissolve 1% (wt/vol) DNase in sterile double-distilled water; aliquot and store at − 20 °C. The solution is stable for 2–3 months.Media As described in Table 1.Fixing solution Weigh out 4 g of paraformaldehyde into a beaker. Add 10 ml of 10× PBS and 60 ml of water. Stir continuously and add a drop or two of concentrated sodium hydroxide. This should allow the parafor-maldehyde to go into solution immediately. Add 4 g of sucrose, allow to dissolve completely and adjust the volume of the solution to 100 ml. Alter-natively, this solution can be made by diluting out commercially available concentrated paraformaldehyde solution. The solution can be stored for up to a month at − 20 °C. ! cautIon Paraformaldehyde is a hazardous material and a possible carcinogen; use it in a chemical safety hood. Sodium hydroxide is a caustic chemical; use it in a chemical safety hood.
FM4-64 or FM4-64FX Dissolve in DMSO to make a 10 mM stock solution. Protect from light. The stock solution can be stored at − 20 °C for up to 2 weeks.Tyrode’s solution Mix 124 mM NaCl, 3 mM KCl, 2 mM CaCl
2, 1 mM
MgCl2, 10 mM HEPES (pH 7.3) and 5 mM d-glucose in distilled water.
Filter-sterilize using a 0.2-µm filter. This solution can be stored for up to 4 weeks at 4 °C.Tyrode’s solution with CNQX and AP5 Add 10 mM CNQX and 50 mM AP5 to sterile Tyrode’s solution right before experiments.High-KCl Tyrode’s solution Dissolve 2 mM NaCl, 50 mM KCl, 2 mM CaCl
2,
1 mM MgCl2, 10 mM HEPES (pH 7.3) and 5 mM d-glucose in distilled water.
Filter-sterilize through a 0.2-µm filter. This solution can be stored for up to 4 weeks at 4 °C.EQUIPMENT SETUPPasteur pipette for trituration of cells Take a sterile glass Pasteur pipette and flame the tip carefully to obtain a smooth surface. The opening of the tip should be comparable to that of a 1-ml pipette tip.
proceDurepreparation of coverslips ● tIMInG 2 d, 5 d before culture1| Place coverslips in a heat-resistant glass container with a large surface area. Add concentrated nitric acid (70% (wt/wt)). Swirl coverslips in the acid, cover them well and leave them in acid for at least 12 h. The acid can be reused two or three times, although it becomes discolored by exposure to light. ! cautIon Nitric acid should be handled in a fume hood.
2| Remove the nitric acid and rinse the coverslips four or five times in several volumes of double-distilled water. Afterward, for each wash, allow the coverslips to soak in water for 2 h, before replacing the wash with fresh water. At each wash, swirl the coverslips around carefully.
3| Aspirate water carefully and ensure that there are no droplets of water sticking to the coverslips.
4| Cover the container with aluminum foil and bake in a dry-heat oven at 225–270 °C for 12–16 h. pause poInt Baked coverslips may be stored at room temperature (20–25 °C) for 1 month if covered tightly.
coating of coverslips, time-lapse dishes and petri dishes with poly-l-lysine ● tIMInG 1 h, 2 d before culture5| When working in a sterile laminar hood, use sterile forceps to place individual coverslips in single wells in a 12-well dish.
6| Sterilize by placing the open dishes with the coverslips under ultraviolet light for 30 min. We do not routinely sterilize the time-lapse dishes or Petri dishes, as they come in a sterile package.
7| Add enough poly-l-lysine solution to cover the coverslip and the glass bottom of the time-lapse dishes. Similarly, for Petri dishes, ensure that the volume of poly-l-lysine covers the dish entirely.
8| Wrap the dish in aluminum foil to prevent evaporation and leave it overnight at room temperature. crItIcal step It is important to make sure that the poly-l-lysine does not dry out during incubation. If you are working in the laminar hood, be sure to turn the blower off during the incubation.
