Generation and Expansion of Primary, Malignant Pleural Mesothelioma Tumor Lines

Summary

The goal of this method paper is to demonstrate a robust and reproducible methodology for the enrichment, generation, and expansion of primary tumor cell lines from surgically resected pleural mesothelioma.

Abstract

Current methodologies for the expansion of primary tumor cell lines from rare tumor types are lacking. This protocol describes methods to expand primary tumor cells from surgically resected, malignant pleural mesothelioma (MPM) by providing a complete overview of the process from digestion to enrichment, expansion, cryopreservation, and phenotypic characterization. In addition, this protocol introduces concepts for tumor generation that may be useful for multiple tumor types such as differential trypsinization and the impact of dissociation methods on the detection of cell surface markers for phenotypic characterization. The major limitation of this study is the selection of tumor cells that will expand in a two-dimensional (2D) culture system. Variations to this protocol, including three-dimensional (3D) culture systems, media supplements, plate coating to improve adhesion, and alternate disaggregation methods, could improve this technique and the overall success rate of establishing a tumor line. Overall, this protocol provides a base method for establishing and characterizing tumor cells from this rare tumor.

Introduction

Malignant pleural mesothelioma (MPM) is a rare tumor highly associated with asbestos exposure. Although immunotherapy-based approaches have shown encouraging results, there is a paucity of treatment options available to patients that develop this disease, and the overall 5-year survival rate is low^1,2. Efforts are underway at multiple institutions to better understand this disease and identify novel therapeutic targets that may improve patient outcomes. While there are multiple mesothelioma mouse models, access to primary mesothelioma tumor cells is more limited³. In vitro expansion of primary mesothelioma tumor cells would provide a valuable model system that can be utilized to study tumor cells directly and their interaction with autologous immune cells such as tumor-infiltrating lymphocytes. While there have been reports on the expansion of primary mesothelioma tumor cells lines, these are few and do not provide a detailed standard operation procedure (SOP). Furthermore, few cell lines are available from commercial sources such as American Type Culture Collection (ATCC). While the availability of primary tumor lines is limited, it has been demonstrated that tumor cells can be expanded from pleural effusions and directly from the tumor tissue^4,5. In addition, expanded tumor cell lines have been shown to preserve the molecular profile of the original tumor^4,5,6.

Our laboratory is studying the tumor-immune microenvironment of MPM and has developed a method for expanding primary MPM tumor cells lines from surgically resected cases. This method is adapted from our experience in establishing primary metastatic melanoma tumor cell lines. The goal of this work is to provide a detailed, practical approach to primary mesothelioma tumor line expansion using a 2D model system and subsequent phenotypic profiling. Given the recent success of checkpoint blockade strategies targeting CTLA4 and PD-1 in the first-line setting², the ability to generate many primary tumor lines could further the understanding of both tumor intrinsic mechanisms of resistance as well as provide an important model system to assess T-cell recognition, thus deepening our understanding of the immune response in MPM.

The major limitation of this protocol is that every tumor contains a different microenvironment, and there is a high degree of variability of expansion success. In addition, this method selects for tumor cells that can expand in a 2D culture system. Other methods that involve the production of a 3D spheroid or organoid culture could provide an alternative approach that may allow for a higher success rate of expansion or result in the ability to derive cell lines that are unable to expand in a traditional 2D system. Such 3D cultures have been demonstrated to be useful for the generation of tumor types that are difficult to expand, for example, pancreatic cancer⁷.

Protocol

All methods described here have been approved by the Institution Review Board (IRB) of the University of Texas MD Anderson Cancer Center. This pertains to standard-of-care, surgically resected MPM tumors removed following informed consent.

