Protocol for generation of multicellular spheroids through reduced gravity

Summary

Multicellular spheroids are useful models for drug testing or studying tumor biology, but their production requires specialized approaches. Here, we present a protocol to produce viable spheroids by slow rotation around a horizontal axis using standard culture tubes. We describe steps for both seed and starter culture, and maintenance and expansion of spheroids. We detail assessment of spheroid size, count, viability, and immunohistochemistry. This protocol reduces gravitational forces that lead to cell clumping and is amenable to high-throughput use.

Graphical abstract

Highlights

• •Generation of multicellular spheroids from malignant and non-malignant cells
• •Method to reduce gravitational forces to form spheroids in suspension
• •Steps to use seed and starter cultures to generate viable spheroids
• •Steps to expand and maintain spheroids in standard culture tubes

Publisher’s note: Undertaking any experimental protocol requires adherence to local institutional guidelines for laboratory safety and ethics.

Multicellular spheroids are useful models for drug testing or studying tumor biology, but their production requires specialized approaches. Here, we present a protocol to produce viable spheroids by slow rotation around a horizontal axis using standard culture tubes. We describe steps for both seed and starter culture, and maintenance and expansion of spheroids. We detail assessment of spheroid size, count, viability, and immunohistochemistry. This protocol reduces gravitational forces that lead to cell clumping and is amenable to high-throughput use.

Before you begin

Overview/introduction

The use of cells cultured in monolayers is a conventional approach to the study of cell biology. Three-dimensional (3-D) cell culture and the generation of multicellular spheroids (MCS) have distinct advantages over these approaches as they can incorporate cellular and stromal interactions and enable cell growth within a structure that better recapitulates tissue environments. When generated from tumor cells from patient-derived samples or from established cancer cell lines, MCS are suitable models for drug screening or for studying tumoral biological processes involved in cell invasion, metastases or angiogenesis. However, the morphology of MCS and the phenotypic behavior of cells can be influenced by the methods used to prepare the spheroids. There are several different methods for generation of MCS. Commonly used techniques for cell culture include the use of ultra-low adherence plates, spinner flasks, rotating wall vessels, microfluidic devices, scaffold/matrix models, hanging drop cultures and magnetic levitation.^1,2,3,4,5,6 Each of these methods has limitations, and some are not only labor intensive but lack consistency in size or scalability that preclude their broader use for drug screening (Figure 1). Herein we describe a simplified protocol for spheroid formation from diverse cell types in suspension that enables intercellular interactions to proceed by reducing gravitational effects. A clinostat is used to provide a slow rotation around a horizontal axis that reduces the effects of gravitational forces and prevents cells from settling. This method provides a simple and scalable process for cell spheroid formation. The use of this protocol results in the generation of MCS with consistent size and shape, and that avoids several of the limitations observed with other methods.

Figure 1.

Comparison of methods for spheroid generation

A comparison of features related to generation and yield is depicted for each. +++ high, ++ moderate, + low, - not applicable.^1,2,3,4,5,6

This protocol describes the generation of non-malignant and malignant cell based MCS. Primary human hepatocytes (HH) were used for generating non-malignant spheroids. Hepatocellular carcinoma cell lines (HepG2 and Hep3B) and cholangiocarcinoma cell lines (SNU-1079 and HuCCT1) were used to develop malignant cell spheroids. Supporting cell populations such as immortalized LX2 human hepatic stellate cells (HSC), primary human dermal fibroblasts (HDF), and human umbilical vein endothelial cells (HUVEC) were used to create organ relevant spheroid models that mimic in vivo cell-cell interactions occurring within tissues.

The following cell combinations are described in the subsequent sections:

• Primary human hepatocytes + hepatic stellate cells (HH-HSC) + fibroblasts (HH-SF) + endothelial cells (HH-SFE).

• Cancer cells + stellate cells (CS) + fibroblasts (CSF) + endothelial cells (CSFE).

This protocol is not limited to the cell types described and can be adapted for use with other types of cells.

Cells and cell culture

Timing: 1 h

• 1.
  Culturing and passaging human malignant cells: HepG2, Hep3B, SNU-1079, and HuCCT1.

  • a.
    Make collagen-1 coating solution as described in the materials and equipment section.
  • b.
    Add 7 mL of collagen-1 coating solution to cover bottom of 10 cm cell culture dish, let sit for 20 s and remove.
  • c.
    Leave dish uncovered in sterile tissue culture hood for 1 h to dry.
  • d.
    Plate 6 × 10⁵ cells/dish and passage when cells reach 70%–80% confluency.
• 2.
  Culturing and passaging non-malignant human hepatocytes (HH).