Washing of coverslips after coating ● tIMInG 1 h, 1 d before culture9| When working in a sterile laminar hood, aspirate the poly-l-lysine carefully.
10| Add 1 ml of sterile water into each well and aspirate; repeat this step three times. crItIcal step Ensure that the glass surface does not dry out during the washing procedure.
11| Aspirate water, add maintenance medium and leave the coverslips in the tissue culture incubator until you are ready to plate cells, but preferably not for more than 24 h. If it is necessary to leave them for more than 24 h, change the medium to fresh main-tenance medium a few hours before plating the cells.
removal of brains from p0 mouse pups ● tIMInG 3–6 min for each animal! cautIon All experiments must be performed in accordance with relevant institutional and governmental guide-lines and regulations.
12| Sterilize instruments by heating them in a dry sterilizer or washing them with 70% (vol/vol) ethanol. Dry thoroughly if ethanol is used.
13| Prepare 60-mm dishes with dissection medium. If you are culturing from individual mice, prepare microcentrifuge tubes with 1 ml of dissection medium (table 1).
14| Euthanize the mouse pup by decapitation and separate the head from the body (Fig. 2).
15| Place the head on a dish and hold down the sides with forceps.
16| Gently dissect the skin on the top of the head and hold down the skin on either side with the forceps.
17| By using fine scissors, cut open the skull by making an incision at the base of the brain. Separate the two halves of the skull and remove carefully. Take care to not cut through the brain tissue when removing the skull bone.
18| By using forceps, pinch off the brain from the base and transfer it to a 60-mm dish containing dissection/dissociation medium (table 1).
19| To culture neurons from individual pups, put each brain in a separate dish. For pooled cultures, put two or three brains into one 60-mm dish. crItIcal step Ensure that the brains are submerged in the medium, and do not let them dry out at any point.
Dissection of the hippocampus and cortex ● tIMInG 4–6 min for each mouse20| Separate the two halves of the brain by making a sagittal cut along the midline. Discard the cerebellum (Fig. 3).
21| Place the brain such that the outer surface of the hemisphere faces the bottom of the dish.
22| Under a dissecting microscope, gently remove the midbrain and thalamic tissue to leave an intact hemisphere containing the cortex and hippocampus.
23| Turn the tissue over so that the hippocampus faces the bottom of the dish.
24| By using forceps, pin down the anterior cortex, taking care not to damage the hippocampus.
a b c d
e f g h
Figure 2 | Illustration of the technique to remove brains from P0 mice. (a,b) Decapitate pups and remove the head. By using a fine scissors, make a midline incision at the skin surface close to the hindbrain region. (c) Follow the incision to the extreme rostral region. (d) Make a small incision at the base of the skull and follow along the midline. (e) Separate the two halves of the skull to reveal the brain. (f) Gently place the forceps underneath the brain and separate it from the underlying tissue. (g,h) Gently remove the intact brain (g) and quickly place it into dissection medium (h). It is important to be extremely fast and careful to avoid contamination. All animal procedures were carried out in accordance with approved protocols from the Institutional Animal Care and Use Committees at UNMC and UCSF.
25| Use another pair of forceps to pick and grab the meninges carefully and gently peel them off, ideally as a single piece. Check for the remaining pieces of meninges and remove them completely. crItIcal step At this stage during mouse development, the meninges can be sticky and difficult to remove, compared with the embryonic stage. However, it is important to ensure that the meninges are completely removed, so that they do not contribute any non-neuronal cells to the culture.
26| Orient the tissue so that the hippocampus is on the top. At this stage, the hippocampus can be identified by its C-shaped structure and opacity, which differ from the neighboring cortical tissue. The cortex can also be dissected out and processed similarly for cortical cultures.
