A simple method to generate human airway epithelial organoids with externally orientated apical membranes

Abstract

Organoids, which are self-organizing three-dimensional cultures, provide models that replicate specific cellular components of native tissues or facets of organ complexity. We describe a simple method to generate organoid cultures using isolated human tracheobronchial epithelial cells grown in mixed matrix components and supplemented at day 14 with the Wnt pathway agonist R-spondin 2 (RSPO2) and the bone morphogenic protein antagonist Noggin. In contrast to previous reports, our method produces differentiated tracheobronchospheres with externally orientated apical membranes without pretreatments, providing an epithelial model to study cilia formation and function, disease pathogenesis, and interaction of pathogens with the respiratory mucosa. Starting from 3 × 10⁵ cells, organoid yield at day 28 was 1,720 ± 302. Immunocytochemistry confirmed the cellular localization of airway epithelial markers, including CFTR, Na⁺/K⁺ ATPase, acetylated-α-tubulin, E-cadherin, and ZO-1. Compared to native tissues, expression of genes related to bronchial differentiation and ion transport were similar in organoid and air-liquid interface (ALI) cultures. In matched primary cultures, mean organoid cilia length was 6.1 ± 0.2 µm, similar to that of 5.7 ± 0.1 µm in ALI cultures, and ciliary beating was vigorous and coordinated with frequencies of 7.7 ± 0.3 Hz in organoid cultures and 5.3 ± 0.8 Hz in ALI cultures. Functional measurement of osmotically induced volume changes in organoids showed low water permeability. The generation of numerous single testable units from minimal starting material complements prior techniques. This culture system may be useful for studying airway biology and pathophysiology, aiding diagnosis of ciliopathies, and potentially for high-throughput drug screening.

INTRODUCTION

Though organ cultures of proximal airway epithelia have been studied since the first half of the 20th century, it was not until much later that the first confluent cultures of cells isolated from animal and human airways were described (1–3). Early airway cultures lacked full differentiation. A significant advance in producing cultures that more closely mimicked the native respiratory tract mucosa was the growth of isolated guinea pig tracheal cells on porous culture supports at an air-liquid interface (ALI) (4). Though not considered three dimensional (3-D) per se, these planar cultures largely recapitulated the 3-D architecture of the airway mucosa. Subsequent advances in growth substrates, media and coculturing techniques have improved modeling of the mucociliary epithelium of the human nose, trachea and bronchi, permitting disease-relevant, in vitro studies of cellular and molecular events occurring within the mucosal lining of the airway (5–7).

Though ALI cultures have been and remain excellent models of the airway epithelium, there has been increasing interest in other 3-D culture models of airway cells. These self-organizing structures are used for modeling various organs and tissues, including tissues of complex organs like the lungs (8–10). With the exception of tissue explants, the earliest 3-D airway cell models that did not involve attachment of the cells to a supporting structure were simple spheroids generated from nasal or bronchial epithelial cells, generally obtained by mucosal brushing, but also from enzymatic digestion of surgically removed nasal polyps or fetal airway tissues (11–16). Epithelial fragments are typically released from the starting material by brief enzymatic digestion often in combination with agitation or flushing the fragments through a hypodermic needle (11, 13), though some have placed the tissue fragments directly into culture media (17). Typically, the isolated cell fragments are suspended in media and attachment to the culture vessel is prevented by the absence of culture vessel matrix coatings along with the use of mechanical rocking or repetitive shaking of the culture vessels during the initial culture period, from as few 1–4 h to as long as 10 days (13, 17). If the floating airway epithelial fragments are minimally digested and include numerous differentiated cells, spheroids showing a differentiated external surface with outwardly oriented apical membranes form within a few days. Other investigators have prepared spheroids from completely dissociated airway cells or from cells placed briefly in monolayer culture (11, 14, 15, 18). However, ciliogenesis, though still outwardly oriented, is invariably delayed taking up to 35 days. In an alternative approach, Guimbellot et al. (19), modifying a method of Müller et al. (20), allowed nasal epithelial sheets to attach to a culture vessel finding that nonadherent cells in the epithelial sheet formed nasosheroids within 2–5 days, and when transferred to suspension cultures these spheroids could be maintained for at least 12 wk. Thus, studies of airway spheroids demonstrated that dissociated or partially dissociated, nonadherent airway epithelial cells aggregate and undergo some degree of remodeling to form 3-D structures that maintain externally oriented differentiation for a significant period of time.

Recently, there has been considerable interest and experimental effort in the development of organoid cultures as models that mimic aspects of organ complexity. Simian and Bissell (21) provide an excellent general review and perspective of the history of organoid cultures. They prefer Shamir and Ewald’s (22) definition of organoids as the “most inclusive and accurate” describing intestinal organoids, stating “organoids can refer to clonal derivatives of primary epithelial stem cells that are grown without mesenchyme or can refer to epithelial-mesenchymal cocultures that are derived from embryonic stem cells or induced pluripotent stem cells.” Specifically, referring to lung organoids and acknowledging that organoid cell models do not yet fully recapitulate the cellular complexity of the lung, Barkauskas et al. (8) define lung organoids as “self-assembling structures generated from lung epithelial progenitor cells cultured in 3-D, with or without mesenchymal support cells.”

