Culturing and imaging glioma stem cells in 3D collagen matrices

Abstract

Methods to maintain human glioma stem cells as neurosphere cultures and image their dynamic behavior in 3D collagen matrices are described. Additional approaches to monitor glioma stem cell differentiation into mesenchymal-type cells, along with example data are included. Together, these approaches enable glioma stem cell differentiation to be controlled while maintaining the cells in culture, as well as allowing cell dynamics to be captured and analyzed. These methods should be helpful for those seeking to understand the molecular mechanisms driving the invasion of glioma cells through three-dimensional environments.

INTRODUCTION:

Glioma cells are extremely effective at moving through three-dimensional environments, like those found within the brain (Beadle et al., 2008; Venkataramani et al., 2022; Zepecki et al., 2019). This is precisely why this malignant disease is so lethal. Glioma cells are able to invade through the narrow confines of the CNS, leading to diffuse tumor margins and difficulty in removing all of the invasive cells through surgical resection. Thus, it is beneficial to understand the molecular mechanisms driving glioma migration with the goal of creating therapeutic strategies to limit their movement and prolong patient survival.

Migrating cells in general are highly responsive to their physical environment (Yamada & Sixt, 2019). Factors like dimensionality, porosity and composition can each dictate the precise mechanisms by which cells move. For example, non-muscle myosin II activity is dispensable for movement across 2D surfaces, but absolutely essential for glioma migration through 3D matrices (Beadle et al., 2008). Therefore, the in vitro tissue culture model used to investigate the mechanisms of glioma invasion should be selected partially based on the types of environments through which these cells move in the brain.

These protocols are designed for studying the molecular mechanisms of glioma cell movement using an in vitro model that maintains physiological relevance and at a level of detail that in vivo can be inaccessible and cost-prohibitive. Type I collagen is an in vitro 3D model that has been used to investigate the mechanisms of glioma invasion (Payne & Huang, 2013), as well as the mechanisms of single cell migration (Doyle et al., 2015). We note that Type I collagen is found minimally in the brain, and the matrix therefore does not recapitulate the typical glioma biochemical environment, thus presenting a methodological limitation. However, 3D collagen is accessible, has a highly tunable architecture, and has been successfully used to elucidate molecular mechanisms in glioma cells. Additionally, the method for monitoring differentiation is visual and devised for convenience and low cost, and is therefore limited in its ability to provide a definitive readout of differentiation.

Here we present several protocols that describe how to culture and maintain glioma stem cells, how to embed them in 3D collagen gels, and provide examples of how to measure their motility, as well as monitor the differentiation of these cells into a mesenchymal subtype.

BASIC PROTOCOL 1

Culturing human glioma stem cells as neurospheres

This protocol describes how to culture primary glioma stem cells (GSCs) as neurospheres in suspension. It includes a version of the culture medium originally used to establish that GSCs are optimally maintained in the same serum-free culture conditions as normal neural stem cells, i.e., with medium that critically contains Neurobasal medium, basic FGF and EGF (“NBE”) (Lee et al., 2006). The same reagents and general procedure have been previously used (Guetta-Terrier et al., 2021; Zepecki et al., 2021; Zepecki et al., 2019) to maintain GSCs derived from patient tumors by the procedure described in Galli et al. (2004).

Materials:

Human primary glioma stem cells (patient derived, used at passages ~18–30); kindly provided by Dr. Nikos Tapinos

Complete Neurobasal media (see recipe in Reagents and Solutions)

StemPro Accutase cell dissociation reagent (Gibco / Fisher Scientific, cat. no. A1110501)

Neurobasal-A medium (Gibco / Fisher Scientific, cat. no. 10888022)

Cell culture microscope

37°C water bath

Biological safety cabinet

Automated pipette with serological pipet tips

15 mL conical centrifuge tube

Cell culture centrifuge

P1000 pipette and filtered tips

Cell counter (e.g., hemocytometer with trypan blue)

60 mm culture dish (CELLTREAT, cat. no. 229660; or ultra-low attachment from Corning, cat. no. 3251)

Protocol steps with step annotations:

Preparation

• 1Confirm by phase-contrast microscopy that neurospheres are an appropriate size to passage (Fig. 1).

  Neurospheres should be approximately 150–400 μm in diameter when ready to passage. In our experience, it takes about 7–14 days of growth for neurospheres to reach this size after re-plating. The core of a neurosphere will start to darken at larger sizes (especially closer to 500 um) due to cell death from hypoxia (Fig. 1). This is normal as some neurospheres in culture will grow faster than others and exhibit darkened cores. We routinely passaged neurospheres that included some of these dark cores without issue, but tried to passage cultures before all the neurospheres exhibited this characteristic.

• 2Warm the Complete Neurobasal media and Neurobasal-A medium to 37°C, and StemPro Accutase to room temperature.

  Prewarming media prevents cell stress caused by cold media. Heating Accutase to 37°C inactivates it. Aliquot Accutase into 1.5 mL vials to conveniently warm more quickly to room temperature.

Figure 1.

Phase-contrast image of GSC neurospheres in culture. Larger neurospheres show some core darkening indicative of cell death due to hypoxia. Scale bar represents 400 μm.

Procedure

All steps except those that require centrifugation should be performed under sterile conditions.

• 3Transfer neurospheres and media to a 15 mL conical tube.

  Neurospheres are visible by eye when approximately 150 μm and larger. Pipet gently at this stage to avoid rupturing neurospheres. If using a small pipette tip (e.g. p1000) rather than serological pipet, be particularly gentle with large spheres which are more vulnerable to shear force through the tip opening. Alternatively, cut the tip for a wider opening. In cases where some neurospheres adhere to the culture dish, they can be dislodged by pipetting a stream of media onto them or by tapping the dish on a stiff surface.

• 4Centrifuge at 100 × g for 5 minutes using a swinging bucket rotor, or 10 minutes using a fixed angle rotor.
• 5Remove supernatant. Resuspend the pelleted cells in 200 μL - 400 μL of StemPro Accutase.

  Media inactivates Accutase by dilution. A small volume of Accutase therefore requires removal of as much media/supernatant as possible without accidentally removing loosely pelleted cells. If some supernatant cannot be removed, it may be necessary to compensate by increasing the volume of Accutase.

• 6Set a P1000 pipette to 25 μL less than the volume of the added Accutase, and triturate neurospheres for ~10–20 seconds immediately and every 1–3 minutes thereafter to help dissociate cells, while allowing them to settle at the bottom of the tube between each trituration. This step should not exceed 10 minutes in total to maintain cell viability.

  Setting the pipet to 25 μL less than solution volume helps avoid introducing air bubbles which can also reduce cell viability. Triturating (pipetting up and down) too forcefully can result in cell death, but too gently will not dissociate spheres.

• 7Add Neurobasal-A immediately after neurospheres appear by eye to have disaggregated (i.e., when you see a cloudy solution without obvious spheres), bringing the solution volume to 5 mL.
• 8Centrifuge cells and media at ~225 × g for 5 minutes.
• 9Remove supernatant. Resuspend the pelleted cells in 1 mL of Complete Neurobasal medium.
• 10Count the cells using your preferred method.
• 11To maintain the culture, plate the disaggregated cells at an appropriate density (Table 1), and incubate at 37°C and 5% CO₂ in humidified air.