27| By using forceps or fine scissors, separate out the hippocampus carefully. Collect it in a 60-mm dish containing fresh dissection/dissociation medium. For culture of individual hippocampi or cortices, collect each pair of hippocampi or cortices in a 1.5-ml Eppendorf tube containing 1 ml of dissection/dissociation medium. crItIcal step It is important that the dissection be done as quickly as possible to ensure cell viability and health.
cell dissociation and plating ● tIMInG ~1–1.5 h28| Place all the hippocampi or cortices in a 15-ml centrifuge tube for a pooled mouse culture. For culturing individual mouse cultures, leave hippocampi or cortices in Eppendorf tubes. Wait until all tissues settle to the bottom and remove all but 5–10% of the medium.
29| Add 10 ml of fresh dissection/dissociation medium, wait for the tissue to settle to the bottom of the tube, aspirate and repeat twice. For tissue in Eppendorf tubes, wash it twice with 1 ml each of dissection medium. crItIcal step It is important to be very gentle with the washing, so as to not damage the hippocampi or break them apart. Smaller pieces of tissue are more difficult to handle during the washing procedure.
30| Resuspend the tissue in 4.5 ml of fresh dissection medium and add 0.5 ml of trypsin solution; swirl the tube around gently to mix. For tissue in individual Eppendorf tubes, scale the volumes down to 450 and 50 µl, respectively.
31| Incubate at 37 °C in a water bath for 20 min. crItIcal step It is important to ensure that this incubation does not proceed for longer than 20 min.
32| Add 0.5 ml of DNase solution and incubate in a laminar flow hood at room temperature for 5 min. For tissue in Eppendorf tubes, add 20 µl of DNase and incubate for 2–5 min.
33| Aspirate the medium and wash the tissue twice gently with 10 ml of fresh dissection medium for each wash. For tissue in Eppendorf tubes, use 1 ml of dissection medium for each wash. crItIcal step Be extremely gentle while washing the hippocampi and cortices because their tissue structures have now been loosened by trypsin incubation.
a c
d e f
bFigure 3 | Steps for dissection of the hippocampus from the intact brain. (a) Place the brain dorsal side up in dissection medium. (b) Separate the hindbrain region and make a sagittal incision to separate the two hemispheres. (c) Place each hemisphere’s cortex side down and remove any noncortical forebrain tissue. (d) Hold the hemisphere in place using forceps; use caution so as not to damage the hippocampus. (e) Remove the meningeal tissue with another pair of forceps. Dissect out the hippocampus (f) and the cortex. All animal procedures were carried out in accordance with approved protocols from the Institutional Animal Care and Use Committees at UNMC and UCSF.
34| Wash twice, each with 10 ml (or 1 ml, for tissue in Eppendorf tubes) of plating medium. crItIcal step Use temperature-equilibrated plating medium from this point forward, as the sera will inactivate the trypsin.
35| Resuspend hippocampi or cortices in 2.5 ml (for tissue from six to ten animals) or 0.5 ml (for tissue from individual animals in Eppendorf tubes) of plating medium.
36| Place the lid of a 100-mm bacteriological-grade Petri dish face down on the floor of the hood. Place the base of the dish on the top of the inverted lid such that the base is at about 30° resting both on the top of the lid and on the floor of the hood. This is to ensure the use of minimal surface area for dissociating the hippocampi. Bacteriological-grade dishes do not support the adherence of neurons to the dish and hence result in greater yield.
37| Carefully place the tissue and the solution in the base of the 100-mm dish such that it occupies the volume at the bottom of the dish (Fig. 4).