Using clonal populations of mouse and FACS-enriched human airway basal cells, Rock et al. (23) produced the first airway epithelial organoid cultures (tracheospheres). They reported that resuspending the basal cells in media mixed with soluble basement membrane constituents produced by Engelbreth-Holm-Swarm mouse sarcoma cells (Matrigel) generated organoids that developed, differentiated, and maintained an internally orientated ciliated apical surface. The development of intestinal organoids by Sato et al. (24) and the description of methods for in vitro differentiation of induced pluripotent stem cells (iPSCs) into ALI cultures with a proximal airway phenotype by Wong et al. (25) provided the framework for generating organoid cultures from embryonic stem cells (ESCs) and iPSCs as starting material. Subsequently, several groups developed protocols for producing organoids with a bronchial phenotype (26–28). Notably, each of their methods produced organoids with inward apical configuration characterized by cilia lining a central cavity. Miller et al. (29, 30) showed that cells cultured from human lung bud tip cells harvested from 12-wk gestational age fetuses formed airway organoids following Smad activation. This method also generated organoids with internal polarity.

Although an important advance, airway organoids produced using these highly specialized methods limit their broad use in personalized medicine (31). Thus, others, building on the earlier studies of Rock et al. (23) sought to produce nasal, bronchial, and tracheal organoid cell models starting from pluripotent cells obtained from human airway tissues. These organoid cell models included passaged nasal and tracheobronchial epithelial cells (32–36), conditionally reprogrammed cells nasal and bronchial epithelial cells (35, 37, 38), and cells obtained from lung tissue or lung lavage fluids (39, 40). Again, organoids produced from adult airway cells cultured under various conditions produced spheroidal structures with internally orientated apical membranes. Tan and coworkers (41) found that mixing human bronchial epithelial cells with human lung fibroblasts and human microvascular lung endothelial cells in Matrigel matrix produced cellular aggregates that reorganized into multitubular structures of varying sizes and shapes, and that these organoids expressed both proximal airway and distal lung epithelial markers. Although basal media formulations and culture additives have varied, investigators have generally grown organoids in cell culture plate membrane inserts. Organoid cultures have included growth factor reduced Matrigel or its equivalent as an additive to the media and/or as a coating placed atop the semipermeable culture insert membrane.

The aim of this study was to generate and characterize 3-D organotypic airway epithelial cultures prepared from isolated human respiratory epithelial cells, without preculture manipulations, induction of pluripotency, isolation of stem cells, or addition of mesenchymal stromal cells. We used cell signaling manipulation with only two signaling factors to promote maintenance and differentiation: R-spondin 2 (RSPO2), a canonical Wnt/β-catenin pathway agonist, and Noggin, a bone morphogenic protein (BMP) signaling antagonist (24). We demonstrate that application of these two factors is sufficient to generate airway epithelial organoids (tracheobronchospheres) directly from patient-derived cells with lasting features of differentiation, providing an alternative strategy to generate 3-D airway cultures. Furthermore, the methods described herein produced airway organoids in which the development of externally orientated polarity is favored, in contrast to previously described methods. We show that these methods also produce an apically outward phenotype when organoids are generated from serially expanded, conditionally reprogrammed cells.

MATERIALS AND METHODS

Study Approval

All protocols involving the use of human tissues and cells were approved by the University of California San Francisco Committee on Human Research. Patients with cystic fibrosis (CF) undergoing lung transplantation provided written informed consent to allow the use of leftover tissue samples. Deidentified normal airway tissues were obtained from lungs collected by the Northern California Transplant Donor Network.

Isolation of Human Airway Epithelial Cells

Human tracheobronchial epithelial (HTBE) cells were obtained from the trachea and mainstem bronchi of lungs deemed unsuitable for transplantation, and cystic fibrosis bronchial epithelial (CFBE) cells (all F508del/F508del) were collected from the mainstem and first lobar bronchi from patients undergoing lung transplantation. The surface epithelium was stripped from the specimens and the cells were isolated as previously described (5). Briefly, the airway tissue was rinsed four times in phosphate buffered saline (PBS) containing 5 mM dithiothreitol and twice in PBS alone and then incubated at 4°C overnight in PBS containing protease (Sigma type XIV, 0.4 mg/mL; Sigma-Aldrich, St. Louis, MO) supplemented with penicillin (100 U/mL), streptomycin (100 mg/L), gentamicin (50 µg/L), and amphotericin B (2.5 µg/mL). The following day, the enzyme solution was removed and replaced by a 1:1 mixture of Dulbecco’s modified Eagle’s medium and Ham’s F-12 Nutrient Mixture (DF12) containing 5% fetal calf serum (FCS) and the antibiotics listed above. The addition of FCS served to neutralize any residual protease as the cells are dislodged from the tissue by vigorous agitation. The denuded tissue was removed, and the clusters of airway epithelial cells remaining were dispersed by repeated agitation in a 10-mL pipette or by a short incubation in 2 mL of 0.05% trypsin, 0.02% EDTA in 0.9% NaCl wt/v (saline-trypsin-versene; STV). Trypsin was neutralized with DF12 containing 5% FCS. An aliquot of cells was stained with trypan blue, and total cell yield, percentage of viable cells, and the presence or absence of beating ciliated cells were assessed using a phase microscope and hemocytometer. A previous study showed that the cells isolated from postmortem tracheobronchial tissues typically consist of 70–99% nonciliated epithelial cells with ultrastructural features consistent with basal or intermediate cells, 1–24% ciliated epithelial cells, and 0–8% goblet cells (42). Samples used in these studies were limited to those with ≥90% viability. The isolated cells were aliquoted into storage vials and frozen overnight at −80°C in a cell freezing container (Thermo Fisher Scientific, Waltham, MA) containing isopropyl alcohol before transfer to liquid nitrogen for long-term storage. All of the donor cell stocks used in this study had been previously tested for their ability to generate confluent, differentiated cultures when grown as cell sheets at ALI.

