Real-Time Monitoring of Human Glioma Cell Migration on Dorsal Root Ganglion Axon-Oligodendrocyte Co-Cultures

Cancer Research

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Summary

Here we present an ex-vivo mixed monolayer culture system for the study of human glioma cell (hGC) migration in real-time. This model provides the ability to observe interactions between hGCs and both myelinated and non-myelinated axons within a compartmentalized chamber.

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Zepecki, J. P., Snyder, K. M., Tapinos, N. Real-Time Monitoring of Human Glioma Cell Migration on Dorsal Root Ganglion Axon-Oligodendrocyte Co-Cultures. J. Vis. Exp. (154), e59744, doi:10.3791/59744 (2019).

Abstract

Glioblastoma is one of the most aggressive human cancers due to extensive cellular heterogeneity and the migration properties of hGCs. In order to better understand the molecular mechanisms underlying glioma cell migration, an ability to study the interaction between hGCs and axons within the tumor microenvironment is essential. In order to model this cellular interaction, we developed a mixed culture system consisting of hGCs and dorsal root ganglia (DRG) axon-oligodendrocyte co-cultures. DRG cultures were selected because they can be isolated efficiently and can form the long, extensive projections which are ideal for migration studies of this nature. Purified rat oligodendrocytes were then added on purified rat DRG axons and induced to myelinate. After confirming the formation of compact myelin, hGCs were finally added to the co-culture and their interactions with DRG axons and oligodendrocytes was monitored in real-time using time-lapse microscopy. Under these conditions, hGCs form tumor-like aggregate structures that express GFAP and Ki67, migrate along both myelinated and non-myelinated axonal tracks and interact with these axons through the formation of pseudopodia. Our ex vivo co-culture system can be used to identify novel cellular and molecular mechanisms of hGC migration and could potentially be used for in vitro drug efficacy testing.

Introduction

Glioblastoma is one of the most aggressive and lethal tumors of the human brain. The current standard of care includes surgical resection of the tumor followed by radiation1 plus concomitant and adjuvant administration of temozolomide2. Even with this multi-therapeutic approach, tumor recurrence is inevitable3. This is partly due to the extensive migratory nature of the tumor cells, which invade the brain parenchyma creating multiple finger-like projections within the brain4 that make complete resection unlikely.

In recent years, it has become evident that the aggressiveness of glioblastoma is due, in part, to the presence of a population of cancer stem cells within the tumor mass5,6, which exhibit high migratory potential7,8, resistance to chemotherapy and radiation9,10 and the ability to form secondary tumors11. GSCs are capable of recapitulating original polyclonal tumors when xenografted to nude mice5.

Despite the wealth of knowledge regarding the genetic background of glioblastomas, studies on glioma cell (GC) migration are currently hindered by a lack of efficient in vitro or in vivo migration models. Notably, while glioma cell-axonal interactions modulated by cellular and environmental factors are a core component of glioma invasion, to our knowledge there is currently no experimental system with the ability to model these interactions12,13,14. To address this deficiency, we developed an ex vivo culture system of primary hGCs co-cultured with purified DRG axon-oligodendrocytes that results in elevated expression of differentiated tumor markers as well as extensive migration and interaction of hGCs with myelinated and non-myelinated fibers. This ex vivo platform, due to its compartmentalized layout, is suitable for testing the effects of novel therapeutics on hGC migration patterns.,

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Protocol

The protocols for collection, isolation, and propagation of patient-derived human glioma cells were approved by the IRB committee of Rhode Island Hospital. All animals were maintained according to the NIH Guide for the Care and Use of Laboratory Animals. All animal use protocols were approved by the Institutional Animal Care and Use Committee of Rhode Island Hospital.