1. Preparation of tumor digestion media and other associated media

1. To prepare tumor digestion media, add 5 mL of 100% Pen/Strep (Penicillin/Streptomycin) to 500 mL of sterile Roswell Park Memorial Institute (RPMI) 1640 medium in a laminar flow hood.
   1. Transfer the medium to a 500 mL, 0.22 µm polyethersulfone (PES) sterile filter for vacuum filtration.
      NOTE: Filtration and aspiration steps should be performed with a standard vacuum pressure of 75-150 torr.
   2. Label and date the tumor digestion medium and store it at 4 °C.
      NOTE: The medium can be stored for 1 month at 4 °C.
2. To prepare the tumor-digesting enzymatic cocktail, add 2.5 mL of 3% collagenase type 1, 1 mL of 1.5 mg/mL hyaluronidase, 20 µL of 250,000 units/mL DNAse 1 to 16.5 mL of digestion medium in a 50 mL conical tube in a laminar flow hood.
   NOTE: The total volume of the tumor-digesting enzymatic cocktail is 20 mL; however, only 10 mL is needed per tumor sample. As such, the remaining cocktail can be stored at -20 °C until needed. Do not refreeze aliquots.
3. To prepare complete tumor medium, add 5 mL of 100% Pen/Strep and 50 mL of fetal bovine serum (FBS) to 500 mL of sterile RPMI 1640 in a laminar flow hood.
   1. Transfer the medium to a 500 mL, 0.22 µm PES sterile filter for vacuum filtration.
   2. Label and date the complete tumor medium and store it at 4 °C.
      NOTE: The medium can be stored for 1 month at 4 °C.
4. To prepare reduced-serum medium for starvation, add 5 mL of 100% Pen/Strep and 5 mL of FBS to 500 mL of sterile RPMI 1640 in a laminar flow hood.
   1. Transfer the medium to a 500 mL, 0.22 µm PES sterile filter for vacuum filtration.
   2. Label and date the reduced-serum medium and store it at 4 °C.
      NOTE: The medium can be stored for 1 month at 4 °C.
5. To prepare freeze medium, add 5 mL of dimethyl sulfoxide (DMSO) to 45 mL of FBS in a laminar flow hood.
   1. Transfer the medium to a 50 mL, 0.22 µm PES sterile filter for vacuum filtration.
   2. Label and date five 15 mL conical tubes.
   3. Transfer 10 mL of freeze medium to each tube and store the tubes at -20 °C.
6. To prepare antibiotic-free medium for mycoplasma testing, add 50 mL of FBS to 500 mL of sterile RPMI 1640 in a laminar flow hood.
   1. Transfer the medium to a 50 mL, 0.22 µm PES sterile filter for vacuum filtration.
   2. Label and date the antibiotic-free medium and store it at 4 °C.
      NOTE: The medium can be stored for 1 month at 4 °C.
7. To prepare fluorescence-activated cell sorting (FACS) buffer for phenotypic analysis, add 16.6 mL of sterile bovine serum albumin to 500 mL of sterile 1x phosphate-buffered saline (PBS) in a laminar flow hood.
   1. Label and date the FACS buffer and store it at 4 °C.
      ​NOTE: FACS buffer does not require filter sterilization as long as both components are stored sterile. FACS buffer can be stored prior to opening for 6 months at 4 °C.