  If needed, cell counts may be reduced to 33% of suggested numbers (depending on cell line) without negatively impacting growth.

• 12Replace the media every 2–3 days by either: (a) Repeating steps 2–3 and resuspending in fresh warm Complete Neurobasal media, or (b) if neurospheres can be distinguished by eye, allowing neurospheres several minutes to settle by gravity to the bottom of the dish before carefully aspirating most of the media and replacing with fresh media.
• 13Allow cells to grow for 7–14 days to re-establish neurospheres of appropriate size for passage or plating.

  Note that this time period may vary by cell line, so monitor cells to confirm the best timetable for your particular cells.

Table 1.

Cell seeding guidelines by size of dish for culture maintenance.

Culture dish size	Cell #	Media volume
35 mm	5 × 105	2 mL
60 mm	1 × 106	4 mL
100 mm	3 × 106	10 mL

BASIC PROTOCOL 2

Inducing GSC adherence and monitoring their differentiation into mesenchymal cells

Cells that interact with extracellular matrix such as fibronectin or collagen can differentiate over days to weeks (Payne & Huang, 2013; Singh & Schwarzbauer, 2012; Yu et al., 2018). GSCs can also be induced to adhere to a surface and differentiate by exposure to serum (Joseph et al., 2015). It is therefore important when using GSCs in culture or experiments to have visual reference points to indicate the extent of differentiation. The below protocol uses a fibronectin substrate and serum-supplemented media, as well as fixed cell fluorescent imaging, to monitor and illustrate changes in differentiating GSCs over a 2-week period. This protocol predominantly uses vimentin localization and cell morphology to assess cellular differentiation following plating on fibronectin.

Materials:

All materials listed in Basic Protocol 1, except for 60 mm culture dishes, plus:

Complete DMEM (see recipe in Reagents and Solutions)

4% paraformaldehyde in PBS 1X (see recipe in Reagents and Solutions)

PBS 1X (see recipe in Reagents and Solutions)

0.25% TritonX-100 in PBS 1X (see recipe in Reagents and Solutions)

0.2% Bovine serum albumin (BSA) solution in PBS 1X (see recipe in Reagents and Solutions)

Mouse anti-vimentin antibody (MilliporeSigma, cat. no. CBL202)

Goat anti-mouse AlexaFluor 488 (Thermofisher/Invitrogen cat. no. A-11001)

Rhodamine phalloidin (ThermoFisher/Invitrogen, cat. no. R415)

DAPI (ThermoFisher, cat. no. 62248)

Fibronectin coated glass-bottom dishes (see Support Protocol 1)

Fluorescent microscope with a high-power objective (e.g. 63x)

Protocol steps with step annotations:

Prepare Reagents

• 1Warm fibronectin-coated dishes to room temperature.

Plate cells

These steps should be performed under sterile conditions:

• 2Complete steps #1–7 of Basic Protocol 1.

  If you need to retain some of these cells for maintaining culture but intend to differentiate cells using Complete DMEM or 7.5% FBS as described in step 4, then count cells at this step and aliquot those required for maintenance to a separate conical tube.

• 3Centrifuge cells and media at ~225 × g for 5 minutes.
• 4Remove supernatant. Resuspend pelleted cells in 1 mL of complete DMEM.

  As an alternative to complete DMEM, use complete neurobasal media supplemented with 7.5% fetal bovine serum but without bFGF, EGF and heparin.

• 5Count cells using your preferred method.
• 6Make calculations in preparation for plating cells onto 5 fibronectin-coated dishes at a density of, e.g., 2 × 10⁴ cells per dish:

  Calculate the number of cells required for 5 dishes.

  2 × 10⁴ cells * 5 dishes = 1 × 10⁵ cells

  Calculate some number of additional cells to include to allow for pipetting error.

  2 × 10⁴ cells * 0.5 dishes = 1 × 10⁴ cells

  Calculate the total number of required cells.

  1 × 10⁵ + 1 × 10⁴ = 1.1 × 10⁵ cells

  Calculate the total volume of media needed, assuming each plate requires 2 mL of media. E.g.:

  5 dishes + 0.5 dish (to represent pipetting error) = 5.5 dishes

  5.5 dishes * 2 mL per dish = 11 mL

  A low density such as 2×10⁴ helps avoid reaching cell confluence over the 2-week time course, but this may need to be optimized. The “0.5 dish” in the above calculation represents a convenient way to account for potential pipetting error and avoid having insufficient cell-media mixtures when plating.

• 7Transfer the required amount of cells into a new conical tube, and add fresh media to bring the total volume to 11 mL.
• 8Aspirate solution from fibronectin-coated plates immediately before adding GSC cells and media. Do not wash before adding cells.
• 9Triturate cells in the conical tube to ensure even mixing.
• 10Plate 2 mL of the GSC cell-media mixture onto each of five fibronectin dishes. Consider the day of plating as Day 0.
• 11Incubate plated cells at 37°C and 5% CO₂ in humidified air.
• 12Change the media of incubated dishes every 3–4 days by aspirating media and adding fresh, warm media.
• 13At days 1, 3, 6, 10 and 14 after plating, remove a dish from incubation to fix and stain.

Fix and stain cells

These steps do not require sterile conditions. All incubations should be done at room temperature.

• 14Prepare 4% paraformaldehyde (PFA) in PBS 1X, 0.25% TritonX-100 in PBS 1X, and 0.2% BSA in PBS 1X solutions (See Reagents and Solutions section for recipes).

  PFA and BSA solutions should be prepared fresh each time. 4% PFA stored long term is prone to oxidation, and BSA promotes bacterial growth.

• 15To fix cells, aspirate supernatant and add 2 mL of 4% paraformaldehyde to each dish. Incubate for 10 minutes.

  Optionally, wash cells with PBS before adding 4% paraformaldehyde, especially if you encounter high background fluorescent signal. When adding any solution during fixation/staining it is best to pipette gently into the corner of the dish to avoid disrupting or washing away cells in the inner well.

• 16Aspirate 4% paraformaldehyde and wash cells with 2 mL PBS.

  If needed it is okay to pause at this point and resume the protocol sometime within a week. Minimize the delay for best results. Simply add PBS back to the dish(es), seal with parafilm and store at 4°C. Aspirate PBS before resuming the next step.

• 17To permeabilize cells to antibodies, aspirate PBS and add 2 mL of 0.25% TritonX-100 to each dish.
• 18Incubate for 5 minutes.
• 19Aspirate TritonX-100 and wash cells once with 2 mL PBS.
• 20To reduce non-specific binding of antibodies, aspirate PBS, and add to each dish 2 mL of 0.2% BSA as a blocking buffer.
• 21Incubate for 30 minutes.
• 22
  During the 30-minute incubation, dilute the primary antibody into 0.2% BSA solution to make the primary antibody solution (“1°αb solution”) in a 1.5 mL microcentrifuge tube:

  • a
    For each dish, add 200 μL of 0.2% BSA solution to the 1.5 mL tube. E.g., 5 dishes * 200 μL = 1000 μL 0.2% BSA.
  • b
    Add the 1°αb (mouse anti-vimentin) at a ratio of 1:500 to the 0.2% BSA solution.

