Airway segments were collected at the time of necropsy from rhesus macaques with healthy lungs. Tissue was from adult rhesus macaque monkeys, age 2.5 years, 3.5 years and 8.5 years old. Cells obtained from animals older than 9 years did not yield sustainable cultures (Supplemental Table S2
). Macaque basal cells are cultured on collagen coated plates (Section 3.1
). The coating functions as a method of selection to purify the basal cells from the cell isolate after tissue dissociation (Section 3.2
). After attachment and selection, the cells can be maintained and amplified in proprietary media without coating (Section 3.3
). Under maintenance conditions, the cells do not differentiate, and we thus have optimized differentiation conditions for the macaque derived basal cells, commonly used in the field (Section 3.4
), namely air–liquid interface differentiation (Section 3.4.1
) and differentiation in tracheospheres (Section 3.4.2
). mmuBCs can further be analyzed and manipulated by common techniques such as viral transfection (Section 3.4.3
) and flow cytometric analysis and sorting (Section 3.4.4
). For long term storage, cells can be cryopreserved (Section 3.5
). For more details on expected results, yields and outcomes per section, we would like to refer to Section 4
, Table 1
, and for troubleshooting Section 4
, Table 2
3.1. Coating: Time for Completion 02:30 h
Prepare 5 mL of coating medium (see Section 5
) per 10 cm plate and pipet onto plate and ensure even distribution.
Incubate plate for 1–2 h at 37 °C.
Remove coating medium from plate and let dry for 30 min in a laminar flow hood.
Once dried, seal plates with Parafilm.
PAUSE Coated plates can be stored up to 2 weeks at 4 °C.
3.2. Tissue Dissociation and Cell Isolation: Time for Completion 20 h
Cut trachea into 3–4 smaller pieces using surgical scissors or a scalpel.
Dissect the upper, large airways out of the macaque lungs, ensure that as little lung tissue remains on these airways as possible.
Put pieces into clean wash buffer.
Remove any remaining lung, fat, connective and lymphoid tissue.
Cut the airway in 0.5 cm pieces and open the airway with a single longitudinal cut.
Transfer the pieces into a 50 mL conical tube containing 20 mL of cold digestion buffer.
Seal tube with paraffin, put tube into sealable plastic bag and incubate overnight for up to 16 h at 4 °C with gentle agitation.
Inactivate the digestion buffer with 2mL of FBS (to a final concentration of ~10% FBS). The tissue will appear swollen but will still be intact.
Remove pieces from conical tube and put into a 10 cm dish containing wash buffer and scrape off epithelial layer with a scalpel. Keep the digest medium on the side.
CRITICAL STEP Scraping should be done carefully, not cutting into the underlying stroma. We recommend holding a corner of the tissue with forceps and then stroking with the scalpel in a single direction, with a movement similar to spreading butter on toast. Discard remains.
Combine wash buffer with scrapings with digest medium together and centrifuge for 5 min, at 500× g.
Resuspend cells in 2–3 mL of Pneumacult-Ex (containing Penicillin, Streptomycin and Amphotericin B) and count cells or collect pellet for further analysis.
Typically, 1 × 106 cells can be harvested per 2–3 g of starting material.
Cells are counted using an automated counter (Luna II).
3.3. Culture and Maintenance: Time for Completion 4 to 5 days
For resuspension of the pellet, pipette up and down gently 3–4 times.
CRITICAL STEP Do not filter the resuspended pellet, BCs will attach, non-basal cells will not and can be washed off the next day. Cell yield is markedly increased if not filtered.
Plate between 2–6 × 106 cells/10 cm dish (3–11 × 104 cells/cm2).
After plating, leave cells undisturbed for a minimum of 24 h.
The next day, remove medium, wash cells gently with PBS (containing CaCl and MgCl) and add fresh, warm medium.
Change medium every 48–72 h. Medium needs to be removed completely and cells are washed 1× with warm PBS (containing CaCl and MgCl) and 8–10 mL of fresh, warm Pneumacult-Ex are added.
Reduce antifungal drugs gradually after 10–14 days in culture, by reducing the concentration 10-fold with every medium change.
Due to clonal expansion, macaque BC cultures will never reach 100% confluency, subculture at 80% confluence. If cultures are grown for long time at maximum confluency, they will become quiescent.
After the first passage cells do not need to be passaged onto coated dishes (see Section 3.1
). For the first passage, after flow sorting, cell manipulations and after thawing collagen plates are an essential requirement.
Remove old medium but keep for later use.
Wash 2× with warm PBS.
Add 1mL of 0.25% trypsin per 10 cm plate. Scale down trypsin accordingly to plate size.