38| By using a fire-polished glass pipette (Fig. 4), carefully triturate the tissue eight to ten times to dissociate the cells gently and obtain a homogenous cell suspension. At this point, there should be no or minimal chunks of tissue. crItIcal step The trituration of cells should be done slowly and carefully to minimize damage to cells. It is best to avoid any bubbling during the procedure and to try to keep the pipette within the cell suspension at all times. The tip diameter of the glass pipette is crucial. Too small an orifice can result in cell damage, whereas too large an orifice can cause inadequate trituration. It is also necessary to be careful with the length of trituration. Too much can damage cells and too little can cause large chunks of tissue to be retained, with either case resulting in a loss of yield. Use a cell strainer if it is challenging to remove tissue chunks, especially for preparing cortical cultures or hippocampal cultures from more than ten mice.? trouBlesHootInG
39| Take a 10-µl aliquot of the dissociated cells to estimate viable cell density on a hemocytometer. Viable cells can be identified by their glossy appearance, whereas dead cells appear dark. Typical yields are 400,000–600,000 hippocampal cells and 600,000–800,000 cortical cells per animal. An optional alternative is to use trypan blue staining to estimate cell viability. Add 10 µl of cells to 40 µl of maintenance medium with 50 µl of trypan blue. Determine the density of cells on a hemocytometer. In this case, viable cells are colorless, whereas dead and damaged cells are blue. If you wish to perform a high-efficiency transfection, proceed to Box 2.? trouBlesHootInG
40| At this point, the desired number of cells can be plated out or used for nucleofection (Box 2). We routinely add 90,000 cells to an 18-mm coverslip in a 12-well dish (65 cells per mm2) and 45,000 cells per time-lapse dish that contains maintenance medium (65 cells per mm2). Gently mix to ensure even plating on the coverslip. Cells that do not settle on the coverslip will not survive. For biochemical studies, we plate 500,000 to 1 million cells on a 60-mm-diameter Petri dish (44–88 cells per mm2) and 1–2 million cells on a 100-mm-diameter Petri dish (32–64 cells per mm2). crItIcal step It is important to plate the neurons, particularly the cortical neurons, quickly to avoid any clumping. If clumping is an issue, dilute the cell suspension with plating medium and triturate gently again, as described in Step 38.
41| Incubate in the cell culture incubator at 37 °C for 2–6 h.
42| Four hours after plating cells, examine under the microscope to ensure that the cells have settled on the substrate.? trouBlesHootInG
a b c
UnpolishedPolished
Pipette tip
Figure 4 | Trituration of isolated hippocampi/cortex. (a) Comparison of unpolished and polished tips of glass Pasteur pipettes. A plastic 1,000-µl pipette tip is shown for comparison. Note that the polished tip has smooth edges and an orifice size comparable to that of the pipette tip. (b) Intact dissected hippocampi in dissection medium at the start of trypsinization. (c) Setup of Petri dishes for triturating cells. All animal procedures were carried out in accordance with approved protocols from the Institutional Animal Care and Use Committees at UNMC and UCSF.
43| Gently aspirate the medium from each well and add 1.5 ml of fresh maintenance medium warmed to 37 °C to each well in the 12-well dish, 1 ml to each time-lapse dish and 10 ml in the 100-mm Petri dish. crItIcal step It is necessary to ensure that the medium has been warmed to 37 °C and that the manipulation is gentle so as not to damage the cells. We prefer to aspirate the medium using a pipette and not a vacuum-based method. Avoid aspiration or addition of medium directly above cells; instead, allow the medium to trickle down gently through the side wall of the plate or well.
44| Incubate in a cell culture incubator at 37 °C.
45| If desired, 2 d after plating, add cytosine arabinoside (araC; 1-b-d-arabinofuranosylcytosine) to a final concentration of 1–5 µM to inhibit the proliferation of dividing non-neuronal cells. Several investigators continue to maintain araC in the medium through the life of the culture. However, in our hands, we have noticed that occasionally the addition of the araC causes the death of neurons. To avoid this, we replace the araC-containing medium with fresh maintenance medium 24–48 h after adding the araC.