2. Digestion of tumor tissue

1. Transfer 10 mL of tumor-digesting enzymatic cocktail from step 1.2 to a dissociation tube.
2. Transfer the piece of tumor tissue to a sterile 6-well plate lid using a sterile scalpel.
   NOTE: The amount of medium that is transferred with the tumor tissue is sufficient for processing.
   1. Remove all fatty tissue, necrotic tissue, and/or blood clots using a new sterile scalpel and forceps.
      NOTE: Fatty tissue is normally yellowish in appearance and semisolid. Necrotic tissue is darker than the surrounding tissue in appearance and very soft; both fatty and necrotic tissue will break apart easily. Bloody tissue appears bright red and may release blood into the medium when manipulated. Once removed, the resulting tumor tissue should hold shape and be pale or pink, depending on the tissue type.
   2. Cut the entire tumor tissue into 1 mm³ small fragments. To ensure the tissue does not dry out, add 500 µL of sterile 1x Hank's Balanced Salt Solution (HBSS) to the tumor tissue.
3. Transfer tumor fragments into the dissociation tube containing 10 mL of tumor-digesting enzymatic cocktail (step 2.1). Close the tube tightly, place it cap-side down on the tissue dissociator, and add the heater apparatus by applying it like a sleeve over the dissociation tube and pressing to click it in place.
   NOTE: The maximum capacity for each tube is 4 g of tumor and 10 mL volume.
4. Select the preprogrammed setting on the dissociator 37C_h_TDK_1. Check the status after 10 min to ensure there is no clogging error as indicated by the flashing red light.
   NOTE: This program processes at 1,865 rpm for 1 h with temperature on at 37 °C.
   ​NOTE: If clogging occurs, remove the dissociation tube and manually swirl it to dislodge any tissue caught in the lid; then replace the tube onto the dissociator and resume the program.
5. Following the 1 h digestion, remove the dissociation tube and place it in a laminar flow hood. Filter the digested tumor by placing a 70 µm cell strainer on top of a 50 mL conical tube and pipetting the digested tumor using a 10 mL pipette onto the filter. If the filter clogs, switch to a new filter.
   1. Rinse the dissociation tube with 10 mL of fresh tumor digestion media and pass the wash through the filter.
   2. Remove the 70 µm filter and discard it.
   3. Bring the total volume to 40 mL with tumor digestion medium.
6. Centrifuge at 500 × g for 5 min at room temperature.
7. Using a 2 mL aspirating pipette attached to a vacuum source, aspirate the supernatant without disturbing the pellet, and resuspend the cells using 10 mL of sterile complete tumor medium.
8. Repeat step 2.6.
9. Aspirate the supernatant as described in step 2.7. Resuspend the cells in 3 mL of sterile complete tumor medium and transfer the cell suspension to one well of a sterile 6-well plate.
10. Keep the plate in an incubator at 37 °C with 5% CO₂.
11. After 24 h, transfer the used medium from the digested tumor in well 1 to a new well (well 2) in the same 6-well plate. Add 3 mL of sterile complete tumor medium to well 1.
12. Transfer the used medium from wells 1 and 2 and combine into well 3 in the same 6-well plate 24 h after step 2.11. Add 3 mL of sterile complete tumor medium into wells 1 and 2.
13. Aspirate the medium from all 3 wells and pipette 3 mL of complete tumor medium into each well 48 h after step 2.12.

3. Generation of primary tumor cell line

1. Using an inverted phase microscope, determine the percentage of fibroblast contamination in the early passage culture.
   NOTE: Fibroblasts are generally long and irregular and have an ill-defined cellular membrane. They appear thin or flat on a Z scale.
   1. If fibroblast contamination is greater than 20% of the cells in the early passage culture, continue culturing after replacing the complete medium with reduced-serum medium.
   2. Repeat the replacement with reduced-serum medium in step 3.1.1 twice per week for 3-4 weeks.
   3. When the culture reaches 80% confluency, passage the cells by splitting 50:50 into 2 new wells of a 6-well plate. Continue passaging 50:50 in reduced-serum media once the cells reach 80% confluency.
   4. Repeat step 3.1.3 for up to 4 weeks, passaging in larger culture flasks as needed by splitting 50:50 once the culture reaches 80% confluency. If the fibroblast population is not reduced to 10% or less, continue to step 3.2 to attempt the differential trypsinization method to remove fibroblast contamination. Otherwise, proceed to step 3.3.
2. To enrich for tumor cells, gently rinse the well with 1x PBS to remove the medium containing serum.
   1. Add trypsin as follows: 1 mL per well if in a 6-well plate, 2 mL for a T25 flask, 3 mL for a T75 flask.
   2. Place the 6-well plate or flask in an incubator at 37 °C for 1 min. Remove the plate or flask from the incubator and check the adhesion of the cells using an inverted microscope. If the cells are partially lifting or floating, gently remove the suspended cells and trypsin only. Place the cells in a 15 mL conical tube containing an equal or greater volume of complete tumor medium.
      NOTE: This partial collection of cells based on adhesion properties is the first of multiple fractions.
   3. Repeat steps 3.2.1 and 3.2.2 until all the cells are lifted, or there are 4 fractions removed.
   4. Centrifuge all independent fractions at 500 × g for 5 min at room temperature.
   5. Aspirate the supernatant as described in step 2.7, and resuspend the cells using 5 mL of sterile, reduced-serum medium.
   6. Plate the cells in a T25 flask for each fraction and place the flasks in an incubator at 37 °C.
   7. Assess the culture after 48 h to determine which fraction is enriched for tumor cells versus fibroblasts. Discard fibroblast-rich flasks. If fibroblast contamination is <10%, continue expansion in complete tumor medium and proceed to step 4. If fibroblast contamination is 10-30%, repeat steps 3.1.2-3.1.4. If fibroblast contamination is greater than 30%, repeat steps 3.2-3.2.7.
3. If few or no fibroblasts are detected within the early passage culture, continue passaging the cells in complete tumor medium by splitting 50:50 once the cells are 80% confluent in their 6-well plate or flask.