1. Media and buffer preparations

  1. Prepare 50 mL of Neurosphere Media: 1x Neuronal basal medium w/o vitamin A, 1x serum free supplement w/o vitamin A, 2 mM L-glutamine, 20 ng/mL epidermal growth factor (EGF), 20 ng/mL basic-fibroblast growth factor (FGF), 2 µg/mL heparin, and 1x antibiotic-antimycotic (anti-anti).
  2. Prepare 50 mL of supplemented Neuronal Basal Medium: 1x Neuronal basal medium, 1x serum free supplement, 4 g/L D-glucose, 2 mM L-glutamine, and 50 ng/mL nerve growth factor (NGF).
  3. Prepare 500 mL of N2B2 Medium: 1:1 Dulbecco's Modified Eagle Medium-Ham's F12 Nutrient Mixture (DMEM-F12), 1x Insulin-Transferrin-Selenium (ITS-G), 66 mg/mL bovine serum albumin (BSA), 0.1 mg/mL transferrin, 0.01 mg/mL biotin, 6.29 mg/mL progesterone, 5 µg/mL N-Acetyl-L-cystine (NAC), and 1 mM putriscine. Aliquot and freeze at -20 °C.
  4. Prepare 500 mL of C-Medium: 1x Minimum Essential Medium (MEM), D-glucose (final 4 g/L), 10% fetal bovine serum (FBS), and 2 mM L-glutamine. Aliquot and freeze at -20 °C. Add nerve growth factor (NGF) fresh before use (50 ng/mL).
  5. Prepare 200 mL of Papain Buffer: 1x Earle's Balanced Salt Solution (EBSS), 100 mM Mg2SO4, 30% Glucose, 0.25 M EGTA, and 1 M NaHCO3.
  6. Prepare 10 mL of DMEM-ITS-G Medium: 1x DMEM, 0.5% BSA, and 1x ITS-G.
  7. Prepare 200 mL of PB Buffer: 1x Dulbecco's phosphate-buffered saline (DPBS) without Ca2+ and Mg2+ and 0.5% BSA. Degas the buffer before use and keep on ice.

2. Isolation and culture of Glioma Stem Cell Neurospheres

  1. Isolation of Neurospheres
    1. Collect IRB approved and patient consented fresh human glioblastoma (GBM) tissue from the operating room. Transfer GBM samples on ice in DPBS containing 2x anti-anti to a BL2 certified biological safety cabinet.
    2. Transfer one 1 cm3 GBM sample with minimal necrosis or red blood cell contamination to a 60 mm plate and cut into 1 mm3 fragments; remove any excess DPBS.
    3. Digest the tissue with 5 mL of collagenase/dispase (1 mg/mL) for 30 min at 37 °C and gently swirl the dish every 5-10 min.
    4. Transfer digested fragments to a 50 mL tube and triturate by pipetting with a 10 mL pipette several times to dissociate the tissue.
    5. Add an equal volume of Neurosphere Media and triturate the tissue again; allowing the large fragments to settle.
    6. Remove media without tissue fragments and pass slowly through a 70 µM cell strainer placed over a fresh 50 mL tube.
    7. Repeat steps 2.1.5-2.1.6 until all the tissue has been dissociated, changing the cell strainer as needed.
    8. Spin the cell suspension at 110 x g for 10 min.
    9. Remove the supernatant and resuspend the pellet in 10 mL of ACK lysing buffer to remove red blood cell contamination. Incubate for 10 min at room temperature.
    10. Spin the cell suspension at 110 x g for 5 min.
    11. Remove the supernatant and resuspend cells in 10 mL of Neurosphere Media. Take 10 µL of the cell suspension and dilute with 10 µL of trypan blue. Count 10 µL of the cell suspension using an automated cell counter or hemocytometer and plate in 10 mL of Neurosphere Media at a density of 3 x 106 cells in 100 mm suspension culture plates.
    12. Add 2 mL of fresh Neurosphere Media to the culture 2-3 times a week.
      NOTE: Neurospheres will form in suspension within 3-4 weeks. After subculturing for 2-4 times, confirm the stemness via the limiting dilution assay and tumor recapitulation via xenographic transplantation in immunocompromised mice as previously described 22.
  2. Sub-culturing Neurospheres
    1. When spheres form and reach a diameter between 200-500 µm, transfer neurospheres to a 15 mL tube and spin at 110 x g for 5 min.
    2. Resuspend neurospheres in 1 mL of pre-warmed cell detachment solution.
    3. Transfer to a 1.5 mL tube and incubate at 37 °C for 5 min.
    4. Set a P200 pipette to 150 µL and triturate by pipetting up and down to dissociate neurospheres.
    5. Transfer dissociated cells to a 15 mL tube. Add 4 mL of Neurosphere Media and spin at 300 x g for 5 min.
    6. Remove the supernatant and resuspend cells in 5 mL of Neurosphere Media. Take 10 µL of the cell suspension and dilute with 10 µL of trypan blue. Count 10 µL of the cell suspension using an automated cell counter or hemocytometer and plate at a density of 1 x 106 cells/60 mm dish in a final volume of 4 mL.
    7. Refresh media 2 times a week and subculture cells as needed. Freeze whole neurospheres when desired in Neuronal basal media w/o vitamin A supplemented with 10% DMSO. Neurospheres may be used for experiments after P4.