Maintenance of neurons ● tIMInG up to 28 d46| Twice a week, aspirate half the medium from each well/time-lapse dish and replace it with fresh maintenance medium warmed to 37 °C. These neurons can be maintained in culture for up to DIV 28. The neurons may be used for low-efficiency transfections (Box 1), immunostaining (Box 2) or live imaging (Box 3) anytime during this period (DIV 0–28). The cultures can be used for examining synaptic function by FM4-64 dye uptake between days 11 and 28 (Box 3). crItIcal step It is important to not replace the entire medium. Neurons secrete factors that promote growth and survival.? trouBlesHootInG
? trouBlesHootInGTroubleshooting advice can be found in table 2.
taBle 2 | Troubleshooting table.
step problem possible reason solution
38 Difficulty in triturating hippocampi
Insufficient trypsinization Ensure the correct concentration and incubation time of trypsin, use fresh trypsin
DNA from damaged cells makes cell suspension viscous
Consider using DNase at the right concentration
Insufficient incubation with DNase Increase incubation time with DNase
Opening of Pasteur pipette tip for trituration is too large
Ensure that the opening of the Pasteur pipette tip is adequate
39 Excessive number of dead cells (more than 25% at the time of counting)
Opening of Pasteur pipette orifice is too small or not smooth enough
Ensure that the opening of the Pasteur pipette is the correct size and smooth
Harsh trituration Triturate the cells gently, ensuring little or no bubbling of liquid
Insufficient trypsin activity, strain on cells during trituration
Ensure the right concentration of trypsin or replace the batch of trypsin
Wrong pH of the dissection buffer Check the pH of the dissection buffer
Extended time period between remov-ing tissue and trituration of tissue
Reduce the time for the entire procedure by practicing techniques
Low yield of cells Clumping of cells Ensure adequate and gentle trituration of cells
39 Cells adhere to Petri dish during the trituration process
Use bacteriological-grade dishes that are not tissue culture coated
Insufficient starting material Ensure enough number of hippocampi used
42 Neurons do not adhere to coverslips when plated
Incorrect coverslip glass Ensure that the right coverslip is being used; neurons are very sensitive to the glass surface
Insufficient etching of coverslips Follow the nitric acid soaking and baking protocol; bake coverslips in small batches to ensure adequate etching
Insufficient volume of poly-l-lysine on coverslip
Use enough poly-l-lysine to coat the coverslip entirely
Poly-l-lysine–coated coverslips are stored for too long
Discard and coat fresh coverslips
Incorrect composition of borate buffer
Check the concentration and pH of borate buffer
Poor quality poly-l-lysine Try a different batch of poly-l-lysine
46 Contamination Nonsterile conditions Sterilize tools by autoclaving or using a dry sterilizer (e.g., Germinator 500 from Cell Point Scientific)
Clean working surface areas with 70% (vol/vol) ethanol
Observe sterile working conditions
Separate tools for culture from tools for other experiments
Cells do not appear viable or healthy 4–5 d after plating
Inappropriate medium composition Use fresh components for all media. Do not store the maintenance medium for more than 10–12 d at 4 °C. Do not freeze thaw serum for plating medium more than once. Do not warm aliquots of maintenance medium more than two times
Inappropriate coating of coverslips or coated coverslips were too old
Ensure coverslips are coated with the correct concentra-tion of poly-l-lysine. Do not store coated coverslips at 4 °C for more than a week
Medium change is inappropriate Replace half the medium with fresh medium warmed to 37 °C every 3–4 d
Medium changing procedure takes too long
In changing the medium, work with cultures in the hood for as short a period of time as possible to avoid creat-ing an alkaline medium and killing cells. When changing medium from multiple dishes, it is better to only take out and feed one dish at a time
Box 1 Variability in transfection—low-efficiency transfection
Neurons are not healthy Optimize neuronal culture to obtain healthy cells that can be maintained in culture
Transfection reagents are not fresh or have changed
● tIMInGPreparation of coverslips should be started 5 d before the day of culturing, but it requires only about an hour each day. Preparation of the hippocampal cultures will take 4–8 h, depending on the number of mice. Subsequent maintenance of the cultures requires an hour twice a week to warm and replace the medium, and an additional hour for performing low-efficiency transfections.
antIcIpateD resultsThe culture of hippocampal and cortical pyramidal neurons is widely used and well characterized. Neurons in culture undergo distinct stages of development, starting with extension of lamellopodia, followed by axon and dendrite specification and extension and synapse formation and maturation. All of these stages can be distinguished in these cultures. Note, however, that the cell population is not completely synchronized and hence all the neurons may not simultaneously be at the exact same stage of development2. Consequently, it is possible to detect neurons with different levels of MAP2 staining (supplementary Fig. 1a). Moreover, we detect some DAPI-positive
taBle 2 | Troubleshooting table (continued).