Generating Conditionally Reprogrammed Cells

Conditional reprogramming of HTBE cells was performed using a modification of published methods (43, 44). Briefly, 3T3-L1 fibroblasts (ATCC, Manassas, VA; RRID:CVCL_0123; https://www.atcc.org/products/crl-1658) were suspended in F-medium [3:1 Ham's F-12/DMEM, supplemented with 5% FBS, 5 μg/mL bovine insulin, 8.4 ng/mL cholera toxin, 10 ng/mL recombinant human epidermal growth factor (R&D Systems)], 25 ng/mL hydrocortisone, 10 μM Y27632-ROCK inhibitor (Enzo Life Sciences, Farmingdale, NY), gentamicin (10 µg/mL), amphotericin B (250 ng/mL), penicillin (100 U/mL), and streptomycin (100 mg/L). The suspended cells were irradiated in a Gammacell 3000 Elan (Best Theratronics, Springfield, VA) at 30 Gy. Irradiated cells were then plated (3 × 106 per 10-cm tissue culture dish) and allowed to attach for 2 h. Next, human bronchial epithelium (HBE) cells (1.5 × 106) were added to each dish and allowed to proliferate until they reached ∼80% confluence at which time they were harvested by differential trypsinization. Conditionally reprogrammed cells (CRCs) were either subjected to an additional round of reprogramming or used to generate organoid or ALI cultures.

Air-Liquid Interface Cultures

Primary (passage [P]0) or conditionally reprogrammed HTBE cells (P2–4) were seeded onto Transwell inserts (Costar 0.4 µm pore size; 1.12 cm²; Corning, Corning, NY) at 5 × 10⁵ cells/cm² precoated with 20 µg/cm² human placental collagen (Sigma-Aldrich) and grown at an air-liquid interface as described (45). Initial plating medium consisted of a 1:1 mixture of Dulbecco’s modified Eagle’s medium and Ham’s F12 medium containing 5% heat inactivated fetal calf serum (#SH30071.01HI; Hyclone, Logan, UT) containing gentamicin (50 µg/mL), penicillin (100 U/mL), streptomycin (100 µg/mL), and amphotericin B (2.5 µg/mL; Catalog Nos. GT50, PS20, and FG70, respectively; Omega Scientific, Tarzana, CA). After 24 h, this was replaced with UNC ALI medium, which supports growth and differentiation of human tracheobronchial cells (Supplemental Table S1; see https://doi.org/10.6084/m9.figshare.14489352.v2) (6). Subsequently, medium was changed every 2–3 days. In some experiments, UNC ALI medium was supplemented with 200 ng/mL recombinant human R-spondin 2 (RSPO2; R&D Systems, Minneapolis, MN) or 25 ng/mL recombinant human Noggin (R&D Systems), separately or together.

Generating Tracheobronchial Epithelial Organoids

PureCol (Advanced BioMatrix, Inc., San Diego, CA), which consists of 97% Type I atelocollagen and 3% Type III collagen, was prepared by mixing, on ice, eight parts PureCol, one-part 10× Minimum Essential Media (ThermoFisher Scientific), and 1.1 parts 0.12 M sodium hydroxide. Matrigel (Corning, Inc., Corning, NY) was thawed on ice overnight as recommended by the manufacturer. In preliminary studies, we tested each of the collagen substrates individually as well as a 2:1 mixture of PureCol and Matrigel. Dissociated HBE or CFBE cells (2.5 × 10⁵) were added to the collagen substrates, and the cell-substrate mixtures were plated on permeable filter supports (0.4-µm pore size, 1.2 cm², Corning) that had been placed at 20°C for ∼2 h before plating. UNC ALI Medium (6) was placed in culture plate wells below the culture inserts while the upper portion of the insert containing the collagen substrate mixture was left at an air-gel interface. The cultures were grown in a tissue culture incubator gassed with 5% CO₂ and air in 80% relative humidity at 37°C. On day 14 of culture, developing organoids were recovered from collagen matrix using a 1:1 mixture of 1 U/mL Dispase (StemCell Technologies, Inc., Inc. Vancouver, BC, Canada) and 0.2% collagenase solution (Clostridium histolyticum, Sigma-Aldrich). The digestion of the collagen matrix was performed according to the manufacturer’s instructions (1 h, 4°C), and digestion of the gel with release of cell units was confirmed using a phase microscope before the process was terminated by adding a 10 mM EDTA solution. The organoids were washed twice in phosphate buffered saline (PBS) and once in warm medium with intervening centrifugation (40 g; 3 min). Next, recovered organoids were mixed with a freshly prepared collagen substrate before being re-plated as described above with the exception that the growth medium was supplemented with 200 ng/mL RSPO2 and 25 ng/mL recombinant human Noggin. The cultures were grown for an additional 7 to 31 days, depending on the experiment, though most experiments were performed on day 28. In some experiments, developing organoids harvested at day 14 were replated as above, but without RSPO2 and Noggin. Media below the insert was changed three times per week.