  • a.
    Make poly-L-lysine coating solution as described in the materials and equipment section.
  • b.
    Add 8 mL of poly-L-lysine coating solution to a 10 cm cell culture dish.
  • c.
    Incubate for at least 1 h at 37°C.
  • d.
    Remove coating solution and wash dish with 8 mL of sterile H₂O.
  • e.
    Plate 6 × 10⁵ cells /dish and passage when cells reach 80%–90% confluency.
• 3.
  Culturing and passaging HSC cells.

  • a.
    Coat 10 cm cell culture dish with collagen-1 solution following step 1 (a–c).
  • b.
    Plate 7 × 10⁵ cells/dish and passage at 70%–80% confluency.
• 4.
  Culturing and passaging HDF.

  • a.
    No special pretreatment of culture dish required.
  • b.
    Plate 6 × 10⁵ cells passage at 80% confluency.

Note: HDF’s are slow growing. If large numbers of cells are required, it is recommended to plate up to 8 × 10⁵–1 × 10⁶ cells.

• 5.
  Culturing and passaging HUVEC.

  • a.
    Coat plate with collagen-1 coating solution following step 1(a–c).
  • b.
    Plate 7 × 10⁵ cells/dish and passage at 80%–90% confluency.

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
Anti-alpha smooth muscle actin antibody (α-SMA) (1:500)	Abcam	Cat# ab5694
CD31 Monoclonal antibody (SP38) (1:50)	Thermo Fisher Scientific	Cat# MA5-16337
Chemicals, peptides, and recombinant proteins
Dulbecco’s Modified Eagle Medium (DMEM) high glucose	Cytiva	Cat# SH30243.01
Fetal bovine serum (FBS)	Gemini Bio	Cat# 100-106
Penicillin/streptomycin solution	Gemini Bio	Cat# 400-109
Hepatocyte Medium Kit	ScienCell	Cat# 5021
(Above Kit) Hepatocyte Medium	ScienCell	Cat# 5201b
(Above Kit) Fetal bovine serum (FBS)	ScienCell	Cat# 0025
(Above Kit) Penicillin/streptomycin solution	ScienCell	Cat# 0503
(Above Kit) Hepatocyte growth supplement	ScienCell	Cat# 5252, 5021
Poly-L-Lysine	ScienCell	Cat# 0403
Fibroblast basal medium	ATCC	Cat# PCS-201-030
Fibroblast Growth Kit-Low serum	ATCC	Cat# PCS-201-041
(Above Kit) Ascorbic acid	ATCC	Cat# PCS-999-006
(Above Kit) Fetal bovine serum (FBS)	ATCC	Cat# PCS-999-010
(Above Kit) rh Insulin	ATCC	Cat# PCS-999-022
(Above Kit) rh FGF-b	ATCC	Cat# PCS-999-020
(Above Kit) L-Glutamine	ATCC	Cat# PCS-999-016
(Above Kit) Hydrocortisone hemisuccinate	ATCC	Cat# PCS-999-014
Dulbecco’s Modified Eagle Medium (DMEM) F12	Gibco	Cat# 10568-018
Heparin	Sigma-Aldrich	Cat# H4784
Endothelial Cell Growth Support	Corning	Cat# CB40006
Collagen-1 rat tail	Corning	Cat# 354236
Acetic Acid, Glacial	Thermo Fisher Scientific	Cat# A38-500
0.25% Trypsin/EDTA	Gibco	Cat# 25200-056
Calcein-AM	Invitrogen	Cat# C1430
Propidium iodide	Thermo Fisher Scientific	Cat# R37169
Phosphate buffered saline (PBS)	Cytiva	Cat# SH30256.02
10% Formalin	Thermo Fisher Scientific	Cat# SF100-4
HistoGel	Epredia	Cat# HG4000012
TBS with Tween™ (TBST), 20× solution	Thermo Fisher Scientific	Cat# J77500.K2
Target retrieval solution, citrate pH 6.1 (10×)	Agilent	Cat# S169984-2
Protein block serum-free	Agilent	Cat# X0909
EnVision + single reagent (HRP. Mouse)	Agilent	Cat# K400111-2
EnVision + single reagent (HRP. Rabbit)	Agilent	Cat# K400311-2
DAB+	Agilent	Cat# K346811-2
Hematoxylin solution, Gills No. 1	Sigma-Aldrich	Cat# GHS132
Hydrogen peroxide blocking reagent	Abcam	Cat# ab64218
Experimental models: Cell lines
Primary human hepatocytes (HH)	ScienCell	Cat# 5200
LX2, human hepatic stellate cells	Provided by Dr. GJ Gores	Mayo Clinic Rochester
Primary human dermal fibroblasts normal, adult	ATCC	Cat# PCS-201-012
Human umbilical vein endothelial cells	ATCC	Cat# CRL-1730
HepG2, hepatocellular cancer cells	ATCC	Cat# HB-8065
Hep3B, hepatocellular cancer cells	ATCC	Cat# HB-8064
SNU-1079, cholangiocarcinoma cells	Provided by Dr. J. Copland	Mayo Clinic Florida
HuCCT1, cholangiocarcinoma cells	Provided by Dr. GJ Gores	Mayo Clinic Rochester
Software and algorithms
ImageJ Fiji (2.3.0; Java 1.8.0_322)	Schindelin et al.7	https://imagej.nih.gov/ij/
GraphPad Prism (9.2.0(332))	GraphPad	Prism - GraphPad
Other
10 cm cell culture dish	Thermo Fisher Scientific	Cat# 08-772-22
1.5 mL microcentrifuge tubes	Thermo Fisher Scientific	Cat# 05-408-129
15 mL conical tubes	Falcon	Cat# 352196
TempAssure PCR pull-apart 8-tube strips attached individual optical caps	USA Scientific	Cat# 1402-3900
Vacuum driven filter 0.22 μm	Genesee Scientific	Cat# 25-227
Pipette tips (10 μL, 250 μL, 1,000 μL)	Mettler Toledo	Cat# 30389175, 30389186,30389166
Disposable serological pipettes (5 mL, 10 mL, 25 mL)	Flacon	Cat# 357543, 357551, 357525
Nunc Lab-Tek II Chamber Slide System	Thermo Fisher Scientific	Cat# 154526
Fisherbrand™ Disposable Base Molds	Thermo Fisher Scientific	Cat# 22363552
Countess Cell Counting Chamber Slide	Thermo Fisher Scientific	Cat# C10228
Countess 3	Thermo Fisher Scientific	Cat# AMQAX2000
Microscope (EVOS M5000)	Thermo Fisher Scientific	Cat# AMF5000
SpinLogic BioReactor with Tachometer Control Unit	RPS Research	N/A
CO2 cell culture incubator	Thermo Fisher Scientific	N/A
Biosafety cabinet	NUAire	N/A