  For antibody reagents reconstituted to a concentration of 1 μg/μL, use Table 2 to quickly determine the volume of antibody required for dilution based on the required volume of 0.2% BSA and desired ratio of antibody. E.g., using Table 2, a 1:500 ratio of primary antibody to 1000 μL of 0.2% BSA solution requires 2 μL antibody.

• 23Aspirate BSA solution. Do not wash.
• 24Add 200 μL of 1°αb solution to each dish.
• 25Incubate for 1 hour at room temperature.
• 26
  During the 1-hour incubation, make the secondary antibody solution (“2°αb solution”) by diluting antibodies in 0.2% BSA in a 1.5 mL microcentrifuge tube:

  • c
    Add 200 μL of 0.2% BSA to the 1.5 mL tube. Scale up as necessary for multiple dishes.
  • d
    Add the secondary antibodies to 0.2% BSA at the following ratios:

    • 1:1,000 anti-mouse IgG-488
    • 1:2,000 rhodamine phalloidin
    • 1:10,000 DAPI

  Rhodamine phalloidin and DAPI do not require a primary antibody because they bind directly to filamentous actin (F-actin) and DNA, respectively.

• 27Aspirate the 1°αb solution and wash cells once with 2 mL of PBS.
• 28Add 200 μL of 2°αb solution to each dish.
• 29Incubate in the dark for 1 hour at room temperature.
• 30Aspirate the 2°αb solution and wash twice with 2 mL of PBS.
• 31Seal each dish in parafilm and either store at 4°C until ready to image, or image immediately.
• 32Image using your preferred inverted fluorescent microscope.
• 33Observe stained GSCs for their size and the distribution of the vimentin network. Fig. 2 shows representative images of GSC differentiation over 14 days.

Table 2.

Look-up table for desired dilutions. Table shows examples of what volume (μL) of antibody to add to a given volume of 0.2% BSA solution when the antibody has a 1 μg/μL concentration. The row for 25 μL is for adding an additional volume to account for pipetting error.

0.2% BSA	1:200	1:500	1:1,000	1:2,000
25 μL	0.13	0.05	0.03	0.01
200 μL	1.0	0.4	0.2	0.1
400 μL	2.0	0.8	0.4	0.2
600 μL	3.0	1.2	0.6	0.3
800 μL	4.0	1.6	0.8	0.4
1000 μL	5.0	2.0	1.0	0.5

Figure 2.

Time course study and immunofluorescent staining of GSCs on 2D fibronectin. GSCs fixed at days 1, 3, 6, 10 and 14, and stained for vimentin (green), filamentous actin (red) and the nucleus (blue). Scale bars at 20 μm.

SUPPORT PROTOCOL 1

Preparing fibronectin-coated dishes for cell microscopy

GSCs that are non-adherent in serum-free culture conditions can be induced to adhere by coating a surface with fibronectin matrix proteins (Yu et al., 2018). This protocol describes how to coat a glass bottom dish with fibronectin to effect GSC adherence in preparation for cell motility studies. The same procedure can be used for making fibronectin-coated culture dishes for passaging adherent GSCs, though a protocol for passaging adherent GSCs is not considered here. This protocol should be completed in sterile conditions.

Materials:

Chemicon human plasma fibronectin purified protein (Millipore, cat. no. FC010, 1 mg/mL)

Gibco Hanks’ balanced salt solution (HBSS), calcium, magnesium, no phenol red (Fisher Scientific, cat. no. 14025092)

Biosafety cabinet

15 mL conical centrifuge tube

Automated pipet with serological pipet tips

P200 or P1000 pipette with tips

35mm glass-bottom dishes (WPI, cat. no. FD35-100) or 50mm glass-bottom dish (WPI, cat. no. FD5040)

Protocol steps with step annotations:

Calculate volumes and prepare solution

• 1Fibronectin should be diluted into 1x HBSS at 10 μg/mL (1:100). Calculate the total amount of fibronectin and HBSS required for the number of dishes that will be coated. See Table 3 for reagent volumes by dish size.

  Fibronectin-HBSS solution should be made fresh each time. The specific fibronectin reagent listed in Materials should not be frozen. However, other fibronectin products may permit freezing, in which case it may be possible to aliquot fibronectin-HBSS solution to store at −20°C.

Table 3.

Fibronectin (Fn) solution by dish size.

Dish size	Reagent amounts	Amt of solution / dish
35 mm glass-bottom dish	10 μL Fn + 990 μL HBSS	1 mL
50 mm glass-bottom dish	20 μL Fn + 1,980 μL HBSS	2 mL

Protocol steps and annotations

• 2In a 15 mL conical tube, dilute fibronectin into HBSS and triturate to mix.

  Do not vortex fibronectin because it will precipitate out of solution and not re-dissolve.

• 3Add 1 mL fibronectin/HBSS solution to cover the bottom of each glass-bottom dish.

  Note that if the solution is added only to the inner well of a 35 mm dish, the concentration of fibronectin will be double compared to if the solution covers the entire bottom of the dish.

• 4Incubate at 37°C for 1 hour.
• 5After incubation, dishes may be used immediately or stored for later use. To store, seal each with parafilm and refrigerate at 4°C. Fibronectin coated dishes stored in this manner can be used up to 1 month after preparing.

BASIC PROTOCOL 3:

Embedding GSCs in a 3D collagen matrix to study their invasive behavior

Cells, including glioma cells, migrate differently in 2D and 3D environments (Duval et al., 2017; Tamaki et al., 1997; Yamada & Sixt, 2019). The architecture of 3D collagen can be precisely tuned by adjusting such factors as collagen concentration, pH and temperature to alter physical features like porosity, fiber size and stiffness (Doyle, 2016). 3D collagen has accordingly been used to clarify aspects of glioma cell migration such as cell morphology, velocity, invasiveness, genetic expression, and interaction with various ECM architectures (Fayzullin et al., 2019; Yang et al., 2010). The following protocol describes how to embed GSCs in 3D type-I collagen in preparation for imaging studies of cell migration, and in particular for live-cell phase-contrast imaging, with assistance from Support Protocol 2. This approach can also be used to study other cell attributes such as intracellular pressure (Petrie & Koo, 2014). The preparation of collagen solution is partly adapted from Doyle (2016), and we recommend that glass bottom dishes used in this protocol be silanized as established by Doyle (2009), to reduce the risk of collagen detachment. Finally, the below protocol uses type I collagen at a concentration of 1.7 mg/mL which may be adjusted as needed to control matrix stiffness and porosity (Miron-Mendoza et al., 2010). Collagen should be made and applied to plates under sterile conditions.

Materials:

All materials listed in Basic Protocol 1, except for 60 mm culture dishes, plus:

Type I collagen, rat tail (Corning, cat. no. 354236)

10x Reconstitution buffer (RB) (see recipe in Reagents and Solutions)

10x DMEM (Sigma Aldrich, cat. no. D2429-100ML)

1M NaOH (see recipe in Reagents and Solutions)

Alexa 647 ester dye (optional) (ThermoFisher, cat. no. A20006)

35 mm glass-bottom dishes (WPI, cat. no. FD35-100), optionally silanized

Ice bucket with ice

Vortex mixer

P200 or P1000 pipette + tips

Protocol steps with step annotations:

Make calculations

• 1
  Calculate reagent volumes to make a working collagen solution. As an example, Table 4 summarizes calculations and reagent amounts to make an arbitrary amount of working collagen solution that can be quickly scaled up or down depending on experimental requirements.