3. Compartmented culture of Rat Dorsal Root Ganglia (DRGs), Oligodendrocytes (OPCs) and hGCs

  1. Preparation of compartmented culture dishes
    NOTE: Perform the following steps the days before the planned harvest of the DRGs.
    1. Assemble compartmented culture dishes.
    2. Dilute collagen stock solution to 500 µg/mL in sterile distilled H2O; mix thoroughly.
    3. With a sterile transfer pipette, fill a 35 mm culture dish with 2 mL of collagen solution; remove the solution, leaving a thin film of collagen behind and place it into the next 35 mm dish. Repeat this process, adding more collagen solution as needed, until all dishes have been coated.
    4. Once all plates are coated, place the plates in a 245 mm x 245 mm culture tray and lay three 1 mm x 1 mm gauze pads in the center of the tray.
    5. To polymerize the collagen, wet gauze pads with 1 mL of concentrated ammonium hydroxide and cover the trays for 15 min.
    6. Remove gauze pads and allow the 35 mm dishes to dry in the laminar flow hood.
    7. While dishes are drying, load the barrel of the syringe grease applicator with high vacuum grease. Place the compartmented chambers in a large mouth media bottle filled with distilled water. Sterilize both by autoclaving and allow to cool.
    8. File off the point of an 18-G needled to make a blunt tip. Sterilize in 70% ethanol. Attach the needle to the grease syringe.
    9. Sterilize the pin rake by soaking in 70% ethanol; allow it to air-dry in the laminar flow hood.
    10. Remove the lid from a dry, collagen coated 35 mm dish. Hold the dish between the thumb and pointer finger. Hold the pin rake with the other hand. Apply a firm pressure to create even 200 µM wide scratches across the center of the dish.
    11. Using a pasture pipette, place two drops of Supplemented Neuronal basal Medium in the center of the scratches.
    12. Repeat steps 3.1.10-3.1.11 until all dishes have been scratched.
    13. Dry the compartmented chambers in a laminar flow hood.
    14. With sterile hemostatic forceps, grasp one compartmented chamber by the center divider. Flip the hemostatic forceps so the bottom of the chamber is facing up.
    15. Apply silicone grease to the compartmented chamber, starting at the top. Ensure that grease is placed neatly and overlaps at all corners.
    16. Remove the lid from a 35 mm dish. Invert the dish and place the scratches over the chamber. Tap down on the bottom of the plate gently with a pair of forceps.
    17. Gently flip the plate over by using the hemostatic forceps. Release the forceps.
    18. Place a mound of grease at the base of the center compartment. Fill each chamber with Supplemented Neuronal Basal Medium (NB) and check for leaks. Seal leaks with silicone grease as needed.
    19. Continue assembling all culture dishes and store overnight at 37°, 5% CO2.
  2. Isolation of Rat DRG Neurons and Culture in the Compartmented Chamber
    1. Sacrifice a timed pregnant E16 Sprague-Dawley rat by CO2 asphyxiation or by chemical overdose.
    2. Place the animal in the supine position on a clean surface; disinfect the abdomen with 70% ethanol.
    3. Grasp the skin of the lower abdomen with forceps and lift gently.
    4. Using scissors, make an "I" incision along the midline of the animal, taking care not to puncture the muscles of the abdominal wall.
    5. With a clean pair of forceps, grasp the muscle wall and make a transverse incision with a clean pair of scissors using caution not to puncture the uterus or intestines.
    6. With a pair of blunt forceps, grasp the uterus and gently lift straight up out of the peritoneal cavity.
    7. Using fresh, sterile scissors, clip the connective tissue at the base of each uterine horn.
    8. Place the entire uterus in a sterile 100 mm tissue culture dish.
    9. Carry the uterus to a laminar flow hood for dissection.
    10. Remove embryos from the uterus, one at a time, by clipping through the amniotic sac and gently teasing the embryo out and into a 60 mm dish containing 5 mL of L-15 with 1x pen-strep.
    11. With fine forceps, place 3-4 embryos into a 60 mm dish containing 5 mL of L-15 with 1x pen-strep.