4. Expansion of early passage primary tumor cell line

1. Once the primary tumor culture contains 10% or less of fibroblast contamination, aspirate the medium and replace it with sterile complete tumor medium twice per week.
   NOTE: The optimal medium volume for a T25 is 5 mL; T75 is 12 mL; T150 is 25 mL.
   1. Passage the culture 50:50 into larger flasks as needed once it reaches 80% confluency in the plate or flask.
      NOTE: The culture may need to be passaged at a higher ratio depending on its growth. Adjust so the culture reaches 80% confluency every 3-5 days.
2. Passage until the culture is at passage 4 or above. Once the culture is 80% confluent in a minimum of two T75 flasks, continue to step 5.

5. Characterization and banking of established primary tumor cell line

1. Cryopreserve one T75 flask as an early-passage freeze. For mycoplasma testing of the remaining T75 flasks, continue to step 5.2.
   1. For cryopreservation of early-passage cells, thaw 1 tube of aliquoted freeze medium prepared in step 1.5.
   2. Collect the cells by trypsinization by first washing the plate with 1x PBS as described in step 3.2 and adding trypsin as described in step 3.2.1.
   3. Place the flask in an incubator at 37 °C for 3 min. Remove the flask from the incubator and check the adhesion of the cells using an inverted microscope. If the cells are lifting or floating, gently remove the suspended cells and trypsin. Place the cells in a 15 mL conical tube containing an equal or greater volume of complete tumor medium.
      NOTE: If the cells remain adherent after 3 min, place the flask back in the incubator for an additional 2 min.
   4. Mix the tube well by pipetting gently and remove 20 µL for counting.
   5. Centrifuge the tube at 500 × g for 5 min at room temperature.
   6. While the cells are spinning, count using a lab-specific counting protocol such as trypan blue or acridine orange/propidium iodide (AO/PI).
   7. Resuspend the cells following centrifugation in freeze medium with a minimum of 2 × 10⁶ cells/1 mL freeze medium.
   8. Transfer 1 mL of the cell suspension from step 5.1.7 to prelabeled 1.5 mL cryovials.
   9. Place the cryovials in a controlled-rate freeze chamber and transfer them immediately to -80 °C.
   10. Store the cells at -80 °C for up to 1 week before transferring them to liquid nitrogen for long-term storage.
2. Split a T75 flask 50:50 for mycoplasma testing. Change the medium to antibiotic-free medium prepared in step 1.6.
   1. After 72 h, thaw the mycoplasma detection reagent, substrate, and controls at room temperature for 15 min before testing.
   2. Collect 1 mL of the supernatant from each culture to be tested and place it in a 15 mL conical tube.
   3. Pellet any suspended cells by centrifuging the supernatant at 500 × g for 5 min at room temperature.
   4. Load 100 µL of sample supernatants and controls into a white 96-well plate. Add 100 µL of mycoplasma detection reagent to each sample and control.
   5. Incubate for 5 min at room temperature.
   6. Place the plate in a plate reader and read the luminescence (Reading A, i.e., first Reading).
      NOTE: The mycoplasma kit manufacturer's protocol indicates keeping the default settings on multifunctional plate readers. The read is set at Luminescence endpoint with an integration time of 1 s and gain of 135. The plate reader uses an Auto-scale routine to optimize the gain signal for each experiment. The 1 s integration time is standard default. Both settings are used to adjust the intensity of the luminescent signal interpreted by the plate reader and may need to be adjusted depending on the individual plate reader.
