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Title:

Isolation, purification and expansion of myelination-competent neonatal mouse Schwann Cells

Running Title:

Myelination-competent mouse Schwann Cells

Authors:

Henrika Honkanen1, Outi Lahti1, Marja Nissinen2, Riina M. Myllylä2, Satu Päiväläinen1, Maria H. Alanne2, Sirkku Peltonen3, Juha Peltonen2 & Anthony M. Heape1

Affiliations:

1 Myelin Group, Department of Anatomy & Cell Biology, University of Oulu, Aapistie 7A, 90014 Oulu, Finland.

2 NF1 Research Team, Department of Anatomy & Cell Biology, University of Oulu, Aapistie 7A, 90014 Oulu, Finland.

3 Cell-Cell Interaction Group, Department of Dermatology, University of Turku, Savitehtaankatu 1, 20521 Turku, Finland.

Corresponding author:

Doc. Anthony M. Heape,

The Myelin Group,

Department of Anatomy & Cell Biology,

P.O. Box 5000 (Aapistie 7A),

90014 University of Oulu,

Finland.

E-mail:

Phone: (+358)-8-5375197

Fax: (+358)-8-5375172

Funding

This work was funded by the NeoBio Research Program of the Finnish National Technology Agency (Tekes), and a research grant from the Academy of Finland. AMH is a Research Fellow of the Academy of Finland.


Abstract

Most studies of PNS myelination using culture models are currently performed using dorsal root ganglion neurons (DRGN) and Schwann cells (SC) pre-purified from the rat. However, the potential of the model is severely compromised by the lack of rat myelin mutants, and the published protocols work very poorly with cells from mice, for which numerous myelin mutants are available. Here, we describe the isolation, purification and expansion of wild-type, myelination-competent SCs from the sciatic nerves of 4-day-old mouse pups.

The protocol consistently provides ~95 %-pure mouse SC yields of 1.9 - 3.3 x 106 cells from the sciatic nerves of 12 to 15 four-day-old mouse pups, within 14 - 20 days of the isolation of the nerves. The proliferation rate of the cultured mouse SCs, expressed as “fold growth/week”, ranges from 2.7 to 4.30 under optimal conditions. Mouse SC proliferation usually ceases within 4 weeks, when the cells become quiescent. Growth is re-induced by the presence of sensory neurons; neuregulin is not sufficient for this effect. The SCs isolated by this protocol maintain their ability to form compact myelin in culture, as judged by the segregated expression patterns of early (myelin-associated glycoprotein) and late (myelin basic protein) markers of myelination in a three-dimensional DRGN/SC coculture model. The SC yields are sufficient to perform 100 to 150 individual myelinating DRGN/SC coculture assays.

Key words

Rodent; Peripheral nervous system; Glial cell; Growth; Myelin; Culture; Method.


Introduction

Schwann cells (SC) are the myelin-forming glial cells of the peripheral nervous system (PNS). As they are the direct and indirect targets of numerous hereditary and acquired peripheral myelin diseases affecting Man, there is strong interest in the research community to understand their biology and pathology. This strong interest has, over the course of the last few decades, led to the establishment of numerous animal models of human SC-related diseases and culture systems that have proved invaluable in studies of cellular and molecular processes related to SC biology and, in particular, to myelin formation, maintenance and disease.

Most studies using myelinating culture models are currently performed using dorsal root ganglion neurons (DRGN) and SCs pre-purified from the rat (Brockes et al. 1979; Kleitman et al. 1997). This model has been employed successfully by many research groups to study various aspects of PNS myelination. However, the potential of the model is severely compromised by the lack of rat myelin mutants. A notable exception is the rat model of human Charcot-Marie-Tooth type 1A disease (Serada et al. 1996; Nobbio et al. 2006).

In contrast, there are several spontaneous mouse myelin mutants, and an ever-growing number of bio-engineered mutants. If employed in a DRGN/SC coculture model, these mutants would be highly valuable for elucidating cell type-specific biomolecular and cellular events taking place during normal and pathological myelination. In particular, when used with mouse DRGNs and SCs this model would provide the unique opportunity of performing mixed phenotype/genotype cocultures, exploiting the abundance of mice with spontaneous or bio-engineered mutations expressed by their SCs, or sensory neurons.