  1. Choose your desired final working concentration of collagen; e.g., 1.7 mg/mL.

     • A lower concentration increases gel porosity and vice versa.
  2. Calculate the final volume of collagen solution, using: (a) stock collagen concentration, which varies depending on the lot; (b) a stock collagen volume of 512 μL, which is proportional to other reagents; (c) the desired final working concentration of collagen chosen above.
     For example, for a stock collagen concentration of 4.40 mg/mL, the final volume is calculated by:
     C₁ V₁ = C₂ V₂
     (4.40 mg/mL)(512 μL) = (1.7 mg/mL)(V₂)
     V₂ = 1325 μL
     where:
     C₁ = stock collagen concentration = 4.40 mg/mL
     V₁ = stock collagen volume = 512 μL
     C₂ = final collagen concentration in working solution = 1.7 mg/mL
     V₂ = final volume of working solution
  3. Calculate the amount of Complete Neurobasal media needed to bring the solution up to the final volume:
     1325 − (512 + 64 + 64 + 7.5 + 5) = 672.5 μL Complete Neurobasal media needed
• 2
  Calculate the volume of collagen solution required considering the following factors:

  1. Number of dishes.
  2. Volume of collagen required per dish based on desired thickness of the collagen gel. For example, for a 35-mm glass-bottom dish with a 23-mm inner well, 125 μL of collagen solution polymerized at 37°C makes an approximately 200 μm thick gel. Scale as necessary.
     The gel surface may not polymerize at a uniform height due to the solution’s viscosity and the inherent variability in coating the dish. Also, note that when imaging with an inverted microscope, a gel that is too thick may: 1) put the gel’s surface out of focal range for some higher magnification objectives (e.g. 63x), and 2) reduce the clarity of fluorescently tagged/stained cells closer to the gel surface.
  3. Pipetting error due to solution viscosity. Add an additional 25 μL of solution per dish to account for error. For 4 dishes with gel thickness of 200 μm (125 μL), the amount of collagen solution required is:
     (125 μL + 25 uL) * 4 dishes = 600 μL collagen solution
     In this example, the Table 4 recipe makes a final volume in excess of experimental needs, and therefore could be scaled down to 50% for a final volume of 663 uL, as shown in Table 5. Alternate calculations can be made to arrive at volumes exactly matched to experimental needs to eliminate excess, but scaling the provided recipe is typically faster and lowers the risk for error when making calculations on the fly.
• 3Calculate the number of cells required for the collagen solution to ensure the expected number are plated for experiments.

  To allow for pipetting error, step 2 suggests making more collagen solution than is required for plating. It is therefore necessary as described in this step to add more cells to the collagen solution than is required for plating, to accommodate the excess collagen solution and ensure the correct concentration of cells is plated.

  Scenario: Let the example calculations from steps 1 and 2 serve as a guide to prepare enough collagen solution for 4 dishes, at 125 μL collagen solution per dish with room for error. The example calculations from step 1 (Table 4) would produce more than double the collagen solution we need. Therefore, we can actually make collagen at 50% of the original recipe (Table 5) to yield 663 μL collagen solution, including 336 μL of complete neurobasal media in which cells will be suspended. To ultimately plate, e.g., 5 × 10⁴ cells per dish, first calculate the number of cells needed for the collagen solution.

  Variables in calculation:

  1. Required number of dishes (from step 2a). E.g, 4 dishes.
  2. Required volume of collagen solution for experiments (from step 2b&c). E.g., 150 μL * 4 dishes = 600 μL.
  3. Required number of cells for experiments. E.g., 5 × 10⁴ per dish * 4 dishes = 2 × 10⁵ total cells.
  4. Final volume of collagen solution (from step 1b). E.g., 663 μL
  5. Required volume of complete media (from step 1c). E.g., 336 μL.

  Calculation: (663 μL collagen solution / 600 μL used solution) * 2 × 10⁵ total cells = 2.21 × 10⁵ required cells in 336 μL complete media

Table 4.

Recipe for collagen solution. The recipe can be scaled up or down as needed (as exemplified in Table 5). The final volume and the volume of Complete Neurobasal media depend on the concentration of the initial stock collagen. Alexa 647 may be used to dye the collagen if the sample will be additionally stained later for fixed-cell imaging.

Reagent	Amount (μL)
Rat tail Type I collagen concentration = 4.40 mg/mL	512
10x Reconstitution buffer	64
10x DMEM	64
1M NaOH	7.5
Alexa 647 dye (optional)	5
Complete Neurobasal media	672.5
Final volume	1325 μL

Table 5.

Collagen solution recipe at 50% of the Table 4 recipe.

Reagent	Amount (μL)
Rat tail Type I collagen concentration = 4.40 mg/mL	256
10x Reconstitution buffer	32
10x DMEM	32
1M NaOH	3.75
Alexa 647 dye (optional)	2.5
Complete Neurobasal media	336
Final volume	663 μL

Prepare reagents

• 4Silanize glass bottom dishes (see Doyle, 2009).

  Using silanized dishes is highly recommended to minimize the risk of a gel detaching from the glass surface.

• 5Confirm by phase-contrast microscopy that neurospheres are appropriate in size to passage (Fig. 1).

  Neurospheres should be approximately 150–400 μm in diameter when ready to passage. In our experience, it takes about 7–14 days of growth for neurospheres to reach this size after re-plating. The core of a neurosphere will start to darken at larger sizes (especially closer to 500 μm) due to cell death from hypoxia (Fig. 1). This is normal as some neurospheres in culture will grow faster than others and exhibit darkened cores. We routinely passaged neurospheres that included some of these dark cores without issue, but tried to passage cultures before all the neurospheres exhibited this characteristic

• 6Warm the Complete Neurobasal media to 37°C. Warm StemPro Accutase to room temperature; 37°C inactivates Accutase.

  Consider aliquoting Accutase into 1.5 mL vials for faster warming to room temperature.

• 7On ice, chill aliquots of 10x reconstitution buffer, 10x DMEM, 1M NaOH, and a 15 mL conical tube for mixing the solution.

Prepare cells for collagen solution

• 8Complete steps 3–7 of Basic Protocol 1 to dissociate cells in neurospheres.
• 9Count cells using your preferred method.
• 10Transfer into a new 15 mL conical tube a volume of media that contains the total number of cells required for the collagen solution, as calculated in Step #3. (Cells in the original tube can be used to continue culture).
• 11Centrifuge the new tube at ~225 × g for 5 minutes.

Prepare and apply collagen solution

• 12During centrifugation, make the collagen solution on ice. Add reagents from Table 4 to a chilled 15 mL conical tube, except for complete neurobasal media. Reagents should be added in the order they are listed, and in the amounts calculated in Steps #1 and #2. To ensure homogenization, vortex the conical tube briefly (4–5 pulses) or mix well by trituration each time after adding 10x DMEM, 1M NaOH, and Alexa 647 (optional).