Overview of Cell Culture Experiments

In preliminary studies, HTBE cells (P0) were cultured at ALI to study the effects of RSPO2 and Noggin. All studies characterizing HTBE organoids were performed on organoids that were generated from cells that had been cryopreserved directly after isolation, thus also representing P0 cultures. Studies were also done to determine whether airway organoids could be generated from CRCs. Ciliary beat frequency and cilia length were compared in P0 and P2–P4 CRCs in matched cultures grown both as organoids and ALI cultures.

Counting Organoid Units and Measuring Total Cellular Protein

On culture days 7, 14, 21, and 28 developing and differentiated organoids generated from three individuals were recovered from gels as described above and diluted in 10 mL of PBS buffer. The dilute (10 µL) was pipetted onto a hemocytometer and aggregating and maturing cellular units were counted. Cellular units were defined by the organization of multiple cells into spherical three-dimensional round to oval structures with no or only small cavities (aggregating organoids) or spherical structures with multiple central cavities or a single cavity (maturing and mature organoids). Single cells or cell sheets were not counted. To measure protein concentration, initial plating aliquots and recovered organoid units were lysed in 200 µL/well RIPA Lysis and Extraction Buffer (Thermo Fisher Scientific) for 45 min on ice. Lysate was spun down at 13,000 g for 20 min and total protein concentration was determined using a commercially available assay (BCA protein assay kit; Pierce Biotechnology, Rockford, IL).

Sample Preparation for Light or Immunomicroscopy

For light microscopy, transwell inserts and organoids were fixed with 10% neutral buffered formalin for 2 h at room temperature and then held in phosphate buffered saline until further processing. For immunofluorescence microscopy, transwell filters and organoids were fixed with Dent’s fixative (1 part dimethyl sulfoxide to 4 parts methanol) and placed at −20°C for 48 h, then held in phosphate buffered saline until further processing. Organoids were recovered from collagen matrix immersing gels in Corning® Cell Recovery solution at 4°C for 1 h. After a PBS wash, dissociated organoids were collected by centrifugation (40 g; 3 min). Supernatant was drawn off and cellular aggregates were transferred into disposable base molds (7 mm × 7 mm × 5 mm, Thermo Fisher Scientific) and mixed with ∼ 200 µL of agarose gel (SeaKem LE, Lonza, Rockland, ME). Agarose gel containing the organoids was polymerized at −20°C for 11 min. Transwell filters and hardened organoid gels were embedded in paraffin for further studies. Hematoxylin and eosin (H&E)-stained sections (5 µm) were used for histological evaluation.

Immunomicroscopy

Immunofluorescence staining was performed on paraffin-embedded sections or directly on cells attached to a transwell filter for en face imaging. Immunohistochemical staining was performed on paraffin-embedded sections. Antibody verification included positive and negative controls. Staining was performed using commercially available antibodies as listed in Table 1. In all immunohistochemical staining experiments appropriate localization of primary antibody staining was verified using intact tracheobronchial airway specimens. The absence of nonspecific staining was confirmed by replacing primary and secondary antibodies with appropriate nonimmune sera.

Table 1.

Primary antibodies, vendors, dilutions/concentration and Research Resource Identifiers

Antibody	Vendor; Catalog Number	Dilution/Concentration	RRID
Immunofluorescence microscopy for en-face imaging of cell culture inserts
Mouse anti-acetylated-α-tubulin	Sigma-Aldrich Merck, St. Louis, MO; T6793	1:700–1:1000	AB_477585
Rabbit anti-cytokeratin 5	ThermoFisher Scientific, Waltham, MA; PA1-37974	1:500	AB_2134167
Rabbit anti-human club cell protein	BioVendor, Asheville, NC; RD181022220	1:1000	AB_344578
Rabbit anti-Muc5AC	Santa Cruz Biotechnology, Dallas, TX; sc-20118	1:500	AB_2146854
Mouse anti-Muc5B	Santa Cruz Biotechnology, Dallas, TX; sc-393952	1:500	AB_2876857
Immunofluorescence microscopy of paraffin sections
Mouse anti-acetylated-α-tubulin	Sigma-Aldrich Merck, St. Louis, MO; MABT868	1 µg/mL	AB_2819178

RRID, Research Resource Identifiers.

Immunofluorescence staining on cells attached to transwell filters.

Primary cells grown in ALI culture were washed 3× for 5 min with PBS before fixation in 4% paraformaldehyde at 4°C for >24 h. The culture filters were removed from the insets using a scalpel, divided into multiple parts and used for different combinations of stains. Filter parts were washed 2× for 30 min in PBST (0.1% Triton X-100 in PBS) and blocked in PBST-CAS [90% PBS containing 0.1% Triton X-100, 10% CAS-Block reagent (Thermo Fisher Scientific)] for 30 min–1 h at room temperature (RT). Primary antibodies were incubated in 100% CAS-Block reagent overnight at 4°C followed by 3× 15-min washes with PBS. Secondary antibodies were used at 1:250 or 1:500 dilution in CAS-Block reagent and incubated at RT for 2 h or overnight at 4°C, respectively. Secondary antibodies (Thermo Fisher Scientific) included the following: AlexaFluor 555-labeled goat anti-mouse antibody (Cat. No. A-21422; AB_25358442), AlexaFluor 488-labeled goat anti-rabbit antibody (Cat. No. R37116; AB_2556544), AlexaFluor 488-labeled donkey anti-mouse antibody (Cat. No. R37116; AB_2556542), and AlexaFluor 405-labeled goat anti-mouse antibody (Cat. No. A31553; AB_221604). After staining with secondary antibodies, samples were washed 3× for 15 min with PBS. ProLong Gold Antifade Mountant with DAPI (Thermo Fisher Scientific) was used to label nuclei. Actin was stained by incubation (30–120 min at RT) with AlexaFluor 488- or 647-labeled Phalloidin (1:40 in PBST; Thermo Fisher Scientific). Confocal imaging was conducted using a Zeiss LSM700 and Zeiss Zen software (Carl Zeiss Microscopy, White Plains, NY; RRD:SCR_013672; https://www.zeiss.com/microscopy/int/products/microscope-software/zen.html). Confocal images were adjusted for channel brightness/contrast and Z-stack projections using Image J, an open-source processing software (ImageJ, version: 2.0.0-rc-23/1.49m; RRID:SCR_003070; version: 2.0.0-rc-23/1.49m; https://imagej.net/).