Unfortunately, the published protocols employed for the rat myelinating coculture model with prepurified SCs and neurons work very poorly with mouse cells, a situation that is largely, but not exclusively, related to the difficulties of isolating and purifying sufficient quantities of myelination-competent mouse SCs. Consequently, for culture-based studies of mouse PNS myelination, researchers tend to resort to the somewhat less refined, and cellularly heterogeneous (but see Kim et al. 1997) dorsal root ganglion (DRG) explant culture model, in which the neurons and the SCs are necessarily derived from the same animal (for example; Liu et al. 2005).

Here, we describe a protocol for the isolation, purification and expansion of myelination-competent SCs from 4-day-old mouse pups, in quantities sufficient for carrying out medium-to-large series of myelinating cocultures (i.e. in excess of 100 cocultures).


Materials and Methods

Culture media

The compositions of the culture media employed in this study are as follows:

Basic growth medium: High-glucose Dulbecco’s modified essential medium (HG-DMEM; Sigma), containing 10 % inactivated horse serum (Gibco), 4 mM L-glutamine (Sigma), 100 units (u)/ml penicillin/streptomycin (Sigma), 2 ng/ml human heregulin-β1 (Sigma), and 0.5 μM forskolin (Sigma).

SC Growth Medium: HG-DMEM, containing 10 % inactivated horse serum, 4 mM L-glutamine, 100 u/ml penicillin/streptomycin, 2 ng/ml human heregulin-β1, 0.5 μM forskolin, 10 ng/ml human basic fibroblast growth factor (bFGF; Sigma), and 20 μg/ml bovine pituitary extract (PE; Sigma).

HMEM: HG-DMEM, containing 20 mM Hepes, 10 % inactivated horse serum, 4 mM L-glutamine, and 100 u/ml penicillin/streptomycin.

Complement-mediated Cytolysis Medium: HMEM containing 4 µg/ml anti-mouse CD90 anti-Thy-1.2 (Serotec).

DRG Growth Medium: Minimum essential medium with Earl’s salts (EMEM; Sigma), containing 10 % inactivated horse serum, 4 g/L glucose and 50 ng/mL nerve growth factor (NGF).

Differentiation Medium: HG-DMEM/Hams F12 (Gibco) (1:1, vol/vol), containing 1 % N2-supplement (Gibco), and 50 ng/ml nerve growth factor (NGF; R&D Systems).

Myelination Medium: EMEM, containing 5 % inactivated horse serum (Gibco), 0.4 % D-(+)-glucose (Sigma), 4 mM L-glutamine, 50 ng/ml NGF, 0.5 μM forskolin, 20 μg/mL PE, 0.25% N2-supplement, and 50 µg/ml ascorbic acid (Sigma).

Preparation of poly-L-lysine-, collagen-, and laminin-coated culture surfaces

All coating solutions (0.25 mg/ml collagen type 1 in 0.1 N acetic acid; 10 µg/ml laminin from Engelbreth-Holm-Swarm murine sarcoma basement membrane; and 10 µg/ml poly-L-lysine) were prepared according to the instructions of the manufacturer (Sigma).

Collagen type 1 was applied to sterile 35-mm diameter (Ø) tissue culture dishes (Greiner Bio-One) for 3 h at +37 °C, and the surface was washed 3 times with sterile H2O.

Laminin was applied to 35-mm (Ø) tissue culture dishes for 2 h at +37 °C, and the surface was washed 3 times with sterile H2O.

Poly-L-lysine was applied to 35-, or 60-mm (Ø) tissue culture dishes, or 13-mm (Ø) glass coverslips, for 45 min at room temperature, and the surface was washed twice with sterile H2O.

Animals and isolation of nerve tissue

The use of the mice and the protocols employed for the isolation of the tissues were approved by the Animal Care and Use Committee of the University of Oulu, Finland (permit No. 035/04).