Incubate for 3–7 min at 37 °C. Earlier passages will detach faster (3 min) than later passage numbers (up to 10 min maximum).
Gently tap dish and check under microscope that cells have detached, maximally incubate for 10 min in trypsin. Viability and yield tend to drop after 10 min in trypsin.
Stop reaction adding 2 mL of FBS and wash plate with old medium.
Centrifuge cells for 5 min at 500× g.
Remove supernatant and resuspend in 1 mL of fresh medium.
Plate 0.5 × 106 cells per 10 cm plate in 10 mL fresh medium per plate.
as few as 0.1 × 106
cells can be plated in a 10 cm dish, initial growth will be slower. If even lower numbers of cells are plated, conditioned medium is highly recommended (see Section 5
3.4. Manipulation and Differentiation: Time for Completion-Variable
For staining purposes, plate ~1 × 104 cells per well of a chamber slide and fix the day after in 4% paraformaldehyde (PFA) for 20 min.
Wash gently 3× with PBS.
Keep in PBS at 4 °C until needed.
For staining of cells on chamber slide, it is recommended to stain directly within the chamber slide and only remove the grid after completion of staining. A more detailed staining protocol can be found in Supplementary Methods
3.4.1. Differentiation on Air–Liquid Interphase (ALI): Time for Completion 14 days
Coat ALI insert for 30 min with coating medium in the incubator, remove the excess and afterwards dry for 30 min in a laminar flow hood.
Plate at 5 × 104 cells/24-well insert density.
Add 400 µL of Pneumacult-Ex to the well and 200 µL to the cell culture insert.
Culture cells according to maintenance protocol until confluency is reached and a cobblestone pattern is visualized.
Once cells have reached confluency on insert, change medium from Pneumacult-Ex to Pneumacult-ALI medium in a gradual fashion.
CRITICAL STEP Gradual change from Pneumacult-Ex to Pneumacult-ALI is highly recommended as cell layers tended to rupture or detach if an immediate change is performed. Gradual change results in more even, healthier cultures. For optimal culture health, media should be changed every 48 h at this point, until in 100% Pneumacult-ALI.
Day 1 on ALI: Remove the medium from the insert and add fresh medium to the bottom of the well (400–500 µL). The insert is submerged in the medium allowing nutrients to diffuse through the lower membrane while the culture is exposed to the air.
On day 1 replace medium with 70% Pneumacult-Ex and 30% Pneumacult-ALI.
On day 3 replace medium with 50% Pneumacult-Ex and 50% Pneumacult-ALI.
On day 5 replace medium with 30% Pneumacult-Ex and 70% Pneumacult-ALI.
On day 7 replace medium with 100% Pneumacult-ALI medium.
Maintain cells thereafter in Pneumacult-ALI medium.
Completely replace Pneumacult-ALI medium every 48–72 hours.
If ALI cultures are to be analyzed, fix and stain the cultures on the insert. For fixation, the medium in the bottom is removed and PBS is added. The cells are put back into PBS for 2–3 min. The PBS in the bottom is removed, this step is repeated 1–2 times. After washing, 4% PFA is added to the bottom of the well and to the insert for 5–10 min.
Cut the insert from the transwell or stain whole insert within the transwell and cut after staining. A more detailed staining protocol can be found in Supplementary Methods
3.4.2. Differentiation in Tracheosphere Assay: Time for Completion 14 days
Thaw 3D Matrigel on ice.
Resuspend cells in cold Pneumacult-Ex at a concentration of 100 cells/µL.
Add cell culture inserts or small cell culture grade glass cover slips to plates (no coating needed) and UV irradiate for 5–10 min.
Mix cells in 3D Matrigel 50:50 (to a final cell concentration of 50 cells/µL).
Add 90 µL of Matrigel/Cell Mix onto the insert or the glass coverslip.
CRITICAL STEP Matrigel polymerizes quickly once at room temperature. We recommend using cold media, and doing all steps on ice. We usually use ice cold pipette tips for this step. Avoid air bubbles in the 3D Matrigel.
For plating on insert, tap the plate after every 90 µL drop that is added to insert, so that there is an even layer. For plating on glass coverslip, let the drop polymerize without spreading.
Let Matrigel polymerize for 15–20 min in the incubator before adding medium.
For the first 5 to 7 days the cells are cultured in Pneumacult-Ex.
Feed cells every other day with Pneumacult-Ex.
Once multicellular spheres are visible in the microscope (after 5–7 days in culture), transition the medium gradually into differentiation medium.
For differentiation, feed cells for 2 days with 50% Pneumacult-Ex and 50% Pneumacult-ALI and afterwards with 30% Pneumacult-Ex and 70% Pneumacult-ALI. Replace medium completely every 48–72 h.