Immunofluorescence staining on paraffin embedded sections.

Unstained 5-µm sections were deparaffinized and treated with heat induced epitope retrieval to unmask antigens, using a heat retrieval solution (Reveal Decloaker 10×, Biocare Medical, Concord, CA) for 10 min at 125°C followed by cooling at 21°C for 10 min. After permeabilization with 0.1% Triton X-100 for 20 min protein blocking was performed using 5% normal goat serum. Sections were then incubated with primary antibodies overnight at 4°C. Alexa Fluor 488 Goat Anti-Mouse (Thermo Fisher Scientific; Cat. No. A-11001; AB_2534069) or Alexa Fluor 568 Goat Anti-Rabbit (Thermo Fisher Scientific; Cat. No. A-11001; AB_143157) was used to visualize binding of antibodies and sections were stained with secondary antibody for 2 h at RT. Prolong Gold Antifade Reagent with DAPI (Thermo Fisher Scientific) to identify cell nuclei was used as mounting medium. Images were obtained on a spinning disk confocal unit (CSU22, Yokagawa, Electric Corporation, Tokyo, Japan) and equipped with Micro Manager version 1.4 (Fuzhou Tucsen Photonics Co., Ltd Fuzhou Fujian, PRC; www.tucsen.com; https://micro-manager.org/download_MicroManager_Latest_Release) or a Zeiss LSM700 and Zeiss Zen software (Carl Zeiss Microscopy, White Plains, NY; RRD:SCR_013672; https://www.zeiss.com/microscopy/int/products/microscope-software/zen.html). ImageJ (RRID:SCR_003070; version: 2.0.0-rc-23/1.49m; https://imagej.net/) was used for image editing.

Electron Microscopy

Organoids were fixed for 90 min in 2.5% glutaraldehyde buffered with a 0.2 M cacodylate buffer. The cultures were washed in 0.1 M cacodylate buffer followed by post fixation in a 1:1 solution of 2% aqueous osmium tetroxide and 0.2 M cacodylate buffer for 2 h at 4°C followed by a second 0.1 M cacodylate buffer wash. Culture were then dehydrated of cultures with ethanol at 35% for 5 min, 50% for 5 min, 70% for 5 min, 95% for 5 min, 100% for 5 min, propylene oxide 15 min and 1:1 solution of propylene oxide/epon resin for 3 h at room temperature. Organoids were passed through three changes of Epon 812 resin. Drops of fresh Epon 812 were placed in clear polyethylene capsules, and the cultures were transferred to the appropriate capsule to embed. The blocks were cured for 48 h at 60°C. A microtome was used to cut 100 nm sections, which were placed on grids. Images were obtained using a FEI Tecnai G2 (Hillsboro, OR) electron microscope.

Quantitative RT-PCR

Before total RNA extraction from ALI cell cultures, filters were washed 3× for 5 min with PBS and removed from the insets using a scalpel cleaned with RNase away (MbP No. 7002). Before RNA extraction from organoids, cellular units were recovered from collagen matrix immersing gels in Corning Cell recovery solution at 4°C for 1 h. Dissociated organoids were transferred to RNase-free vials (Thermo Fisher Scientific, Ambion, MbP No. AM12400), spun down after 3× 6 min PBS washes and removed from the vials with a scalpel cleaned with RNase AWAY (Thermo Fisher Scientific). Before RNA extraction from native bronchial surface epithelium the tissue was washed for 3× for 10 min with PBS and a 0.3-cm² tissue section consisting of the surface epithelium only was removed using a scalpel, which had been previously cleaned with RNase Away. The RNeasy Mini Kit (QIAGEN, Hilden, Germany) was used, and the cells were collected in RLT buffer + β-mercaptoethanol (10 μL/mL), vortexed for 2 min, and lysed using QIAshredder (QIAGEN) columns. RNA was collected in UltraPure water (Thermo Fisher Scientific) and used for cDNA synthesis with iScript cDNA Synthesis Kit (Bio-Rad). Using SsoAdvanced Universal SYBR Green Supermix (Bio-Rad), quantitative RT-PCR (qPCR) was conducted on a CFX Connect Real-Time System (Bio-Rad) in 96-well PCR plates (Brand Tech Scientific, Essex, CT). Experiments were conducted in biological and technical triplicates and normalized by GAPDH expression levels. Expression levels were analyzed in Microsoft Excel (Microsoft Corporation, Redmond, WA; RRID:SCR_016137; https://www.microsoft.com/en-gb/), and graphs were generated using R (Project for Statistical Computing, RRID:SCR_001905; https://www.r-project.org). Genes studied and corresponding qPCR primer sequences are listed in Table 2.