step problem possible reason solution
Box 1 DNA concentration is not right or DNA quality is not good
Measure DNA concentration and absorbance ratio at 260/280 nm (A260/A280) ratio again. These should be above 1 mg ml − 1 and ~1.8, respectively. Prepare fresh DNA
Too many or too few cells are transfected
Adjust the amount of DNA for transfection
Lipid-DNA complex incubation time is either too long or too short
Make sure the incubation time is between 20–30 min
Low-quality DNA Use high-quality DNA. We have had success with using the EndoFree maxi prep kit from Qiagen (cat. no. 12362)
Variability in transfection—high-efficiency transfection
Inappropriate solution for electroporation
Use the manufacturer’s solution for electroporation
Too few cells are transfected Adjust the amount of DNA for transfection
Low-quality DNA Use high-quality endotoxin-free DNA
DNA concentration is not right or DNA quality is not good
Measure DNA concentration and A260/A280 ratio again. These should be above 1 mg ml − 1 and ~1.8, respectively. Prepare fresh DNA
Too many cells die after nucleofection Ensure that the right number of cells and program are being used. Leave cells in the nucleofection solution for the minimum time possible
MAP2 Tau Merge
a
b
DIV 0
DIV 5
DIV 1 DIV 2
DIV 10 DIV 14
Figure 5 | Images of cultured hippocampal neurons. (a) DIC images of neurons at different stages during culture—DIV 0, 1, 2, 5, 10 and 14. Note how neuronal morphology and arborization develop over time. Scale bar, 10 µm. (b) Immunostaining of DIV 7 neurons with axonal (Tau, green) and dendritic (MAP2, red) markers. Note that axons and dendrites can be clearly distinguished by Tau-specific and MAP2-specific labeling and morphology. Scale bar, 20 µm.
structures that are smaller than neu-ronal nuclei. We suspect that these represent non-neuronal dividing cells that are undergoing cell death after araC treatment or fragmented cell debris from the preparation proce-dure. Note that these structures do not usually have a distinct cell body associated with them (supplementary Fig. 1b). Hence, DAPI staining may not be a reliable indicator of total cell number if not counted carefully, and differential interference contrast (DIC) imaging might be a more reliable indicator of total cell counts. In these cultures, we detect about 6–8% astrocytes, as identified by glial fibrillary acidic protein (GFAP) reactivity (supplementary Fig. 2), and about 20% of the neuron population comprises inhibitory neurons, as identified by immuno-staining to γ-aminobutyric acid (GABA) (supplementary Fig. 3).
Mouse postnatal neurons cultured as described here can be successfully maintained in culture for several days. These neurons extend neurites that have characteristic morphologies (Fig. 5a) and express appropriate dendritic (MAP2) and axonal (Tau) mark-ers (Fig. 5b). These neurons are amenable to transfections. In neurons transfected with a GFP-expressing vector, it is possible to distinguish axons and dendrites by virtue of their morphology after about 10–12 d of culture (Fig. 6a; axons are thin and lack spines; dendrites are thicker and mature dendrites typically contain spines). Neurons develop excitatory synapses as indicated by staining for psd95 (excitatory postsynaptic marker) and vGlut1 (excitatory presynaptic marker) and inhibitory synapses as indicated by gephyrin (inhibitory postsynaptic marker) and vGAT (inhibitory presynaptic marker) labeling (Fig. 6b). Synaptic function can be assessed using the FM4-64 dye uptake assay (Fig. 7).