CRITICAL STEP The tracheospheres should not be moved to 100% Pneumacult-ALI. They are ultimately cultured in 30% Pneumacult-Ex and 70% Pneumacult-ALI medium. Sphere morphology is better in in this 30:70 medium compared to 100% Pneumacult-ALI.
If the cells are not moved to Pneumacult-ALI medium, they will not differentiate. If maintained for too long in Pneumacult-Ex they will become solid spheres of Krt5+ basal cells.
For fixation, resuspend the Matrigel in 4% PFA by pipetting up and down.
After 10–15 min, transfer the slurry of PFA, Matrigel and Spheres into a 1.5 mL Eppendorf tube and fill with PBS (containing CaCl and MgCl).
Centrifuge spheres (300× g, 3 min) and wash the pellet twice with PBS (containing CaCl and MgCl).
Stain spheres as whole mount if they are cultured for less than 14 days, afterwards the spheres become too dense and impermeable to the antibodies. After 14 days in culture, the tracheospheres will need to be sectioned to perform staining. A more detailed embedding and staining protocol can be found in Supplementary Methods
For tracheosphere cultures at clonal density (5–10 cells/µL), the cells need to be cultured in a minimum of 30% conditioned Pneumacult-Ex Medium (see Section 5
). They will take longer to form spheres (usually 10 days). The transition into differentiation medium should not be done before 14 days in vitro.
3.4.3. Transduction of Macaque BC Cultures: Time for Completion 72 h
To demonstrate ability to transduce macaque basal cells, we utilized a preexisting lentivirus that had been developed for infecting human cells.
Plate between 0.5–1.0 × 106 cells/10 cm dish (0.75–1.5 × 104 cells/cm2).
After plating, leave cells for a minimum of 24 h before further manipulation.
For transduction, dilute virus of choice in Pneumacult-Ex and keep on ice.
Remove medium from macaque BC cultures, wash once with PBS (containing CaCl and MgCl). Add medium containing high MOI of virus (MOI > 100) to adherent cells.
Twenty-four hours after transduction, but no later than 36 h, wash the medium off the cells and replaced with fresh Pneumacult-Ex.
Using a Lentiviral construct containing GFP under the CMV promoter, GFP expression is observed as early as 24 h post infection and will increase over the next 72 h (see Supplementary Materials
and Figure 4).
Infected macaque BCs should be used within 7–14 days for experiments because transgene silencing reduced GFP expression. Even after sorting expression can be reduced by up to 60% within 14 days. In order to avoid silencing and establishing stable lines, Lenti-SIV virus constructs should be used [7
GFP+ infected cells can be frozen (see Section 3.5
) and thawed at a later time point.
3.4.4. Sample Preparation for Flow Cytometric Analysis or Purification
Flow Cytometric Analysis: Time for Completion 2 h
Dissociate cells (see Section 3.3
) and resuspend in 1mL of 4% PFA in a 15 mL conical tube for 5 min.
Wash out PFA by adding 13 mL of PBS (containing CaCl and MgCl) and centrifuge (300× g, 5 min).
Resuspend cells in 1 mL of PBS (containing CaCl and MgCl) in a 15 mL conical tube.
Gently agitate cells on a vortex while adding 100% ethanol dropwise to a volume of 10 mL (resulting in 90% ethanol).
Keep cells at −20 °C for 30 min.
PAUSE Fixed and ethanol permeabilized cells can be stored in 90% ethanol for up to 5 days at −20 °C.
For further staining, cells are centrifuged (500× g
, 3 min) and washed twice with Flow Cytometry Buffer (see Section 5
) followed by a 30 min blocking step in Flow Cytometry Buffer and stained with antibodies of interest thereafter. A more detailed protocol can be found in Supplementary Methods
Purification by Flow Sorting: Time for Completion 30 min
Dissociate cells (see Section 3.3
) and resuspend 1 × 106
cells/100 µL of Flow Cytometry buffer.
Add Calcein Blue at a concentration of 1 µL/mL of a 10 µM stock.
Filter cells through the cell strainer cap of polystyrene round bottom tubes (40 µm mash).
Sort cells for endogenous GFP expression (after infection) or cell surface markers, as well as live-dead stain (Calcein Blue).
Sort cells into Conditioned Medium (see Section 5
) containing 10 µM of Rho-kinase inhibitor Y-27632.
CRITICAL STEP Collection in conditioned medium containing Y-27632 and the usage of freshly prepared collagen-coated plates resulted in better recovery of the sorted cells.
Plate sorted cells on freshly prepared collagen plates in conditioned medium containing Y-27632 or use for further downstream analysis (Single Cell Sequencing, RNA isolation, fixation).