  To more easily pipet the collagen, cut the end off a pipette tip to create a wider hole. Avoid repeated pipetting up & down due to viscosity. Try to minimize bubbles as they can disrupt the homogeneity of the gel and weaken its adherence to the glass. Alexa dye is required only to visualize collagen fibers in fluorescent imaging, and is presented here as an option in case you would like to fluorescently fix and stain samples after using them for phase-contrast imaging as described in Support Protocol 2. Omit the dye if there’s no intention for fluorescent imaging.

• 13When centrifugation is complete, remove supernatant, and resuspend cells with the volume of complete neurobasal media required by the collagen recipe (e.g., 336 μL), as calculated according to Step #1c.
• 14Add the complete volume of resuspended cells to the collagen solution and triturate or pulse vortex to mix.
• 15Place a glass-bottom dish on ice, and evenly spread your desired amount of collagen solution with cells (step #2b; e.g., 125 μL) to cover the inner well. Avoid introducing bubbles. Repeat for each needed dish.

  Distribute collagen from the pipet as evenly as possible, but be prepared to use the pipet tip to help spread it out, especially when working with lower volumes. It’s ok to make contact between the pipet tip and glass bottom. Tilting the dish back and forth can help even the coverage.

• 16As the application of collagen and cells is completed for each dish, incubate at 37°C, 5% CO₂, for between 45 minutes to 1 hour.

  Cold and acidity prevent collagen polymerization. Warming to 37°C and neutralizing the pH should trigger collagen polymerization within 15–30 minutes.

• 17Add 2 mL of complete neurobasal media to each dish and incubate for ~48 hours in preparation for live-cell phase-contrast velocity imaging. See Support Protocol 2.

  Cells require at least > 24h to adhere, polarize, and begin to migrate.

SUPPORT PROTOCOL 2

Phase-contrast imaging with a tiled-matrix to study cell migration in a 3D gel

GSCs may migrate beyond a single field of view during live imaging lasting longer than several hours. It is therefore beneficial when imaging their migration to capture a matrix of adjacent fields of view, or “tiles”, in the XY plane and later assemble them into a single large field of view. This helps ensure that enough cells remain in focus for the duration of the experiment for analysis such as cell velocity or migration dynamics, as inevitably some cells in the 3D gel will move away from the plane of focus. The following protocol outlines the set up and key parameters for imaging live GSC cells in a 3D gel over time, using phase-contrast microscopy with a tiled-matrix capture.

Materials:

Human GSC cells in 3D collagen (see Basic Protocol 3)

Complete neurobasal media (see recipe in Reagents and Solutions)

Any relevant pharmacological treatments

Inverted confocal microscope with:

Phase-contrast capability

Phase-contrast objective, especially in the range of 20x-40x

Incubation chamber (37°C, 5% CO₂, humidified air)

Motorized stage

Microscopy software capable of automated capture of a tiled matrix over time, including the ability to automatically move the stage to pre-saved positions.

FIJI/ImageJ, or other image-processing software able to stitch together tile images.

Protocol steps with step annotations:

Prepare Reagents

• 1Warm complete neurobasal media to 37°C.
• 2Dilute any required pharmacological treatments, such as chemical inhibitors, into aliquots of complete neurobasal media.

Protocol steps and annotations

• 3Before beginning the experiment, aspirate old media and add 2 mL per dish of new media, including any required pharmacological treatments mixed into the new media.
• 4Allow cells to incubate for at least 30 minutes, or longer as required by relevant pharmacological treatments.
• 5
  Start the microscope and prepare the relevant hardware, including:

  1. Set up the microscope’s incubation chamber at 37°C and 5% CO₂ with humidified air. Allow sufficient time (usually 15–30 minutes) for the chamber to equilibrate before placing any dishes.
  2. Choose a phase-contrast objective, e.g., 32x.

  An objective in the magnification range of 20x-40x is typically appropriate for 3D phase-contrast live imaging. Magnification lower than 20x is sufficient for measuring velocity but makes it difficult to resolve migratory structures. Magnification higher than 40x decreases the thickness of the plane of focus, and thereby increases the probability that cells in 3D will move out of the plane of focus. Ensure that a dish is seated perfectly level, since a wide imaging area will exaggerate the effects of a tilled plate and may unintentionally result in imaging too near the top or bottom surface of the gel.

• 6After cells have completed incubation and the incubation chamber has equilibrated, retrieve the dish(es) from incubation and secure them in the incubation chamber.

  Ensure dishes are secure from moving due to motion of the motorized stage. E.g., use a small amount of dental wax to adhere a dish to the imaging tray to immobilize. Ensure that the immobilization method does not cause the dish to tilt at all.

• 7Start the microscopy software.
• 8Looking through the ocular lens, bring cells into focus near the center of the sample and inside the gel somewhere between the top and bottom surface along the z-axis.
• 9Looking through the ocular lens, visually scan the sample along the X, Y and Z axes to find a field of view that appears to have an appropriate density of cells (Fig. 3).

  The gel may be thinner at its center than at the perimeter of the inner well due to some adhesion to the sides. Be cautious about choosing a field of view at the perimeter due to increased gel thickness and a slight upward slope in the glass base as it approaches the perimeter, as these factors can degrade image clarity and require more laser power. Also note that high cell density in the field of view can obscure a clear view for easily tracking cells, so when possible, avoid areas where cells frequently overlap.

  A good guideline is to ultimately track the velocity of 15–20 cells, and remember that the number can be distributed throughout the total area of the tiled matrix. Because cells may move out of the plane of focus over the duration of the experiment, consider trying to find a density of in-focus cells that is 2–4 times the number eventually needed.

• 10
  Ensure selection of a focal plane that is near the middle of the gel for the chosen field of view.

  1. Either looking through the ocular lens or using the software’s live view, use the focus knob to move the plane of focus along the z-axis to find the approximate top and bottom bounds of the gel. Note these boundary heights in μm as stated by the software.
     Typically, imaging software displays the focal plane height via the “z-stack” feature.
  2. Calculate the approximate gel thickness as the difference in boundary heights. E.g., if software indicated that the gel top surface was at 1139 μm and bottom surface at 937 μm, the gel thickness would be 202 μm.
     Note that the boundary height values displayed by software (e.g.., 1139 and 937 μm in this example) are measurements that relate to the microscope’s focal distance; their absolute values do not have special meaning for the purpose of finding gel bounds. The numbers merely serve as relative reference points for the calculation.
  3. Calculate the mid-point plane of the gel by dividing the gel thickness by 2 and adding that number to the bottom boundary z value.
     E.g.: 202 μm / 2 = 101 μm
     937 μm + 101 μm = 1038 μm
     1038 μm is the midpoint of the gel for the chosen field of view.
     In other words, 1038 μm is the focal distance that in this example corresponds the midpoint of the gel.
  4. Use the focus knob to set the focal plane to the midpoint of the gel (e.g., 1038 μm as displayed in software) and adjust along the z-axis as needed to find a plane containing the most in-focus cell bodies that is not within ~50 μm of the top or bottom of the gel (Fig. 4). This reduces the influence that mechanical effects from glass or open surfaces may have on cell motility.
     GSCs in 3D collagen can have highly dynamic protrusions that may unpredictably extend above and below the plane of focus, while cell bodies move less rapidly and therefore are better candidates for determining where to focus for tracking cell motility. It is therefore important to maximize the number of in-focus cell bodies rather than in-focus protrusions. Also, cell bodies typically move in the direction of their protrusions, so when possible it is preferable to include cells with protrusions closer to the plane of focus, to increase the probability that cells stay close to the plane of focus for the duration of imaging.
• 11In software, set parameters to capture a matrix of tiled images over time. Consider the following parameters in particular.