Table 2.

Primer sequences for real-time qPCR

Gene	Primer Sequences (5′ to 3′)
Markers of differentiation
FOXJ1	hqFoxJ1-F1: ATCTACAAGTGGATCACGGAC hqFoxJ1-R1: GAGGCACTTTGATGAAGCAC
MCIDAS	hqMcidas-F1: TGAGCAATACTGGAAGGAGG hqMcidas-R1: CCTGTTTCTGGGTCAATGTC
TP63	hqP63-F1: CGTGAGACTTATGAAATGCTG hqP63-R1: TGAAGATGGAGACTGTATTGAG
KRT5	hqKrt5-F1: GCAGTACATCAACAACCTCAG hqKrt5-R1: CTACATCCTTCTTCAGCATCAC
SCGB1A1	hqCCSP-F1: CCTGATCAAGACATGAGGGA hqCCSP-R1: TAATTACACAGTGAGCTTTGGG
CGRP	hqCGRP-F1: GGACTATGTGCAGATGAAGG hqCGRP-R1: CGTGTGAAACTTGTTGAAGTC
Secretory/goblet cells
MUC5AC	hqMuc5AC-F1: GGAACTTCAACAGCATCCAG hqMuc5AC-R1: GAGCATACTTCTCATTCTCCAC
MUC5B	hqMUC5B-F1: TGTTCCTCAACTCCATCTACAC hqMUC5B-R1: CTGACAAACACCTGCATGAG
MUC1	hqMUC1-F1: AACCTCCAGTTTAATTCCTCTC hqMUC1-R1: AATTGTACCACCACAGATCC
Ion channels
TMEM16	hqTMEM16-F1: AGACTCTCCTATCCCTTCTCC hqTMEM16-R1: TCTCATAGACAATCGTGCTCC
SCNN1A	hqENaC-F1: TCTCTGCTGGTTACTCACGA hqENaC-R1: GCTCCTTGAAGAAGATGTTGAC
ATP1A1	hqNaKATPase-F1: ATGTCTATGATTGACCCTCCC hqNaKATPase-R1: CCCTCTGATATGATGCCCAC
CFTR	hqCFTR-F1: CCTATGACCCGGATAACAAGGA hqCFTR-R1: GAACACGGCTTGACAGCTTTA
ATP12A	hqATP12a-F1: GGCACAGACATCATTATGGGTC hqATP12a-R1: GATGCAGACTTGTCGCTGG
Reference gene
GAPDH	hqGAPDH-F1: GGAGCGAGATCCCTCCAAAAT (Tm 61.6) hqGAPDH-R1: GGCTGTTGTCATACTTCTCATGG (Tm 60.9)

FOXJ1, Forkhead box J1; MCIDAS, multiciliate differentiation and DNA synthesis associated cell cycle protein; TP63, tumor protein P63; KRT5, keratin 5; SCGB1A1, secretoglobin family 1A member 1; CGRP, calcitonin gene-related peptide; MUC5AC, Mucin 5AC; MUC5B, Mucin 5B; MUC1, Mucin 1; TMEM16, transmembrane protein 16; SCNN1A, sodium channel epithelial 1 subunit alpha; ATP1A1, ATPase Na⁺/K⁺ transporting subunit alpha 1; CFTR, cystic fibrosis transmembrane conductance regulator; ATP12A, ATPase H⁺/K⁺ transporting, nongastric, alpha polypeptide; GAPDH, glyceraldehyde 3-phosphate dehydrogenase.

Cross-Sectional Diameter and Cross-Sectional Surface Area Calculation

Light microscopy images of formalin-fixed, paraffin-embedded organoids generated from three CF individuals with the F508del/F508del genotype and three non-CF individuals taken at ×400 magnification were analyzed for cross-sectional diameter and cross-sectional surface area using a calibrated scale with ImageJ (RRID:SCR_003070; version: 2.0.0-rc-23/1.49m; https://imagej.net/).

High-Speed Video Imaging of Cilia

High-speed videos of cilia in organoid and ALI cultures from three matched individuals were obtained by differential interference contrast (DIC) imaging on an Axiovert inverted microscope with a ×63 water objective lens (Zeiss) and a high-speed camera (Phantom Miro, Vision Research, Wayne, NJ) at 1,000 fps. Videos and images were evaluated for ciliary beat frequency (CBF) in beats per second (Hz) and cilia length with ImageJ, RRID:SCR_003070; version: 2.0.0-rc-23/1.49m; https://imagej.net/).

Osmotic Response Studies

A perfused microchannel device that was microfabricated with polydimethylsiloxane, as reported previously (46), was used to immobilize organoids to study their volume response to osmotic challenges. Organoids were superfused continuously in the microchannel device under conditions in which >95% exchange of perfusate fluid was accomplished in under 2 s. Perfusate solutions consisted of PBS (∼300 mOsm), PBS diluted 1:1 with distilled water (150 mOsm), and PBS with added mannitol to give osmolalities of 500 or 1300 mOsm. Low-magnification images of multiple immobilized organoids were recorded by transmitted light microscopy at a rate of 1 Hz to deduce the kinetics of organoid volume by image analysis, as described (46), and transepithelial osmotic water permeability was computed from the kinetics of organoid volume, osmotic gradient size, and organoid size.