The protocol employed for the isolation and growth of mouse SCs is based on the method of Brockes et al. (1979), as modified by Kleitman et al. (1997), for the rat SCs.

Four-day-old CD-1 mouse pups (12 - 15 pups/preparation, unless stated otherwise), raised in the Laboratory Animal Centre (University of Oulu, Finland), are sacrificed by CO2 exposure followed by decapitation, and cleaned by brief submersion in 70 % ethanol. Both sciatic nerves are exposed, dissected out under a binocular microscope, and placed in a sterile culture dish (3-cm Ø, Greiner Bio-One) containing ice-cold phosphate-buffered saline (PBS) without CaCl2 or MgCl2 (Sigma), where they are maintained until the nerves have been harvested from all of the mice. The nerves are then carefully stripped of connective tissue with fine forceps and transferred to a new culture dish containing 3.0 ml ice-cold PBS. The cleaned nerves are shredded with forceps until they resemble cotton balls. The sciatic nerves are kept on ice for not more than 1 hour.

Dissociation and plating of nerve fragments

The shredded nerves, together with the PBS (3 ml), are transferred from the culture dish to a 15-mL plastic Falcon tube containing 0.5 ml 2.5 % trypsin (Sigma), 0.5 ml 10 mg/ml collagenase A (Roche), and 3 ml PBS. The culture dish is rinsed with 3 ml of PBS, and the rinsate is added to the Falcon tube containing the nerve/enzyme mixture (total volume 10 mL; final enzyme concentrations: 0.125 % trypsin and 0.05 % collagenase A). The enzymatic digestion is allowed to proceed for 30 min at +37 °C, mixing occasionally, after which the mixture is centrifuged at 190 x g for 5 min in a benchtop centrifuge (Centra CL2 Thermo IEC). The supernatant is removed, taking care not to aspirate the fluffy nerve fragments, and the residual enzymes are flushed from the cellular pellet by 3 consecutive washes with 7 ml of high-glucose DMEM (Sigma) containing 10 % horse serum (Gibco) and 5-min centrifugations at 190 x g. The cells are resuspended in 2 ml of Basic Growth Medium (see above), plated on a poly-L-lysine-coated 60-mm plastic tissue culture dish, and placed in a cell culture incubator (Forma Scientific) at + 37 °C, with a 5 % CO2 humidified atmosphere. Two days later, the Basic Growth Medium, including any material that has not already adhered to the culture dish, is transferred to a fresh poly-L-lysine-coated 35-mm tissue culture dish. Two mL of fresh Basic Growth Medium are added to the original culture dish containing the attached cells, and both dishes are replaced in the cell culture incubator for a further 2 days.

Purification of Schwann cells – complement-mediated cytolysis

Complement-mediated cytolysis of the contaminating fibroblasts with mouse anti-mouse CD90 anti-Thy-1.2 antibody (Dong et al. 1999) is performed 96 h (4 days) after the enzymatic dissociation and plating of the nerve fragments. The Basic Growth Medium is removed from the cultures and the attached cells are rinsed once with Ca2+- and Mg2+-free Hank’s balanced salt solution (HBSS) (Gibco) in 20 mM Hepes Buffer (Cambrex), and once with HMEM (see above), before adding 1 ml of the Complement-mediated Cytolysis Medium (see above). After incubation for 15 min at +37 °C, 200 µl rabbit HLA-ABC complement sera (Sigma) is added and the incubation is continued for another 2 h at +37 °C. Cytolysis is terminated by rinsing the cells twice with Ca2+- and Mg2+-free HBSS in 20 mM Hepes Buffer.

The complement-mediated cytolysis is repeated before the first passage (see below) if fibroblasts are still visible in the cultures. Fibroblasts are routinely identified by phase-contrast microscopy as flattened polymorphic cells with large, round, non-luminescent nuclei.

Expansion of Schwann cells

After the complement-mediated cytolysis of the fibroblasts, 2 mL SC Growth Medium (see above) are added to each culture and the plated cells are maintained in a cell culture incubator (Forma Scientific) at +37 °C, with a 5 % CO2 humidified atmosphere, changing the medium every 2 days. The cells are passaged when the dishes are ~80 % confluent, i.e. after 7-12 days in culture, depending on the initial SC population size and the cell proliferation rate.