  This protocol has been successfully executed using the below parameters but may require adjustment depending on your exact microscope setup and any variation in cell preparation.

  1. Scan area: Zoom out to the maximum to increase the field of view.

     A larger scan area reduces the number of tiles needed in the tiled matrix to cover the intended imaging area.

  2. Tile scan: 3 rows, 4 tiles per row (Fig. 5).

     The magnification of the objective and the density of in-focus cells in the intended capture area will influence the matrix size you choose (e.g. 3×4, 4×4, etc), where higher magnification or lower cell density require more tiles. Adjust the number of tiles and their arrangement to find the right coverage for your needs. Scan time increases as number of tiles increases, and cannot exceed the Imaging Interval.

  3. Positions: Save the XYZ coordinates as a “position” when imaging multiple samples or multiple sites in a single sample.

     This feature is also useful for temporarily saving coordinates when searching for the best field of view in a sample.

  4. Imaging interval: 5 minutes

     Adjust based on the temporal granularity you need, factoring in the anticipated speed of cells (faster cells may need a shorter interval), and whether the interval is long enough to accommodate the time the microscope requires to capture all tiles and multiple dishes. Note also that for some microscopes it is good practice to allow some idle “downtime” between intervals to prevent overheating or mechanical issues.

  5. Imaging duration: 12 hours

     Adjust imaging duration as needed. As a guideline, less than 6 hours may provide insufficient data. Greater than 12 hours increases evaporative loss of media (usually significant after 24 hours).

• 12Do a preliminary scan of the chosen field of view by activating the features described in step #11a-c, and begin imaging.
• 13Repeat steps #8–10 and #12 if the preliminary results are unsatisfactory and require iterative adjustment, or repeat for any additional imaging positions for the current or other samples.
• 14When all preliminary imaging is satisfactory, activate the features described in step #11d-e.

  When using multiple saved positions, it is good practice to check that in the time elapsed while saving positions, the chosen fields of view saved earlier have not in the meantime drifted from their positions.

• 15Begin imaging, and monitor the first few cycles of images to ensure there is no focus drift.
• 16When imaging is complete, save all data, import tiles into FIJI/ImageJ, and stitch together using the Combine feature.

  It is critical to check that tiles are aligned such that no there are no gaps over overlaps between tiles that would cause errors in measured distances. If such discrepancies exist and cannot be fixed, only make measurements within a given tile. In FIJI/ImageJ, rather than manually stitching, consider creating a stitching macro (series of automated steps).

Figure 3.

Phase-contrast image of GSCs suspended in 3D collagen. Out of focus areas indicate parts of cells that are above and below the plane of focus. Scale bar represents 50 μm.

Figure 4.

Illustration emphasizing that the plane of focus should be near the middle of the gel.

Figure 5.

Phase-contrast image of GSCs embedded in 3D collagen. Single image tiles are stitched together into a 4×3 matrix to present a contiguous field of view. Variable illumination of each image tile causes a gridded appearance after assembly. Scale bar represents 200 μm.

REAGENTS AND SOLUTIONS:

0.1% Bovine serum albumin (BSA) solution

0.1g Bovine serum albumin (BSA) (MP Bio, cat. no. 160069)

10 mL ddH₂O or Milli-Q water

0.22 μm 50 mL Steriflip vacuum filter (Millipore, cat. no. SE1M179M6) or 0.22 μm syringe filter equivalent

50 mL centrifuge tube

Combine 0.1 g BSA with 10 mL ddH₂O in a 50 mL centrifuge tube to make 1% BSA solution. Remaining steps should be done in sterile conditions. Use a 0.22 μm flip-filter (or syringe-filter equivalent) to sterilize the 1% BSA solution. In a new 15 mL tube combine 1 mL of 1% BSA solution with 9 mL of sterile ddH₂O to make 0.1% BSA solution.

0.2% BSA in PBS

0.1g Bovine serum albumin (BSA) (MP Bio, cat. no. 160069)

PBS 1X (see recipe in this section)

15 mL centrifuge tube

Dissolve 0.2 g of BSA into 10 mL of PBS 1X in a 15 mL centrifuge tube. BSA solution should be prepared fresh for each use to avoid bacterial growth. This recipe makes sufficient solution to fix and stain 4 samples using Basic Protocol 3. (See Table 6.)

Table 6.

A quick reference for making 0.2% BSA in PBS solution as needed.

Number of dishes	BSA	PBS 1X	Total volume
1	0.005 g	2.5 mL	2.5 mL
4	0.02 g	10 mL	10 mL

0.25% TritonX-100

Triton X-100 stock (Sigma-Aldrich/BioXtra, cat. no. T9284)

PBS 1X (see recipe in this section)

50 mL centrifuge tube, x2

Vortex mixer

Dissolve 400 μL of stock TritonX-100 into 39.6 mL PBS 1X in a 50 mL centrifuge tube to make a working stock of 1% TritonX-100; keeps at least 12 months at room temperature. Make a 0.25% TritonX-100 working solution by dissolving 10 mL of 1% TritonX-100 into 30 mL PBS 1X in a 50 mL centrifuge tube. Vortex stock and working solutions to facilitate complete dissolution. (See Table 7.)

Table 7.

A quick reference for making TritonX-100 solutions.

Solution	stock TritonX-100	1% TritonX-100	PBS 1X	Total volume
1% TritonX-100 in PBS	400 μL	-	39.6 mL	40 mL
0.25% TritonX-100 in PBS	-	10 mL	30 mL	40 mL

4% paraformaldehyde (PFA) in PBS:

16% Paraformaldehyde in aqueous solution (Electron Microscopy Sciences, VWR cat. no. 100503-914)

PBS 1X (see recipe in this section)

Dissolve 500 μL of 16% PFA into 1.5 mL of PBS 1X for each dish requiring fixation, all in a 15 mL conical tube.

10x DMEM

Dissolve 1 packet of DMEM powder (Sigma-Aldrich, cat. no. D2429) into 50 mL of ddH₂O or Milli-Q water, and filter via 0.22 μm 50 mL Steriflip vacuum filter (Millipore, cat. no. SE1M179M6). Aliquot in volumes of 500 μL, store at −20°C (lasts indefinitely), and thaw at 4°C (lasts 1 month).

10x Reconstitution buffer (RB)

0.88 g of NaHCO₃ (0.26M)

1.906 g of HEPES (0.2M)

40 mL of Milli-Q H₂O

Combine reagents and filter with a 0.22 μm 50 mL Steriflip vacuum filter (Millipore, cat. no. SE1M179M6). Aliquot at volumes of 500 μL and store at −20°C (lasts indefinitely), and thaw at 4°C (lasts 1 month).