Statistics

Statistical evaluation of organoid diameter, cilia length, and CBF were done by Student’s t test (http://www.physics.csbsju.edu/stats/t-test.html). Statistical significance was defined at a P-value ≤ 0.05. Boxplots were generated in R (R Project for Statistical Computing, RRID:SCR_001905; https://www.r-project.org). qPCR samples in Supplemental Fig. S1 (see https://doi.org/10.6084/m9.figshare.14489169) were compared using two-tailed Wilcoxon rank-sum tests, without adjustment for multiple comparisons (https://astatsa.com/WilcoxonTest/). Ciliation and cavitation analysis was evaluated by χ²-test (http://www.physics.csbsju.edu/stats/contingency.html) and subjected to Bonferroni correction for multiple comparison. For ciliary length and CBF, the F-test function was used to determine if the variances of the two samples were unequal.

RESULTS

Choosing Signaling Manipulations to Promote Basal Cell Expansion and Maintenance

Establishment of airway organoids and tracheobronchospheres depends on the availability of sufficient numbers of airway basal cells, which are the adult tissue-specific stem cells. We sought to generate organoids without the use of manipulations such as induction of pluripotency, isolation of stem cells, or addition of mesenchymal stromal cells to the cultures. In many protocols, basal cells are either induced from iPSCs using a stepwise differentiation protocol or purified from large quantities of isolated airway epithelial cells. As these steps represent a significant limitation for establishing airway mucociliary organoids from small amounts of patient material, we wondered if signaling manipulations that promote basal cell expansion and maintenance could be used during organoid culture. Canonical Wnt/β-catenin signaling was shown to promote the expression of the basal cell master transcription factor ΔN-TP63, which is necessary and sufficient to suppress differentiation from basal cells, and which was shown to upregulate proliferation (47–49). Additionally, it was demonstrated that the inhibition of BMP/Smad signaling is required for basal cell expansion in vitro, and that basal cells are devoid of nuclear Smad localization in the mouse airway in vivo (50). Furthermore, inhibition of BMP signaling was shown to enhance the differentiation of cultured airway epithelial cells without skewing cell type composition, likely due to the greater availability of basal cells for the generation of mature epithelial cell types (51, 52).

Motivated by these observations, using the ALI cell model we tested if the combined stimulation of canonical Wnt signaling and inhibition of BMP signaling could promote basal cells more efficiently than previously reported for each signaling manipulation alone. We chose the addition of the Wnt agonist RSPO2 and the BMP antagonist Noggin to manipulate signaling. Furthermore, as promoting basal cells can suppress the specification of differentiated epithelial cell types, we assessed the effects of RSPO2 and Noggin treatment when applied at the onset of the in vitro cultures as well as their application at a later phase of the differentiation protocol (after initial cell fate specification of epithelial cells). We hypothesized that delaying the addition of RSPO2 and Noggin could have a positive effect on basal cell maintenance, but without preventing specification and differentiation of ciliated and secretory epithelial cell types.

We used primary HTBE cells grown in ALI culture, then analyzed differentiation as well as effects on the basal cell population by immunofluorescence (IF) and confocal microscopy, and by qPCR (Supplemental Fig. S1). Treatment of cells during ALI days 1–7 and analysis at ALI day 7 confirmed previously published effects on HTBE cells: IF revealed that RSPO2 treatment reduced the numbers of differentiating multiciliated cells (MCCs; Ac.-α-tub. staining) and club cells (marked by strong CC10 expression), while Noggin treatment promoted differentiation of MCCs and club cells (Supplemental Fig. S1A). Additionally, mucin staining (Muc5ac and Muc5b) was reduced in RSPO2-treated samples and remained largely unchanged after Noggin treatment (Supplemental Fig. S1A). In line with the IF data, qPCR analysis showed that RSPO2 reduced the expression of all differentiation markers (FOXJ1 and MCIDAS for MCCs, SCGB1A1 for club cells, and MUC1, MUC5AC, and MUC5B for mucous secretory cells) and upregulated expression of definitive airway basal stem cell markers (TP63 and KRT5), while differentiation markers and TP63 were slightly upregulated in Noggin-treated samples (Supplemental Fig. S1B). Combined application of RSPO2 and Noggin during the initial phase of ALI differentiation largely resembled the effects of RSPO2 treatment alone, suggesting that the stimulation of Wnt signaling had a dominant effect on differentiation ability from basal cells (Supplemental Fig. S1B).

In contrast to treatments applied during the specification phase of ALI culture, the application of RSPO2, Noggin, or RSPO2 + Noggin after cell fate specification (during ALI days 21–28) did not affect the formation of MCCs or expression of FOXJ1, which is required for MCC maintenance, while RSPO2 and RSPO2 + Noggin treatments both led to an increase in TP63 and KRT5 expression (Supplemental Fig. S1, C and D). This indicated enhanced maintenance and/or expansion of basal cells. Interestingly, the number of cells expressing CC10 at high levels was diminished and SCGB1A1 was decreased in RSPO2 and RSPO2 + Noggin treated cultures (Supplemental Fig. S1, C and D). These results suggest either a reduced maintenance of club cells or simply an attenuation of SCGB1A1/CC10 expression from cells which maintain club cell identity. In contrast, mucous secretory cells were not negatively affected by the same treatments as Muc5b/MUC5B expression was increased (Supplemental Fig. S1, C and D), a finding previously described following stimulation of canonical Wnt signaling (47).