   7. Remove the plate from the plate reader and add 100 µL of the mycoplasma detection substrate to each sample and the controls.
   8. Incubate for 10 min at room temperature.
   9. Place the plate in the plate reader and read the luminescence (Reading B, i.e., second Reading).
   10. To determine mycoplasma positivity, determine the ratio of Reading B to Reading A.
       NOTE: Mycoplasma positivity ratios are described in the mycoplasma kit manufacturer's protocol. Readings A and B are acquired with the same settings and simply reflect the reading order.
       1. Consider a ratio < 1 to be mycoplasma-negative.
       2. Consider a ratio of 1-1.2 to be inconclusive and repeat step 5.2.
       3. Consider a ratio > 1.2 to be mycoplasma-positive and monitor this culture; terminate the culture if necessary.
3. Flow cytometry characterization of mycoplasma-negative tumor cells
   1. Dislodge the tumor cells using cell-dissociation buffer.
   2. Count the cells and resuspend the cells at 1 × 10⁶/mL in FACS buffer from step 1.7.
   3. Remove 100 µL of cell suspension into individual tubes for surface staining.
   4. Centrifuge at 500 × g for 5 min at room temperature.
   5. Aspirate the supernatant and block Fc receptors by incubating the cells in 500 µL of 5% goat serum in FACS buffer at room temperature for 10 min.
   6. Add 2 mL of FACS buffer and centrifuge the suspension at 500 × g for 5 min at room temperature.
   7. Aspirate the supernatant and add the surface stain mix of antibodies (Table 1) and FACS buffer so that the final volume is 100 µL. Cover the samples and stain them on ice for 30 min.
   8. Repeat step 5.3.6.
   9. Aspirate the supernatant and fix the cells by resuspending in 200 µL of 1% paraformaldehyde (PFA) plus 0.25% EtOH. Incubate at room temperature for 20 min with the samples protected from light.
   10. Repeat step 5.3.6. Resuspend the cells in 200 µL of FACS buffer for acquisition using an appropriate flow cytometer.
       NOTE: Mesothelioma tumor cells are expected to be positive for mesothelin and N-cadherin. Some may also express CD90.
4. Expand in complete tumor medium and bank as described in steps 4.1 and 5.1, respectively.

Results

To determine fibroblast contamination of early-passage cultures, cells are assessed using an inverted phase microscope to identify the frequency of fibroblasts relative to the other adherent cells present. Figure 1 shows examples of increasing fibroblast contamination of 80% (Figure 1A), 50% (Figure 1B), and 30% (Figure 1C), compared to a culture with no fibroblast contamination (Fi...

Discussion

While this protocol is straightforward, there are a few critical steps that must be closely followed. The early-passage freeze is important to preserve the ability to repeat the tumor-cell enrichment process if initially unsuccessful. The ability to assess fibroblastic contamination by eye to decide the correct splitting and media starvation technique is vital to preventing fibroblast overgrowth in the culture. In addition, the differential trypsinization method requires careful observation of the cells during incubation...