Complete DMEM with FBS (7.5%)

500 mL Dulbecco’s Modified Eagle Medium without phenol red (DMEM; Cytiva, cat. no. SH30284.01)

40 mL of 100% fetal bovine serum (FBS; final 7.5%; R&D Systems, cat. no. S11150)

5 mL penicillin-streptomycin-glutamine (100X; Gibco, cat. no. 10378016)

Filter FBS with a 0.22 μm 500 mL Steri-flip filter (CELLTREAT, cat. no. 229707) before combining reagents.

Complete Neurobasal media

47.75 mL of Neurobasal-A medium (Gibco / Fisher Scientific, cat. no. 10888022); alternatively, Neurobasal-A medium minusphenol red (Gibco / Fisher Scientific, cat. no. 12349015)

500 μL of 2mM GlutaMAX supplement (100x stock; Gibco / Fisher Scientific, cat. no. 35050061)

500 μL of Antibiotic Antimycotic (100x stock; Gibco / Fisher Scientific, cat. no. 15240062)

1 mL of B27 supplement minus Vitamin A (100x stock; Gibco / Fisher Scientific, cat. no. 12587010)

50 μL of 2 mg/mL Heparin solution (2 μg/mL final) (STEMCELL Technologies Inc, cat. no. 07980)

100 μL of recombinant human basic fibroblast growth factor, reconstituted at 10 μg/mL (154 a.a., bFGF, 20ng/mL final) (PeproTech, cat. no. 100–188)

100 μL of recombinant human epidermal growth factor, reconstituted at 10 μg/mL (HB-EGF, 20 ng/mL final) (PeproTech, cat. no. 100–47)

0.1% bovine serum albumin (BSA) (see recipe in this section)

Tabletop centrifuge

Biological safety cabinet

P1000 and P100 pipettes and tips

Automated pipette with serological pipet tips

0.2 mL PCR tubes

1.5 mL centrifuge tubes

15 mL conical tubes

50 mL conical tube

Reconstitute bFGF and EGF: Briefly (~5 seconds) spin down the stock vial of growth factor to ensure powder moves to the bottom for resuspension. In sterile conditions, resuspend 100 μg of bFGF in the manufacturer’s vial using 500 μL of 0.1% BSA solution. Transfer the 500 μL of reconstituted bFGF into a 15 mL conical tube containing 9.5 mL of 0.1% BSA solution for a total of 10 mL of bFGF in 0.1% BSA, and gently mix by trituration. Do not vortex growth factors. Aliquot the bFGF in 0.1% BSA solution as 110 μL portions (to allow for pipetting error) into 0.2 mL pcr tubes. A stock vial of 100 μg makes about 90 aliquots and the process takes about 60–90 minutes. Repeat the preceding steps for reconstitution of EGF.

Aliquot GlutaMAX, Antibiotic Antimycotic, B-27 and Heparin: Aliquot these reagents into volume-appropriate 1.5 mL or 0.2 mL tubes for ease of use and to avoid subjecting reagents to additional freeze-thaw cycles. Store at indicated temperatures.

Assemble media: Add reagents to a conical tube as listed in Table 8, scaling the volumes up or down as needed. Once assembled, Complete Neurobasal media maintains full activity for 1 week due to the decline in biological activity of bFGF and EGF thereafter. It is therefore recommended that new media be made after 1 week, although some labs are known to extend this period up to 2–3 weeks. Note: for transfections, Antibiotic Antimycotic may reduce transfection efficiency and therefore may need to be omitted from the media. In this instance, replace the lost volume with Neurobasal-A media.

Table 8.

Example recipe for 50 mL aliquot of Complete Neurobasal Media

Reagent	Amount	Final concentration	Stock	Stock storage (°C) and expiration
Neurobasal-A	47.75 mL	1x	500 mL bottle, 1x	4 (12 months; protect from light)
GlutaMAX	500 μL	1x	100 mL bottle, 100x (200mM)	room temperature, 4 or −20 (24 months)
Anti Anti	500 μL	1x	100 mL bottle, 100x	−20 to −5 (12 months)
B-27	1 mL	1x	10 mL bottle, 50x	−20 to −5 (12 months)
Heparin	50 μL of 2mg/mL	2 μg/mL	2 mL vial, 2 mg/mL	4 (date given, but generally ~18 months)
bFGF	100 μL	20 ng/mL	100 μg powder	−20 or −80 (<12 months) / 4 (1 week)
EGF	100 μL	20 ng/mL	100 μg powder	−20 or −80 (<12 months) / 4 (1 week)

NaOH solution

0.40 g NaOH pellet

10 mL ddH2O or Milli-Q water

Dissolve NaOH into water and sterilize using a 0.22 μm filter. Aliquot at volumes of 500 μL and store at −20°C (lasts indefinitely).

PBS 1X

1 PBS tablet (ph 7.4; Millipore, cat. no. 524650)

1 L of ddH₂O or Milli-Q water

Autoclave to sterilize after preparation.

COMMENTARY:

Background Information

Collagen gels have been used to study cell biology for many years, and more recently have been used to study the molecular mechanisms underlying migration of glioma cells (Cockle et al., 2015; Fayzullin et al., 2019). The protocols presented here describe how to use 3D collagen matrices to discover new cellular and molecular biology underpinning glioma cell dynamics. The protocols also detail techniques for culturing and monitoring differentiation of glioma stem cells for use in an in vitro system. Glioma stem cells more closely recapitulate the original tumor than clonal lines and thus are especially useful for studying molecular mechanisms (Chen et al., 2012). The in vitro collagen model does not reproduce the biochemical composition of the in vivo microenvironment, but is more accessible, cost effective, easier to image, and critically can represent important physiological structural features. To investigate biochemical factors, consider as an alternative or in addition to the collagen model using decellularized brain tissue or live brain slices.

Critical Parameters:

Culturing glioma stem cells as neurospheres

Patient derived GSC lines are known to vary in growth rate, for example doubling every 2 days to every 14 days, due to intertumoral heterogeneity that is inherent to gliomas (Grube et al., 2021). It may therefore be necessary to adjust the number of cells recommended for seeding (Table 1) depending on the growth rate you observe. When dissociating GSC neurospheres, the risk of cell death rises in proportion to longer exposure to Accutase and more vigorous trituration. Cells generally suffer no ill effects from Accutase exposure up to 10 min. However, finding the right trituration technique can take practice: too gentle fails to mechanically break apart cells, but too much shear force damages cell membranes. An indication for the need to triturate more firmly is if, after 8–9 minutes of suspension in Accutase, spheres remain significantly large or there is little to no cloudiness. Two factors that increase shear force are higher velocity of fluid flow and smaller diameter tips; try adjusting these factors to optimize dissociation. When practicing the right level of trituration, err on the side of too gentle, and recognize that it may not be possible to fully dissociate very large spheres without some cell death (though the cores of such spheres may already be dead due to hypoxia). It is also important to note that different cell lines can vary in sensitivity to enzymatic and mechanical factors that will influence your technique and rate of dissociation.

Adherent GSC growth rate and differentiation medium

Cell growth rate varies by patient derived line, as previously mentioned. Consider this factor when determining the number of cells to plate, to avoid confluent dishes before the 2-week time course ends. DMEM with serum is typically used as a differentiation medium (Ledur et al., 2017), but because neurobasal medium is better suited to neural cells, you may opt to differentiate cells by adding 7.5% FBS serum to complete neurobasal medium without bFGF, EGF and heparin (Zepecki et al., 2021).