    12. Working under a dissecting microscope and with one embryo at a time, euthanize by decapitation.
    13. Lie the embryo ventral side up and remove the limbs and the tail.
    14. With fine forceps, make a midline incision in the animal.
    15. Remove internal organs and tissue to expose the dorsal structures, particularly the spinal cord which should be visible upon completion.
    16. Place one blade of micro-dissecting scissors between the vertebral column and the spinal canal and carefully cut through the vertebral column to expose the spinal cord. Take care not to clip through the spinal cord.
    17. With fine forceps, lightly grasp the rostral end of the spinal cord and slowly lift out of the embryo. The DRGs will be attached to the spinal cord.
    18. Transfer spinal cords and attached DRGs to a 35 mm dish containing 2 ml of L-15 with 1x pen-strep. Place on the ice.
    19. Once all spinal cords have been isolated, use fine forceps to individually pluck DRGs from the spinal cord. Place DRGs into a fresh 35 mm dish.
    20. If nerve roots are present on DRGs, clip them away.
    21. Remove prepared 35 mm dishes from the 37 °C, 5% CO2 incubator. Remove the media from the previous day. Place 80 µL of Supplemented Neuronal Basal Media (NBF) containing 10 µM 5-Fluoro-2'-deoxyuridine (FUDR) in the center compartment and 250 µL of media in each outer compartment.
    22. Place 2 ganglia in each center compartment. Return dishes to the 37 °C, 5% CO2 incubator.
    23. The following day add 2.5 mL of NBF.
    24. Feed the cultures following the schedule in Table 1, altering the schedule based on the day chosen to begin the DRG prep.
    25. On day 21 (or when axons reach the end of the distal compartments), either seed cultures with a GBM neurosphere (Proceed to step 3.2.26) and live image or myelinate the cultures with oligodendrocytes cultures (Proceed to step 3.3) and then seed with a GBM neurosphere after myelination.
    26. Replace Supplemented NB Medium in each distal compartmented chamber that will be seeded with a GBM neurosphere with Supplemented NB Medium containing 10% FBS.
    27. With a P20 pipette set to 10 µL, remove one GBM neurosphere from the culture dish. The GBM neurosphere should measure approximately 200 microns in size.
    28. Place the tip of the pipette in the distal chamber and expel the GBM neurosphere slowly so that it gently falls onto the axons in the portion of the distal chamber that is closest to the center chamber, over the axons near the center chamber. Be careful not to let the pipette tip disrupt the axons.
    29. Leave the culture in the biosafety cabinet for 1 h at room temperature to allow the GBM neurosphere to attach.
    30. Once the neurosphere has attached, very carefully replace the media in the distal compartment with Supplemented NB Medium.
    31. Live image cultures for 3-7 days, adding media as needed. In this case, a controlled CO2 live cell enclosure attached to the microscope (e.g., Zeiss Axiovert) was used to continuously monitor cell migration for 7 days using brightfield. Images were acquired every 10 min using the associated software. Repeat the same protocol using hGCs that have been stably transfected to express GFP and images were captured using a combination of brightfield and 488 nm laser.
      NOTE: Neurospheres may be transduced with lentivirus or transfected with plasmids or siRNAs. Compartments may also be treated with desired small molecule inhibitors to study migration.
  3. Myelination of DRG Axons with Oligodendrocytes
    1. The day before oligodendrocyte isolation, replace the NB Medium in the DRGs with C-Medium.
    2. Before dissection, place 10 mL of Papain Buffer into a 60 mm dish in the incubator to equilibrate.
    3. In a laminar flow hood, fill one 100 mm dish and one 60 mm dish with ice cold HBSS. Place on ice.
    4. Sacrifice a P2 rat pup by decapitation. Remove the skin using a pair of scissors. After the skin is removed, cut the skull along the midline with a pair of fine scissors.
    5. Gently remove the skull with fine forceps. Using a spatula, gently scoop the brain from the bottom of the skull and transfer to an inverted tissue culture plate lid.