If the cells are plated after sorting, change the medium 24 h after plating to conditioned medium without Y-27632. After another 48 h the medium is changed to fresh Pneumacult-Ex medium.
3.5. Freezing, Long Term Storage and Thawing
3.5.1. Freezing and Long-Term Storage: Time for Completion 30 min
Freeze a minimum of 5 × 105 cells per vial.
Instead of plating, resuspend cells in 900 µL of conditioned medium (see Section 5
) and lastly add 10% DMSO to cryovials.
Invert cryovials 5–10 times gently.
Freeze cells slowly using isopropanol boxes at −80 °C and transferred to liquid nitrogen 48 h later.
Cells can be kept at −140 °C for long-term storage and can successfully be thawed at least 1.5 years after freezing.
3.5.2. Thawing: Time for Completion 30 min
Thaw quickly in a water bath.
CRITICAL STEP To increase the cell viability and yield after thawing, PBS should be used cold and medium should be warmed no higher than room temperature.
Resuspend in 10 mL of cold PBS (containing CaCl and MgCl).
Centrifuge for 5 min, at 300× g, at 4 °C.
Remove PBS and resuspend in 10 mL fresh medium.
After thawing, plate cells on collagen coated plate. With the next passage they can again go onto uncoated dishes.
5. Reagents Setup
5.1. Coating Medium
66 µL (200 µg) of bovine collagenase (PureCol, Advanced Biomatrix, 5005-100ML, 3 mg/mL) in 5 mL of PBS to a final concentration of 40 µg/mL per 10 cm plate
Coating medium needs to be prepared fresh for every use. Diluted and stored collagen is not recommended. The PureCol collagen stock is good in the fridge for up to 1 year.
5.2. Wash Buffer
500 mL of DMEM/F12
5 mL of Penicillin Streptomycin
1 mL Amphotericin C (Fungizone) (25 ng/mL)
Wash buffer is stable at 4 °C for 3–4 weeks. Ideally wash buffer is prepared fresh every time to ensure maximum protection from fungal or bacterial contamination in the cultures.
5.3. Dissociation Buffers
5.3.1. 10× Protease Mix
100 mg Protease (Protease from Streptomyces griseus)
1 mg DNAse
9 mL of cold PBS
The 10× Protease Mix should be frozen down as 2 mL aliquots immediately after preparation. Preparation should be done on ice with cold PBS. Protease Mix is stable at −20 °C for up to two years.
5.3.2. Digest Buffer
0.2 mL Amphotericin B (2.5 µg/mL)
Digest Buffer needs to be prepared fresh before each digest. Wash buffer should be cold when preparing the digest buffer.
5.4. Pneumacult-Ex and Pneumacult-ALI
Pneumacult-Ex and Pneumacult-ALI are prepared fresh before every feed according to the manufacturer’s instructions.
49 mL Pneumacult-Ex base medium
1 mL Pneumacult-Ex supplement
100 µL Primocin
Pneumacult-Ex supplement is aliquoted upon arrival into 1 mL aliquots and stored at −20 °C. Pneumacult-Ex base medium is stored at 4 °C. The supplemented, complete medium is good for up to 2 weeks if stored at 4 °C and repeated warming and cooling is avoided.
44.5 mL Pneumacult-ALI base medium
5 mL 10× Pneumacult-ALI supplement
0.5 mL 100× Pneumacult-ALI maintenance supplement
100 µL Primocin
Pneumacult-ALI 10x Supplement is aliquoted into 5 mL aliquots upon arrival. 100× Supplement is stored at −20 °C and protected from light. Pneumacult-ALI base medium is stored at 4 °C. The supplemented, complete medium is good for up to 2 weeks if stored at 4 °C, in the dark and repeated warming and cooling is avoided.
5.5. Conditioned Medium
To prepare conditioned medium macaque BCs need to be cultured with fresh Pneumacult-Ex for 24–48 h. The medium is collected off the cultures and centrifuged at 1000× g for 5 min. The medium is collected, except the bottom 2 mL in the tube. This medium is then filtered through a 0.2 µm filter and stored at 4 °C for maximally 4 weeks. Before conditioned medium is used on the cells, the conditioned base medium is supplemented with Pneumacult-Ex supplement at a ratio of 1:20 (instead of 1:10).
5.6. Flow Cytometry Buffer
49 mL PBS (containing CaCl and MgCl)
0.5 mL Penicillin Streptomycin
0.5 mL Amphotericin B (2.5 µg/mL)
0.5 g Bovine Serum Albumin
Flow Cytometry Buffer is stable for 2 weeks at 4 °C.