Collagen architecture

Altering different parameters of collagen could impact cell behavior and morphology. Collagen can be modified, for example, by increasing concentration to increase stiffness and reduce porosity, or by polymerizing at different temperatures to alter architecture (Doyle, 2016). Significant alterations to the gel architecture may require that cells adopt differing modes of migration or morphologies for navigation. Use the tunability of collagen to explore aspects of cell motility or troubleshoot issues where cells may be poorly matched to collagen architecture.

Plating density in 3D collagen

Proper plating density will critically determine imaging results. Too many cells will make it difficult to clearly distinguish individual cell bodies and especially protrusions since cells readily overlap in a 3D field. Plating too few cells may be solved by adding tiles to the imaging matrix, but the resulting increased imaging time may extend longer than a desired Imaging Interval.

Troubleshooting:

Table 9 summarizes the most common technical issues we encountered, organized by protocol.

Table 9.

Troubleshooting guide.

Protocol	Problem	Possible Cause	Solution
Basic Protocol 1	Cells fail to properly dissociate	Too much supernatant remains before Accutase is added, which dilutes/stops enzymatic activity.	Try to remove more supernatant without removing cells.
		Neurospheres are too large to easily disassociate.	Grow neurospheres to a smaller size for easier dissociation
		Mechanical dissociation is too gentle.	Either triturate more frequently, firmly, or try a smaller diameter tip.
Basic Protocol 2	Cells approach confluence before the end of 14 days	Cell proliferation rates vary by patient-derived line. Your cells may grow faster than expected compared to the example provided in the protocol.	Plate fewer cells in dishes with longer incubation periods. E.g., 20,000 cells for week 1 dishes, 15,000 for week 2 dishes.
	High cell density in center of dish	After plating, when physically transporting dishes to incubation, rocking or swirling motions can cause suspended cells to aggregate in the center of the dish, where they settle.	When transporting cells, minimize any movements that may cause significant motion in cell media. Do not intentionally create motion in an attempt to redistribute cells, as this may have the opposite effect.
	Cell edges appear folded over, as indicated by very bright staining	Cell edges lifted during the fixing and staining process.	Be careful when ejecting reagents from the pipette to avoid directly hitting cells (eject onto the inner wall of the dish) or eject with less velocity.
	Background fluorescence	Fluorescent antibodies were incompletely washed away.	Do one or more additional washes with PBS.
		Contamination, most evident by positive DAPI staining.	Isolate the source of contamination. Contamination could originate from inadequate sterile technique or potentially have been introduced at some point in the process of isolating GSCs from a patient sample.
Basic Protocol 3	Collagen texture appears patchy or uneven (Fig. 6)	Collagen solution began to warm above 4°C before completing application to the dish.	Quickly mix and apply collagen solution, keeping everything on ice for the duration. Moving quickly is important not only to ensure integrity of collagen but also to avoid prolonged cell exposure to cold temperatures.
		A reagent in the collagen recipe needs replacement.	Though 10x RB and 10x DMEM should keep indefinitely at −20°C, they may alter after a long time, or when kept at 4°C for an extended period. It’s also possible for the pH of NaOH to drift. Remake these solutions. If replacement fails, next try new collagen.
Support Protocol 2	Microscope can’t resolve surface of gel	The gel is too thick for the working distance of the objective to focus on the gel surface (i.e. working distance is too short).	Use a wider objective or a lower volume of collagen solution to make a thinner gel.
	Adjacent tiles, when combined, have gaps or significant overlap	The microscope is misaligned.	A technician should realign the instrument. If this is not possible, only measure the dynamics of a cell within a single tile rather than moving between tiles.

Understanding Results:

Neurospheres that are ready to passage should appear similar to those in Fig. 1 under 400 μm. Occasionally spheres may adhere to each other and grow together as a single non-spherical mass. Neurospheres that appear to be excessively loose or in the process of disaggregating indicate unhealthy cells; check for contamination and that reagents are not expired. To minimize risk of contamination, all cell culture should be done using sterile technique, using safeguards such as filtered pipette tips. If cells are newly thawed, cells that appear unhealthy may not have survived the freezing process rather than being contaminated. If several days after passaging the majority of cells appear as single cells rather than nascent spheres, trituration may have been too forceful and harmed the cells. Expect that some larger spheres may not fully disaggregate during passaging and may appear immediately afterward in culture as small, disorganized clumps. These clumps will seed future neurospheres.

Differentiating cells should significantly enlarge over 14 days and vimentin networks should become less expansive and concentrate around the nucleus. In addition or as an alternative, consider western blotting or immunofluorescent staining for proteins that have been previously suggested to indicate glioma cell stemness, such as CD133, nestin, Nanog and Sox2 (Zepecki et al., 2019). However, note that there is currently no definitive single or combination of markers of GSC stemness (Prager et al., 2020).

GSCs in 3D collagen should exhibit similar morphology and velocity for the duration of imaging. Obvious changes over time suggest that an environmental factor (e.g., altered temperature, CO₂, contamination, etc) is affecting the cells. A cell density as shown in Fig. 5, captured in a 3×4 tiled matrix, should yield 15–18 cells that stay close to the plane of focus over a 12 hr period. It is normal for most cells to appear out of focus.

Time Considerations:

Passaging GSC neurospheres

The procedure takes 35–45 min.

Plating and staining GSCs to monitor differentiation

The procedure for plating cells takes 45–60 min, and the procedure for fixing and staining before imaging takes ~3.25 hr for a single dish, with an additional 5 min for each additional dish. To save time staining, consider fixing cells at days 1, 3 and 6, and staining them together on day 6. Similarly, group fixations from days 10 and 14 and stain on day 14. While the best time to stain and image is immediately after fixation, the grouped staining strategy is reasonable when time efficiencies take priority.

Coating dishes with fibronectin

The procedure takes ~75 min, but 60 min of that time is just incubation.

Embedding GSCs in a 3D collagen matrix

Calculations and preparation take about 20 min. However, if using silanized dishes but none are prepared, allow a half-day to silanize dishes and allow them to dry overnight (see Doyle, 2009). If your experimental plan requires using many silanized dishes, it is advantageous to silanize 20 or more dishes at once and store them in a storage bin with desiccants. Cell preparation takes ~45 min; preparation of the collagen solution takes ~15 min for a single dish, plus <5 min for each additional dish, and 45–60 min of incubation.

Live imaging cells in 3D matrices

Time depends on your microscope setup, the microscope’s imaging speed, user experience, number of positions (within and/or between conditions), and whether time is required for drug treatments. The least predictable steps with regard to time are those that involve locating a suitable field of view. In general, for 1 position allow 60–90 min for microscope setup and all preliminary imaging, and for 4 positions allow up to 3 hr if everything moves slowly.

Figure 6.

Phase-contrast images of the surface of 3D collagen matrices with embedded GSCs. (A) Collagen fibers appear uniformly distributed, as expected. (B) Collagen appears patchy and non-uniform, indicating that a problem occurred at some step of gel creation. See the Table 9 troubleshooting guide for suggestions. Scale bars represent 50 μm.

DATA AVAILABILITY STATEMENT:

Data is available upon request.