Materials and equipment

DMEM high glucose medium

Reagent	Final concentration	Amount
Fetal Bovine Serum	10%	50 mL
Penicillin/Streptomycin Solution	1%	5 mL
DMEM high glucose basal media	N/A	500 mL
Total		555 mL

Store at 4°C for up to 30 days.

Human Dermal Fibroblast medium

Reagent	Final concentration	Amount
Fetal Bovine Serum	2%	10 mL
rh FGF basic	5 ng/mL	0.5 mL
Ascorbic acid	50 μg/mL	0.5 mL
Hydrocortisone hemisuccinate	1 μg/mL	0.5 mL
rh Insulin	5 μg/mL	0.5 mL
L-glutamine	7.5 mM	18.75 mL
Fibroblast Basal Medium	N/A	480 mL
Total		510.75 mL

Store at 4°C for up to 30 days.

Human Umbilical Vein Endothelial Cells Medium

Reagent	Final concentration	Amount
Fetal Bovine Serum	10%	50 mL
Heparin (50 mg/mL)	0.09 mg/mL	1 mL
ECGS (30 mg/mL)	0.03 mg/mL	0.5 mL
DMEM F12	N/A	500 mL
Total		551.5 mL

Store at 4°C for up to 30 days.

Primary Human Hepatocyte Medium

Reagent	Final concentration	Amount
Fetal Bovine Serum	10%	50 mL
Penicillin/Streptomycin Solution	1%	5 mL
Hepatocyte Growth Supplement	1%	5 mL
Hepatocyte medium	N/A	500 mL
Total		560 mL

Store at 4°C for up to 30 days.

HH-HSC culture medium

Reagent	Final concentration	Amount
HH Media	N/A	5 mL
DMEM high glucose	N/A	2 mL
Total		7 mL

Store at 4°C for up to 30 days.