    6. Remove the cerebellum and divide the cerebrum into two cerebral hemispheres. Remove the olfactory bulbs, hippocampus, and basal ganglia below the cerebral cortex of each hemisphere. Place the cerebral cortex in the 100 mm dish containing HBSS. Repeat steps 3.3.4-3.3.6 for the remaining animals.
    7. Working with one cortex at a time, remove the meninges with fine Dumont forceps. Place all meninges-free cortices into fresh 60 mm dishes with HBSS.
    8. Dice the cortical tissue into 1 mm3 pieces. Place on ice.
    9. Place equilibrated Papain Buffer into a 15 mL tube. Add 200 units of papain and 2 mg L-cysteine. Filter sterilize and add 200 µL of sterile DNase I.
    10. Remove HBSS from the diced brain tissue and replace with Papain Buffer from 3.3.2. Place dish in 37 °C, 5% CO2 incubator for 80 min, gently shaking every 15 min.
    11. Transfer the digested tissue to a 50 mL tube and add 2 mL of C-medium.
    12. Triturate with a 5 mL serological pipette to dissociate the tissue; allow larger pieces of tissue to settle. Remove supernatant and place into a sterile 15 mL tube.
    13. Add 2 mL of C-Medium and repeat trituration with a 5 mL serological pipette once more. Switch to a 1 mL pipette tip and triturate until the tissue is completely dissociated. Transfer to the 15 mL tube.
    14. Spin the triturated tissue at 300 x g for 15 min. Carefully remove supernatant and resuspend pellet in 8 mL of DMEM-ITS-G Medium.
    15. Pre-wet a sterile 30 µM cell strainer with 2 mL of PB Buffer. Place cell strainer over a 50 mL tube and filter the cell suspension 1 mL at a time. Rinse the filter with 5 mL of PB Buffer.
    16. Transfer the cell suspension to a 100 mm bacteriological plate; incubate for 15 min in a 37 °C, 5% CO2 incubator to allow microglia to attach.
    17. Remove media and place into a 15 mL tube. Rinse the plate gently with 2 mL of DMEM-ITS-G medium and transfer to the 15 mL tube.
    18. Resuspend the cell suspension gently. Take 10 µL of the cell suspension and dilute with 10 µl of trypan blue. Count 10 µl of the cell suspension using an automated cell counter or hemocytometer.
    19. Spin the cell suspension at 300 x g for 10 min.
    20. Aspirate supernatant completely and resuspend cells in 70 µL of PB Buffer for every 1 x 107 cells. Mix well and incubate at 4 °C for 10 min.
    21. Add 20 µl of anti-A2B5 microbeads for every 1 x 107 cells. Mix well and incubate at 4 °C.
    22. Add 1-2 mL of PB Buffer for every 1 x 107 cells and centrifuge at 300 x g for 10 min in a 4 °C centrifuge.
    23. Aspirate supernatant completely. Resuspend up to a total of 108 cells in 500 µL of PB Buffer. Keep on ice.
    24. Place a magnetic bead column in the magnetic field of a separator.
    25. Prepare the column by rinsing with 500 µL of PB Buffer. Apply the cell suspension to the column. Discard flow-through in a waste container (this contains un-labeled cells).
    26. Wash the column with 500 µL of PB Buffer 3 times. Discard flow-through.
    27. Remove the column from magnetic separator and place in a collection tube.
    28. Pipette 1 mL of PB Buffer onto the column. Flush the magnetically labeled cells by firmly pushing the plunger into the column.
    29. Add 4 mL of C-Medium to the cell fraction and spin at 300 x g for 10 min.
    30. Remove supernatant and resuspend in 5 mL of N2B2 Medium. Take 10 µL of the cell suspension and dilute with 10 µL of trypan blue. Count 10 µL of the cell suspension using an automated cell counter or hemocytometer.
    31. Plate oligodendrocytes at a concentration of 150,000 cells per compartmented chamber containing a DRG with fully extended axons.
    32. The following day change the media in the compartmented chamber to N2B2 to allow for myelination. Replace media every 2-3 days. Myelination will be complete in 14 days.
    33. On day 14, seed culture with a GBM neurosphere as in 3.2.26-3.2.32. Maintain myelinated cultures in N2B2 Medium for the duration of any experiments.
      NOTE: The protocol is presented as a flow chart schematic in Figure 1.