Materials

Name	Company	Catalog Number	Comments
10 mL serological pipettes	BD Falcon	357551
15 mL conical tubes	BD Falcon	352097
2 mL aspirating pipettes	BD Falcon	357558
5 mL serological pipettes	BD Falcon	357543
50 mL conical tubes	BD Falcon	352098
6-well microplates, tissue culture treated	Corning	3516
Accutase	Innovative Cell Technologies	AT104	A protease-collagenase mixture considered to be more effective in preserving epitopes for flow cytometry.
anti-CD325 (N-Cadherin) PE	Invitrogen	12-3259-42
anti-CD90 PE-Cy7	BD Biosciences	561558
anti-Mesothelin APC	Miltenyi	130-118-096
Bovine Serum Albumin 30%	Sigma	A8577-1L
Cell Dissociation buffer, enzyme-free	Thermo Fisher Scientific	13151014
Cell strainer (70 µm)	Greiner Bio-One	542-070
Centrifuge with 15 mL and 50 mL adaptors
Collagenase Type 1	Sigma-Aldrich	C-0130	Dissolve 1.5 g of Collagenase type I into 100 mL of sterile DMEM and aliquot in 5 mL volumes. Label well and store at -20 °C.
Controlled-rate freezing chamber	Thermo Fisher Scientific	15-350-50
Cryovials	Thermo Fisher Scientific	5000-0020
CulturPlate-96	Packard Instrument Company	6005680	White, opaque 96-well microplate.
Dimethyl sulfoxide	Thermo Fisher Scientific	BP231-100
DNAse I (from bovine pancreas)	Sigma-Aldrich	D4527	Aliquot at 50 μL, label well and store at -20 °C. After thawing, keep at 4 °C for up to 1 month.
Dulbecco's Phosphate buffered saline solution 1x	Corning	21-031-CV
Ethanol 200 proof	Thermo Fisher Scientific	A4094
FACS tubes-filter top	BD Falcon	352235
Fetal bovine serum	Gemini-Bio	100-106
GentleMACS C-tubes	Miltenyi	130-093-237
GentleMACS Octo-dissociator with heater apparatus	Miltenyi	130-096-427	Includes specialized heater apparatus sleeves used to apply heat to the C-tubes during dissocation.
Goat serum	Sigma-Aldrich	G9023	Aliquot and store at -20 °C.
Hank's Balanced Salt solution, 500 mL	Corning	MT21022CV
Hyaluronidase	Sigma-Aldrich	H3506	Dissolve 0.15 g of Hyaluronidase into 100 mlLof sterile DMEM and distribute into 1.0 mL aliquots. Label well and store at -20 °C.
Inverted-phase microscope
Laminar flow hood
Live/dead yellow dye	Life Technologies	L-34968
Micropipettor tips, 20 µL	ART	2149P
Micropipettor, 0.5-20 µL
Mycoalert assay control set	Lonza	LT07-518	Aliquot positive controls and store at -20 °C.
Mycoalert Plus mycoplasma detection kit	Lonza	LT07-710	Aliquot reagent and substrate and store at -20 °C. Save remaining buffer and aliquot for negative control and store at -20°C.
Paraformaldehyde 16%	Electron Microscopy Sciences	15710	Prepare stock by filtering through a PVDF syringe filter (Millex cat. no. SLVV033RS) and aliquot under a fume hood. Store at -20 °C.
Penicillin-streptomycin 10,000 U/mL	Gibco	15140-122
Pipet aid
Plate reader with luminescent capabilities	BioTek	Synergy HT	any plate reader with these specifications can be used
RPMI 1640 media	Corning	10-040-CV
Scalpel	Andwin	2975#21
Stericup Quick Release-GP sterile vacuum filtration system, 0.22 µm PES Express PLUS, 500 mL	EMD Millipore	S2GPU05RE
Steriflip-GP filter, 0.22 µm PES Express PLUS, 50 mL	EMD Millipore	SCGP00525
Sterile forceps	Thermo Fisher Scientific	12-000-157
T25 flasks, tissue culture treated	Corning	430639
T75 flasks, tissue culture treated	Corning	430641U
Trypan blue solution 0.4%	Gibco	15250-061
Trypsin EDTA 0.05%	Corning	25-052-CI
ViaStain acridine orange/propidium iodide (AO/PI) solution	Nexcelom	CS2-0106-5ML