HH-HSC-HDF culture medium

Reagent	Final concentration	Amount
HH Media	N/A	5 mL
DMEM High Glucose	N/A	2 mL
HDF Media	N/A	2 mL
Total		9 mL

Store at 4°C for up to 30 days.

HH-HSC-HDF-HUVEC culture medium

Reagent	Final concentration	Amount
HH Media	N/A	5 mL
DMEM High Glucose	N/A	2 mL
HDF Media	N/A	2 mL
HUVEC Media	N/A	2 mL
Total		11 mL

Store at 4°C for up to 30 days.

CS Medium (Cancer cells)

Reagent	Final concentration	Amount
DMEM High Glucose	N/A	10 mL
Total		10 mL

Store at 4°C for up to 30 days.

CSF Mixed Medium (Cancer cells)

Reagent	Final concentration	Amount
DMEM High Glucose	N/A	7 mL
HDF Media	N/A	2 mL
Total		9 mL

Store at 4°C for up to 30 days.

CSFE Mixed Medium (Cancer cells)

Reagent	Final concentration	Amount
DMEM High Glucose	N/A	7 mL
HDF Media	N/A	2 mL
HUVEC Media	N/A	2 mL
Total		11 mL

Store at 4°C for up to 30 days.

Collagen-1 coating solution

Reagent	Final concentration	Amount
Acetic acid (17.4 M)	0.017 M	0.25 mL
Rat tail collagen 1 (3.89 mg/mL)	0.06 mg/mL	3.94 mL
ddH2O	N/A	250 mL
Total		254.19 mL

Poly-L-lysine coating solution

Reagent	Final concentration	Amount
Poly-L-lysine (1 mg/mL)	0.009 mg/mL	70 μL
ddH2O	N/A	8 mL
Total		8.07 mL

Calcein-AM/Propidium iodide stain (Live/Dead)

Reagent	Final concentration	Amount
PBS	N/A	1 mL
Calcein-AM (1 mg/mL)	0.0005 mg/mL	0.5 μL
Propidium iodide (10 mg/mL)	0.005 mg/mL	0.5 μL
Total		1.001 mL

Step-by-step method details

Generation of seed cultures

Timing: 1–2 h

The purpose of this step is to facilitate the cell-to-cell contact between the cell lines and initiate spheroid formation. The following describes the preparation of HH-HSC, CS, CSF, and CSFE spheroids from culture plates as they are combined to form a seed culture.

• 1.Remove culture media from cells in cell culture plates.
• 2.Wash each plate with 7 mL of PBS and remove.
• 3.Add 2 mL of 0.25% trypsin/EDTA to each plate.
• 4.Incubate at 37°C for 3–5 min, until fully detached.
• 5.Add 8 mL of complete media to each plate to neutralize trypsin. Collect cell suspensions in individual 15 mL conical tubes.
• 6.Centrifuge cell suspensions for 5 min at 300 × g and 4°C.
• 7.Remove and discard supernatant and resuspend cell pellets in 2 mL of respective complete media by gently pipetting up and down.
• 8.Combine 10 μL of resuspended cell solution with 10 μL of trypan blue for a total of 20 μL.
• 9.Pipette 10 μL of the solution from step 8 onto a cell counting chamber slide and count cells using an automated cell counter.

Alternatives: Count using a manual hemocytometer.

• 10.
  Preparation of seed cultures.

  • a.
    HH-HSC or CS spheroids.

    • i.
      Required cell numbers 1 × 10⁵ for HH or cancer cells and for 4 × 10⁴ LX2 cells.
    • ii.
      Combine HH with LX2 cells and combine cancer cells with LX2 cells in individual 1.5 mL tubes with a volume of 500–600 μL.
  • b.
    HH-SF or CSF spheroids.

    • i.
      Required cell numbers 1 × 10⁵ for HH or cancer cells and 4 × 10⁴ for LX2 and HDF cells.
    • ii.
      Combine HH or cancer cells with LX2 and HDF cells in a 1.5 mL tube with 500–600 μL media.
  • c.
    HH-SFE or CSFE spheroids.

    • i.
      Required cell numbers 1 × 10⁵ for HH or cancer cells and 4 × 10⁴ for each LX2, HDF, or HUVEC cells.
    • ii.
      Combine HH or cancer cells with LX2, HDF, and HUVEC in a 1.5 mL tube with 500–600 μL media.
• 11.Centrifuge cell suspensions in the 1.5 mL tubes for 5 min at 300 × g at 4°C.
• 12.Prepare mixed culture media as appropriate for cell types and combinations (HH-HSC, HH-SF, HH-SFE, CS, CSF, and CSFE) as outlined in the materials and equipment section.
• 13.Remove and discard media without disturbing the cell pellet.
• 14.Resuspend cells in 60 μL of mixed media from step 12 and transfer to 0.2 mL microfuge tube.