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Representative Results

In order to study the interaction of hGCs with axons, we generated purified DRG axons as previously described15,16,17,18. These purified DRG axons were then seeded with hGCs, which formed GFAP+/Ki67+ tumor-like structures integrated within the axonal network, while individual hGCs migrated either in association or between the axons (Figure 2). To determine how hGCs interact with myelinated axons, we seeded DRG axon cultures with purified rat oligodendrocytes and induced myelination as previously described19,20,21. Addition of hGCs on the myelinated DRG-oligodendrocyte co-cultures showed that hGCs migrate in association with the myelinated axons and away from the tumor mass through the formation of pseudopodia as shown in our recent paper (Figure 3)22. Oligodendrocyte myelination can be determined using Myelin Basic Protein (MBP) staining of the cultures. To quantify the migration of hGCs in these cultures we measured the total area of the culture occupied by the migrating hGCs using Image J software22.

MONDAY WEDNESDAY FRIDAY
DAY 1 NBF
WEEK 1 NB NBF NB
WEEK 2 NBF NB NB
WEEK 3 NB NB

Table 1: Feeding schedule of DRG cultures.

Figure 1
Figure 1: Schematic summary of the protocol. Please click here to view a larger version of this figure.

Figure 2
Figure 2: hGCs form tumor-like structures in co-culture with DRG axons. The culture was fixed and stained for the tumor markers GFAP (red) and Ki67 (green), while the axons were stained with Neurofilament (blue). Scale bar: 200 μm. This figure has been modified from our recently published paper22. Please click here to view a larger version of this figure.

Figure 3
Figure 3: hGCs migrate along myelinated axonal tracks. hGCs expressing the green fluorescent protein (GFP) migrate along myelinated axonal tracks stained red for MBP in the hGC-DRG-oligodendrocyte culture system. Scale bar: 200 μm. Please click here to view a larger version of this figure.

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Discussion

Migration studies for hGCs may be performed by using Boyden chamber systems or scratch assays. However, while these experiments fail to give any information regarding the interactions of tumor cells with other surrounding tissues, the present system can recapitulate GC interactions with myelinated and non-myelinated fibers. Furthermore, to study tumor formation and end-point migration, organotypic slice cultures of the rodent brain or in vivo implantation of glioma cells into the rodent brain or flank have previously been utilized23,24,25. More recent efforts on three-dimensional modeling of glioma cells have utilized systems like collagen layers26,27,28, astrocyte-based scaffolds29,30, extracellular matrix layers31, electrospun nanofibers32, and hydrogels33. While these experiments produce satisfactory end-point results regarding migration, they lack the ability to be studied in real-time by high-resolution microscopy.

Here, we have demonstrated a new approach to studying the migration of hGCs by preparing a DRG co-culture in a compartmented chamber. If access to fresh GBM tissue is available, hGCs can be isolated and cultured reliably. Although hGCs take a long time to form neurospheres initially, the cultures are easy to maintain and subculture once the spheres form. To ensure the success of the hGC culture, the tissue should be processed as quickly as possible. The time between resection and processing should be minimized as much as possible, and samples should always be transported on ice.

The DRG is also easy to isolate, extends its axons longer than cortical neurons and can be easily myelinated in culture with oligodendrocytes. hGC neurospheres or dissociated hGCs are co-cultured in the distal compartments of the chambers, allowing for live cell imaging and real-time quantification of cellular interactions with axons and myelinated fibers. Additionally, this protocol is not limited to only DRG cultures, as cortical neurons may also be isolated and myelinated with few modifications to this protocol. This model may also be used to study other forms of brain cancer.

While the DRG is relatively simple to isolate and maintain, the addition of the compartmented chamber is time consuming and creates a multitude of technical limitations that require training and practice to succeed. Critical to the success of this protocol is uniform collagen coating and scratching of the culture dishes because uneven or excessively thick collagen tends to peel. Extreme care should also be taken while placing silicone grease on the compartmented chambers. If even pressure is not used while dispensing the silicone grease, there will be gaps along the bottom of the compartmented chamber that will require sealing with excess grease to prevent media leakage. Additionally, minimum pressure should be used when adhering the compartmented chambers to the culture dish floor. If axons fail to cross out of the middle compartment after week one and the DRG looks otherwise healthy, it is likely that too much pressure was applied when placing the compartmented chamber or an excess amount of grease was used.