In neurons that have been transfected with the high-efficiency nucleofection with vector and shRNA-expressing constructs, it is possible to detect the knockdown of the protein of interest by western blot analysis (supplementary Fig. 4). Moreover, cells are quite amenable to time-lapse imaging. Figure 8 shows an example of a dendritic segment with spines in a GFP-expressing neuron imaged over time. Such imaging allows for clear examination of spine morphology over time. It is also possible to couple such studies with co-transfections with fluorescently labeled synaptic markers, such as psd-95, to simulta-neously examine spine and synapse dynamics over time.
The ability to culture mouse postnatal neurons offers the potential to investigate multiple aspects of neuronal structure and function. However, the great utility of cultured neurons is in the ability to be manipulated with low- and high-efficiency transfections. Transfected neurons are ideal for studying various aspects of neuronal structure and function by imaging or by electrophysiology. In addition, these cells can be further manipulated by overexpression or knockdown of proteins. By using genet-ics to manipulate gene expression and high-efficiency transfections to mark the populations, one can generate mixed cultures of mouse neurons from
Figure 6 | Morphology and immunostaining of neurons for synaptic markers. (a) Confocal microscope images of a DIV 17 neuron expressing GFP. The dendrites and axons can be distinguished clearly by their morphology. Note the presence of spines on the dendrites and the thin morphology of the axon. Scale bar, 20 µm. (b) Immunostaining of DIV 14 GFP-transfected neurons with antibodies to excitatory and inhibitory pre- and postsynaptic markers. The majority of inhibitory synapses (presynaptic—vGAT, blue; postsynaptic—Gephyrin, red) form on dendritic shafts (*, left image), whereas the majority of excitatory synapses (presynaptic—vGlut1, blue; postsynaptic—psd95, red) form on spine heads (*, right image) and can be clearly distinguished. Scale bars, 5 µm.
*
FM4-64 MergeGFP
*
Figure 7 | Synaptic functional assay on cultured neurons. A GFP-transfected neuron at DIV 17 subjected to FM4-64 dye uptake protocol to examine functional synapses (*). Scale bar, 5 µm.
Note: Supplementary information is available in the online version of the paper.
acknoWleDGMents We thank the past and present members of the Reichardt laboratory for their support and insight. This work was supported by US National Institutes of Health Grant F32-MH079661 (G.M.J.B.), the Simons Foundation (L.F.R.) and start-up funds from the Munroe-Meyer Institute, University of Nebraska Medical Center (J.A.).
autHor contrIButIons G.M.J.B., S.-H.L. and J.A. designed experiments. G.M.J.B., S.-H.L., D.S., Y.Y., Y.-G.N. and J.A. performed the experiments. G.M.J.B., S.-H.L., D.S., Y.Y. and J.A. collected, analyzed and interpreted data. J.A. and L.F.R. supervised the experiments. G.M.J.B., S.-H.L., L.F.R. and J.A. wrote the manuscript.
coMpetInG FInancIal Interests The authors declare competing financial interests: details are available in the online version of the paper.
Published online at http://www.nature.com/doifinder/10.1038/nprot.2012.099. Reprints and permissions information is available online at http://www.nature.com/reprints/index.html.
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Figure 8 | Neurons in culture are amenable to time-lapse imaging. Time-lapse imaging of two dendritic shafts with associated spines in GFP-transfected DIV 14 neurons. Note that the dendritic shaft shows no evidence of damage even upon prolonged imaging. Note how some spines are relatively stable (#), whereas others show alterations in morphology (*) over time. Scale bar, 2 µm.
two different strains of mice to dissect their abilities to form functional connections with each other24. In addition, transfec-tion allows for temporal control of gene deletion; for instance, cultures from floxed mice can be transfected with a plasmid encoding Cre recombinase to abolish the expression of the gene of interest at various time points. These cultures could also be successfully used for routine biochemical studies and proteomics25. With the use of high-efficiency transfections, these cells are amenable to biochemical studies of a large cohort of neurons.
In summary, primary mouse cultures generated successfully following the protocol described can be used for a variety of cell biological and biochemical studies. These cultures, prepared with a modest level of difficulty and care, are quite robust. They can offer great insights into neuronal cellular architecture and function.