Generation of starter cultures

Timing: 1–2 h

The purpose of this step is to initiate spheroid formation from the cells combined in the seed cultures.

• 15.Place tubes onto clinostat in a 37°C incubator.

Alternatives: Any device that rotates and prevents cells from settling will be effective. Thus, machines such as a random positioning machine (RPM) or a low-speed tissue culture rotator can be used. When rotating strictly around a single horizontal axis, samples should be placed as close to the center axis as feasible to induce a simulated microgravity effect and avoid centrifugal forces.

• 16.Set the clinostat to 18 RPM and allow it to run for 90 min.

CRITICAL: A low speed of rotation is critical for the reduction of gravitational forces by averaging out these forces without inducing centrifugal forces that could lead to aggregation. Depending on the device and the diameter of the cell vessel that is used, the speed or rotation can range from 8-40 RPM. The residual gravity force can be calculated and RPMs adjusted as needed, using the formula a = ω² ∗r where ω is the rotation velocity (in rad/s) and r is the inner radius of the cell vessel (in m).⁸

• 17.Remove tubes from the clinostat and add 100 μL of spheroids specific mixed media to the tubes.
• 18.Vortex tubes for 5 s.

CRITICAL: This step is a critical step that is needed to produce uniform and evenly sized spheroids. For validation of spheroid quality refer to “Image and analysis of spheroids” section.

CRITICAL: Ensure that cells remain well suspended after vortexing. If cell aggregation occurs and cells are allowed to settle at this step only a small number of spheroids will form.

• 19.Fill tubes completely with mixed media and return to the clinostat.
• 20.Set clinostat to 18 RPM and allow to run 15–18 h.

Expansion and maintenance of spheroid cultures

Timing: 0.5 h

The purpose of this step is to promote the growth of a large number of spheroids by increasing the available growth area and nutrients. Upon completion the spheroids have developed for 72 h and can be used for experiments as desired or continue to be maintained in culture.

• 21.Remove tubes from the clinostat.

Note: For validation of spheroid quality refer to “Image and analysis of spheroids” section.

• 22.Using sterilized scissors cut the top of 200 μL pipette tips and transfer spheroids into 1.5 mL tubes and fill them completely with appropriate media.

CRITICAL: Cutting pipette tips increases the opening diameter which allows for spheroids to be transferred without causing physical disruption.

• 23.Return tubes to the clinostat at 18 RPM and allow it to run for 24 h.

Maintenance

• 24.Remove tubes from the clinostat.
• 25.In the sterile cell culture hood place samples in a tube holder. Allow spheroids to settle by gravity to the bottom of the tube for 20–40 s.
• 26.Carefully remove 1.3 mL of media from tube and replace the same volume with fresh complete media.
• 27.Manually ensure that spheroids are suspended through rotation of the tube.
• 28.Place tubes back into tube holder and set the clinostat to 18 RPM allow it to run for 24 h.

Imaging and analysis of spheroids

Timing: 1–2 h

The purpose of this step is to confirm the successful incorporation of the different cell types and to verify the overall viability of the spheroid.

Ongoing tracking of spheroid development

• 29.Remove tubes from clinostat and bring to bright field microscope.

Note: At any desired time during the 72-hour period tubes can be removed from the clinostat and imaged by light microscopy to ensure that spheroid development is proceeding as desired (Figures 2A–2F). Spheroids can be retrieved from the tubes at any time for assessments of total count, viability, or immunohistochemical testing. A simple bar clamp could be used to grasp the end of the tube (Figure 2B).

Alternatives: Locking wrench, locking pliers, or tape. In general, any tool that can secure the tube over the objective can be used to produce a stable image.

• 30.Position the tube over the objective and acquired desired images (Figure 2C).

Note: The spheroids at this stage are prone to clumping together. To help mitigate this, it is recommended to always keep tubes laid out parallel to the surface of the work area. If multiple tubes are being imaged at the same time, consider splitting up the tubes into two working groups.

Figure 2.