For passaging, the cells are rinsed two times with Ca2+- and Mg2+-free HBSS in 20 mM Hepes buffer, followed by a brief incubation (less than 2 minutes) with ~0.5 mL 1 x trypsin versene (Sigma). Trypsination is stopped by the addition of 1-2 mL of horse serum, followed by rinsing with high-glucose DMEM containing 10 % horse serum. After centrifugation (190 x g for 5 min), the cells are again rinsed with 5-10 mL high-glucose DMEM containing 10 % horse serum, recentrifuged and suspended in 1.0 mL of SC Growth Medium, and counted in a Bürker-Türk cell counting chamber.

The SCs, resuspended in SC Growth Medium, are replated on poly-L-lysine-coated culture dishes at a density of 5 - 6 x 104 cells/ 35-mm dish (with 1 mL medium), or 1.5 – 1.9 x 105 cells/60-mm dish (with 2 mL medium), and grown at +37 °C, with a 5 % CO2 humidified atmosphere, changing the medium every 2 days, until the cultures reach ~80 % confluence. The cells are then trypsinated and counted, as described above.

For frozen storage, the trypsinated cells are centrifuged, resuspended in foetal bovine serum (FBS; Gibco) containing 7% dimethylsulfoxide (DMSO; Sigma), and stored in liquid nitrogen.

Characterization of the Schwann cell cultures

Phase-contrast microscopy. Routinely, SCs are identified as elongated bi-, or tripolar cells, with an oval luminescent soma. Fibroblasts appear as flattened, polymorphic, non-luminescent cells with a large, round nucleus.

Immunofluorescence microscopy. SCs were identified by triple fluorescence microscopy as follows. Cells harvested after one complement-mediated cytolysis and two sub-culture/trypsination cycles, performed as described above, were plated in SC Growth Medium on poly-L-lysine-coated glass coverslips (Ø 13.0 mm) in 35-mm culture dishes, at a density of 20000 cells/coverslip, and cultured for 48 h, as described above. The culture medium was then removed and, after rinsing the coverslips with PBS, the cells were fixed with 2.5 % para-formaldehyde for 10 min at room temperature, permeabilized with ice-cold methanol for 6 min, and again rinsed with PBS. Non-specific binding surfaces were blocked by incubation for 15 min, at room temperature, with 1 % bovine serum albumin (BSA) in PBS.

Primary antibodies (Abcam): mouse monoclonal anti-S100β antibody and rabbit polyclonal anti-Glial Fibrillary Acidic Protein (GFAP), were mixed and diluted in PBS containing 1 % BSA, to final dilutions of 1:200 and 1:1000, respectively.

Secondary antibodies (Molecular Probes): Alexa Fluor® 488-conjugated goat anti-mouse IgG (H+L) and Alexa Fluor® 568-conjugated goat anti-rabbit IgG (H+L), were mixed and diluted in PBS containing 1 % BSA, both to final dilutions of 1:100.

The fixed cells, on the coverslips, were incubated with the primary antibody mixture for 45 min at +4 °C, washed 3 x 5 min with PBS containing 1 % BSA, incubated with the secondary antibody mixture for 30 min at + 4 °C, and washed once with PBS. Finally, the cells were incubated with 0.2 µg/ml Hoechst nucleus dye (Sigma) in PBS for 10 min, washed once with PBS and 3 times with H2O, and then mounted on a glass microscope slide. The slides were observed under a Nikon Eclipse E600 fluorescence microscope, photographed using a QImaging MicroPublisher 5.0 RTV camera, and analyzed with QCapture PRO Software.

The total number of cells (Hoechst-positive), the number of SCs (S100β-positive, GFAP-positive and Hoechst-positive), and the number of fibroblasts (Hoechst-positive, but S100β- and GFAP-negative), were counted in 10 random fields (30-50 cells/field). The abundance of each cell type is expressed relative to the total number of Hoechst-positive cells in each field.