hGC migration along myelinated and non-myelinated axonal tracts in the brain has not been well described due to the lack of efficient and reproducible ex vivo models. We describe here the development of an innovative ex vivo culture system to study how glioma cells migrate along axons and myelin, a crucial component in developing new treatments that specifically target tumor cell migration. Our culture system is versatile since there is fluidic isolation between compartments, facilitating the ability to study various novel treatments or differing concentrations of substances that affect glioma cell migration. An additional advantage of our co-culture system is the ability to monitor hGC migration and the effects of the various treatments in real-time. The protocol described here is currently being used as the basis for the development of more sophisticated biomimetic 3-dimensional ex vivo systems using exclusively human cellular components that could be used to assess toxicology and efficacy of novel drugs.

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Disclosures

The authors have nothing to disclose.

Acknowledgments

This work was supported by internal funds of the Department of Neurosurgery, Brown University to N.T.

Materials

Name Company Catalog Number Comments
100 mm Suspension Culture Dish Corning 430591
2.5S NGF ENVIGO B.5025
60 mm Suspension Culture Dish Corning 430589
ACK Lysing Buffer Thermo Fisher A1049201
Ammonium Hydroxide Solution Fisher Scientific A669-500 Concentrated
Animal-Free Recombinant Human EGF Peprotech AF-100-15
Animal-Free Recombinant Human FGF-basic (154 a.a.) Peprotech AF-100-18B
Anti-A2B5 MicroBeads, human, mouse, rat Miltenyi Biotec 130-093-392
Antibiotic-Antimycotic (100X) Thermo Fisher 15240062
AutoMACS Rinsing Solution (PBS, pH 7.2) Miltenyi Biotec 130-091-222
B27 Supplement Thermo Fisher 17504044
B27 Supplement, minus vitamin A Thermo Fisher 12587001
Bacteriological Plate BD Falcon 351029
Biotin Sigma B4639
BSA Sigma A9418
Campenot Chamber Tyler Research CAMP-10
Cell Culture Dish Corning 430165 35mm X 10mm
Cell Strainer BD Falcon 352350 70 uM, Nylon
Cell Strainer BD Falcon 352340 30 uM, Nylon
Collagenase/Dispase Roche 11097113001
Cultrex Rat Collagen I Trevigen 3440-100-01
D-Glucose Sigma G5146
DMEM Thermo Fisher 10313021
DNase I Sigma D7291
Dow Corning High-Vacuum Grease Fisher Scientific 14-635-5D
Dumont #5 Forceps Roboz RS-5045
E16 Timed Pregnant Sprague Dawley Rat
EBSS Sigma E7510
EGTA Sigma E3889
FBS Hyclone SH30070.02
FUDR Sigma F0503
GlutaMAX Supplement Thermo Fisher 35050061
Ham's F-12 Nutrient Mix Thermo Fisher 11765054
HBSS Thermo Fisher 14175095
Hemostatic Forceps Roboz RS-7035
Heparin Sodium Salt, 0.2% in PBS Stem Cell Technologies 07980
Hypodermic Needle, 18G BD 511097
Insulin-Transferrin-Selenium G Thermo Fisher 41400045
L-Cysteine Sigma C7477
L-Glutamine Thermo Fisher 25030081
Leibovitz's L-15 Medium Thermo Fisher 11415064
MACS BSA Stock Solution Miltenyi Biotec 130-091-376
MACS MultiStand Miltenyi Biotec 130-042-303
MEM Thermo Fisher 1190081
Mg2SO4 Sigma M2643
MiniMACS Separator Miltenyi Biotec 130-042-102
MS Columns plus tubes Miltenyi Biotec 130-041-301
NAC Sigma A8199
NaHCO3 Sigma S5761
Neurobasal Medium Thermo Fisher 21103049
Neurobasal-A Medium Thermo Fisher 10888022
Ordinary forceps
P2 Sprague Dawley Rat Pups
Papain Worthington LS003126
Penicillin-Streptomycin Thermo Fisher 15140148
Pin Rake Tyler Research CAMP-PR
Progesterone Sigma P8783
StemPro Accutase Cell Dissociation Reagent Thermo Fisher A1110501
Syrine Grease Applicator Tyler Research CAMP-GLSS
Transferrin Sigma T2036
Uridine Sigma U3003

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