Imaging process

(A–F) (A) Tubes mounted on clinostat, (B) tube in bar clamp, (C) and tube setup for imaging on microscope. Spheroid formation of HH-HSC cells at 18 h (D), 48 h (E), and 72 h (F). Scale bar = 750 μm.

Imaging for total count and yield assessment

• 31.Remove tubes from the clinostat.
• 32.In the sterile cell culture hood place samples in a tube holder. Allow spheroids to settle by gravity to the bottom of the tube for 20–40 s.
• 33.Carefully remove the supernatant and replace it with 500 μL of PBS.
• 34.Using a cut 1,000 μL pipette tip transfer the spheroids into one chamber of a four-chamber microscope slide.

Alternatives: A non-chambered microscope slide can be used in place of a chamber microscope slide; however, a smaller volume of PBS (300 μL) should be used to avoid loss.

• 35.Image spheroids using a bright field microscope.

Note: We suggest aiming to capture all spheroids in the sample for an accurate total count. Ensure that edges of spheroids are clearly defined otherwise if spheroids are overlapping accurate measurements will not be possible. Imaging may require capture of multiple fields of view.

• 36.Save Images as .tiff files and transfer them to a computer with access to ImageJ Fiji image processing software (open-source software).

Note: When saving images ensure that the scale bar is included in the image. The scale bar will be used for size measurements in the ImageJ Fiji software.

• 37.Using a cut pipette tip, transfer and divide spheroids from the chamber slide evenly into two 1.5 mL tubes.

Viability assessment

• 38.Allow spheroids to settle to the bottom in one of the 1.5 mL tubes.
• 39.Remove PBS.
• 40.Incubate spheroids in 300 μL of calcein-AM/propidium iodide solution for 4 min. Refer to materials and equipment section for recipe.

CRITICAL: Spheroid fluorescence imaging should be completed promptly after 4-minute incubation time, if staining is continued for significantly longer, the spheroids may become overstained and can no longer be interpreted, this is particularly crucial for propidium iodide stain.

• 41.Image spheroids via bright field microscopy and fluorescence microscopy.

Note: For fluorescence microscopy use an excitation wavelength of 482nm (GFP channel) to excite calcein-AM and 542nm (RFP channel) for propidium iodide.

• 42.Save images as individual files (Bright field, GPF, RFP) and merged files (Figure 3).

Figure 3.

Viability assessment of multicellular spheroids

(A–C) HH-HSC spheroids (A), HH-SF spheroids (B), and HH-SFE spheroids (C) were generated and analyzed after 72 h. Brightfield, calcein-AM (live cells), propidium iodide (dead cells), and the merged images are shown. The spheroids largely comprise of viable cells with only a small number of dead cells. Scale bar = 300 μm.

Immunohistochemistry

• 43.Allow the spheroids in the second 1.5 mL tube (from step 37) to settle to the bottom.
• 44.Remove PBS.
• 45.Add 300 μL of 10% formalin to the tube and incubate at 20°C–22°C for at least 24 h.

Note: The solution is toxic and required safety laboratory procedures for storage should be observed.

• 46.Ensure that spheroids are settled to the bottom of the tube and carefully remove the properly discard 10% formalin.
• 47.Wash with 500 μL of PBS.
• 48.Warm HistoGel using water bath or microwave as per manufacturer.
• 49.Allow spheroids to fully settle and remove PBS.
• 50.Add 50 μL of warmed HistoGel to the tube.
• 51.Using a cut pipette tip transfer HistoGel containing spheroids to disposable histology mold and place spheroids in a small bead.
• 52.Let slightly solidify at 20°C–22°C for 1–2 min.
• 53.Fill the rest of the mold with warm HistoGel and place on ice for 5 min.
• 54.Remove solidified HistoGel from the mold and place in tissue cassette.
• 55.Keep tissue cassette in 70% Ethanol.
• 56.Perform sample paraffinization, deparaffinization, and rehydration as per standard in-house protocol.
• 57.Retrieve antigen using target retrieval solution, citrate pH 6.1 (10×) (Agilent, #S169984-2) for 25 min at 100°C.
• 58.Allow slides to cool in solution at 20°C–22°C for 25 min.
• 59.Rinse slides with water 2–3 times and place in TBST.
• 60.Block endogenous peroxides with 0.3% hydrogen peroxide (Abcam, #ab64218) for 5 min at 20°C–22°C rinse slides with TBST.
• 61.Incubate slides in serum free protein block (Agilent, #X0909) for 5 min at 20°C–22°C.
• 62.
  Proceed with immunohistochemical staining for cell type verification.

  • a.
    Incubate slide with 300 μL of primary antibody for 60 min at 20°C–22°C, after incubation rinse with of TBST.

    • i.
      For HSC & HDF (Figure 4B), use α-SMA primary antibody (Abcam, #ab5694) at a dilution of 1:500.
    • ii.
      For HUVEC (Figure 4C), use anti-CD31 primary antibody (Thermo Fisher Scientific, #MA5-16337) at a dilution of 1:50.
  • b.
    Incubate slide with 300 μL of predilute Envision Plus anti-rabbit labeled polymer (Agilent, #K400311-2) for 30 min at 20°C–22°C, rinse slides with TBST.
  • c.
    Incubate slides with 300 μL of predilute DAB+ (Agilent, #K346811-2) for 5 min at 20°C–22°C, rinse with distilled water.
  • d.
    Counter stain slide with 300 μL of predilute Gills I hematoxylin (Sigma-Aldrich, #GHS132) for 30 s, rinse with water.
  • e.
    Dehydrate, clear, and mount slides.

Figure 4.

Immunohistochemical analysis of multiple sections of HH-SFE spheroids

(A) Hematoxylin and eosin (H&E).

(B) α-SMA expression identifies the presence of hepatic stellate cells and fibroblasts around the periphery of the spheroid.

(C) CD31 expression identifies the location of HUVEC endothelial cells within the center of the spheroid. Scale bar = 150 μm.

Image analysis

Timing: 1–2 h

Here we quantify the number of spheroids, determine the spheroid size and estimate the spheroid volume.

• 63.Open the ImageJ Fiji application.⁷ Access the folder containing spheroid images, drag and drop the image into ImageJ Fiji.
• 64.Select the line tool from the toolbar and measure the image scale bar by left clicking and holding to trace the scale bar’s length.

• 65.
  Select the analyze tab from the menu options.

  • a.
    Set the scale from the drop-down menu.

    Note: A dialog box will appear with the “distance in pixels”, the “known distance”, the “pixel aspect ratio”, and the “unit of the length”.

  • b.
    Edit the “known distance” and “unit length” options to the distance and unit in the selected image select the “global” box and click “ok”.

    Note: Consider what units are desired for final volume calculation. If volume will be reported in μm³ or mm³ ensure that “known distance” and “unit of length” are set accordingly. Ex. For reporting in μm³ if image scale is 750 μm “known distance” to 750 and “unit of length” to μm. Reporting in mm3 with 750 μm scale set “known distance” to 0.750 and “unit of length” to mm.

• 66.
  Select the line tool from the menu options.

  • a.
    Draw a line across the longest diameter of the spheroid then click “M” on the keyboard this will save the measurement.
  • b.
    Draw a line across the smallest diameter of the same spheroid and click “M”.

Note: Repeat steps 66a-b for all spheroids in the image.

Note: The longer diameter should be designated as the “length” of the spheroid, and the shorter diameter as the “width” of the spheroid.

• 67.Copy data from ImageJ Fiji into an excel spreadsheet.

Note: Format data so that the length and width of each spheroid in the data set are identifiable.

• 68.To estimate spheroid volumes, apply the formula volume = π$d^3/6$, where $d$ is equal to the average of the length and width of the spheroids.

Expected outcomes

This protocol describes the generation of 2–4 cell type spheroids using both benign and malignant cells. Throughout the development of the method, we observed that all spheroids increased in size and circularity within the first 72 h, with progressive definition of a distinctive outer periphery over time. This protocol results in the formation of large, circular, spheroids (200–400 m in diameter) that are separate from each other and not aggregated. The number of spheroids that form in each sample tube varies with the cells used (Table 1), but the ease of set-up provides a simple route to scalability and mass production.

Table 1.

Volume and total count of HH-HSC, CS, CSF, and CSFE spheroids

	HH-HSC/CS				HH-SF/CSF				HH-SFE/CSFE
	Spheroid count	Avg. length (μm)	Avg. width (μm)	Volume (mm3)	Spheroid count	Avg. length (μm)	Avg. width (μm)	Volume (mm3)	Spheroid count	Avg. length (μm)	Avg. width (μm)	Volume (mm3)
HH	9	538	488	0.092	15	403	353	0.048	6	738	637	0.228
HepG2	45	306	256	0.020	100	300	231	0.011	48	205	186	0.005
SNU1079	67	233	197	0.006	44	356	304	0.022	16	423	356	0.038

The resultant spheroids are robust and highly viable, as demonstrated by calcein-AM/propidium iodide staining (Figure 3), and are suitable for therapeutic testing, invasion, and migration studies, or for immunohistochemical analysis. We observed a lack of a central necrotic core in all of our models which would be desirable for cytotoxicity studies such as for drug screening.

We have observed the following milestones. After 18 h, initial independent spheroids with uneven edges are observed. After 48 h, spheroids have formed and have uniform, spherical shapes. After 72 h, it is possible to observe spheroids contracting while maintaining a round, uniform shape (Figures 2D–2F). At this stage, spheroids can be manipulated without risk of dissociation because cells have formed strong cell-cell connections.

Limitations

We have demonstrated the successful spheroid development of MCS using 5 different types of human cells (HH, HepG2, Hep3B, SNU-1079, and HuCCT1), in conjunction with up to three supporting cell types (HSC, HDF, HUVEC). The efficacy and yield of spheroid formation using this protocol may vary with the type of cells used. Thus, it is recommended that the investigator conducts preliminary experiments to determine the optimal seeding density, culture conditions and cell ratios for desired MCS formation.

The method used for determination of spheroid volume assumes that the spheroid being measured is spherical. Due to the 2D imaging for determination of dimensions, this assumption may not always be accurate; therefore, if accurate quantitation of spheroid volume is desired, we recommend measuring all or a large representative sample and/or replicating experiments.

Troubleshooting

Problem 1

Cell clusters have merged after the first 18 h incubation of the starter culture on the clinostat (Figure 5) (step 18).

Figure 5.

Example of accidental spheroid merging after 18 h of rotation

Scale bar = 100 μm.

Potential solution

The risk of unwanted spheroid merging/clumping can be reduced by the following actions: Visually ensuring that spheroid units are well resuspended after starter culture incubation. If initial vortex of 5 s does not resuspend cellular cluster increasing vortex time to 10–15 s may help to prevent clumping.

Problem 2

Spheroids merge after 48 or 72 h on clinostat (step 29).

Potential solution

The risk of merging/clumping at this later stage of spheroid development may be mitigated by, increasing expansion culture tube size, or reducing starter culture cell numbers to reduce their density. This may reduce the spheroid to spheroid contact and help to keep the spheroids from clumping and merging. If certain cell types have an increased tendency to form a single spheroid during later stages of the described protocol, we recommend retrieval for desired experimentation or analysis at an earlier time point of 48 h instead of 72 h.

Problem 3

Overlapping spheroids during final image acquisition (steps 34 & 35).

Potential solution

Check spheroids under microscope to ensure that none are overlapping before starting image acquisition. If spheroids are overlapping increase PBS volume by 100 μL and gently tap the side of the chamber slide to distribute the spheroids.

Problem 4

Spheroids have low viability (step 40).

Potential solution

There are several possible reasons for low viability of cells within MCS. An assessment of cell viability at the start of the protocol prior to use will avoid the use of unhealthy cells. A possible cause of reduced viability within spheroids is excessive duration of staining with calcein-AM/propidium iodide, as long-term exposure to propidium iodide can be toxic. To mitigate this risk, MCS should be imaged without delay after staining with calcein-AM/propidium iodide at defined time points. Other potential causes of decreased viability include the duration of the image acquisition and transfer-induced damage. During imaging, spheroids may be exposed to suboptimal temperatures and CO₂ conditions; reducing the duration of this exposure may improve their overall viability. This can be accomplished by imaging groups of 3–5 tubes containing spheroid samples simultaneously. Transferring spheroids may cause them to sustain mechanical damage. To avoid this, pipette tips should have a wide opening that reduces lodging of spheroids within the pipette tip.

Problem 5

Spheroids becoming trapped in tube while transferring (steps 22, 34, and 37).

Potential solution

Spheroids may become trapped in tube during transfer. Cutting pipette tips while necessary, can make spheroid retrieval more difficult. When transferring, allow the spheroids to settle to the bottom of the tube. Then, using a cut pipette tip gently resuspend the spheroids, the now controlled floating spheroids can be captured in one motion. This should allow for the retrieval of most of the spheroids. If additional spheroids remain trapped in tube add additional media and repeat process.

Resource availability

Lead contact

Further information and requests should be directed to Tushar Patel; patel.tushar@mayo.edu.

Materials availability

Not applicable as this protocol is not associated with any newly generated materials.

Data and code availability

Not applicable.