Taken together, these experiments confirm previously published results of Wnt stimulation and BMP inhibition on differentiation and basal cell maintenance. Furthermore, they indicate that the use of RSPO2 + Noggin likely promotes organoid generation and maintenance when applied after an initial phase of differentiation circumventing the inhibitory effects of Wnt/ß-catenin on epithelial cell fate specification.

Optimization of Conditions for Generating Airway Epithelial Organoids

Primary airway epithelial organoids were cultured from wild type (wt) HTBE cells and epithelial cells isolated from the large bronchi of patients with CF, homozygous for the F508del mutation. Organoid growth conditions, including cell matrix composition and volume, as well as seeding density, were optimized using HTBE cells. PureCol, a bovine collagen solution composed of 97% type I and 3% type III collagens; Matrigel, a heterogeneous mixture of matrix proteins and growth factors; and a 1:2 mixture of both solutions were tested. PureCol alone produced a more tubular conformation of airway epithelial cell clusters, which was not desirable for further studies (data not shown). Airway organoids seeded in a Matrigel or a mixed matrix consisting of 2 parts PureCol and 1-part Matrigel formed spherical cell units. Optimal growth in the mixed collagen matrix was found at a seeding density of 2.5 × 10⁵ cells/cm² (300,000 cells per 1.2 cm² Transwell insert). At a seeding density of 0.83 × 10⁵ cells/cm² relatively few organoids developed, while at 4.17 × 10⁵ cells/cm² cells formed confluent cell sheets along with organoid units.

To achieve optimal organoid differentiation, developing organoids were harvested at 2 wk and then replated and cultured in the presence of RSPO2 and Noggin. At this time the initial gels had begun to disintegrate, a limitation that was remedied by transferring the organoids to freshly prepared matrix. However, a consequence of replating was the loss of ∼60% of the organoids (Fig. 1A). On average, the final organoid yield in the mixed collagen matrix on day 28 was 1,720 ± 302 (n = 15 cultures, from three different donors), significantly greater than in organoids cultured in Matrigel alone (360 ± 235 cell units, n = 15 cultures, from three individuals, P ≤ 0.005). Organoid protein content steadily increased over the course of 28 days of culture. Notably, protein content increased from day 14 to 21 despite the loss of organoid resulting from replating, indicating ongoing growth and differentiation (Fig. 1B). This increase was attributed to the stimulatory effects of RSPO2 + Noggin on organoids growth and differentiation, as replating organoids in their absence resulted in a loss of 84% and 88% of cellular units at days 21 and 28, respectively, along with an associated decrease in organoid protein content [698 ± 71 µg/mL at day 21 and 640 ± 75 µg/mL at day 28 (each n = 6 from three individuals)].

Figure 1.

Characterization of organoids grown from primary human airway epithelial cells. A: organoid cultures (n = 5 cultures each from three individuals) were initiated with 2.5 × 10⁵ cells/cm², and the number of cellular units was determined at weekly intervals. Media changes and additions of RSPO2 and Noggin were done on day 14. B: total protein of HTBE cells at seeding and at indicated days of organoid development. Despite loss of organoid cellular units after collection of organoids and replating, total organoid protein increased following the addition of RSPO2 + Noggin indicating their positive effect on organoid vitality. For each time point one sample each using seeding aliquots from three individuals was analyzed. C: brightfield images of H&E-stained sections of organoids grown from HTBE cells. The amorphous background material present in some panels is gelatin used as a carrier during tissue processing. At day 7 the HTBE cells have aggregated into a small spherical organoid is a small internal cavity developing. At day 14 organoids have enlarged and multiple small cavities are present. Indicative of remodeling, the cavities contain dying or degenerating cells and elsewhere, pyknotic nuclei of apoptotic cells are present. At day 21 in organoid shown the central cavities have coalesced but contain cellular debris and the outer wall remains thickened. At day 28 this mature organoid has a single cavity and its surface is composed of multiciliated cells displaying outward polarity (inset). D: brightfield image of H&E-stained section of 28-day organoid grown from F508del/F508del CFBE cells. This organoid has two cavities both with internally orientated ciliated apical membranes. E: brightfield image of H&E-stained section of 28-day organoid grown from F508del/F508del CFBE cells. This organoid with a single cavity shows outwardly orientated polarity except for a small focus of cavitary cilia (arrows). Widened intracellular spaces may be cleavage sites where this organoid is undergoing ongoing remodeling towards complete external apical polarity. F: mean numbers of wt and CF organoids (average percentage per donor) displaying various locations of cilia at culture day 28 (sample size indicated, wt, n = 120 from three individuals with three technical replicates from one donor and one technical replicate from two donors; F508del/F508del, n = 80 from two individuals with one donor with two technical replicates and one donor with one technical replicate). G: frequency of organoids grown from wt and F508del/F508del cells displaying solid morphology (average percentage per donor), a single cavity or multiple cavities at day 28 of culture (sample size indicated, wt, n = 120 from three individuals with three technical replicates from one donor and one technical replicate from two donors; F508del/F508del, n = 80 from two individuals with one donor with two technical replicates and one donor with one technical replicate). H: comparison of cross-sectional diameter of formalin-fixed, paraffin-embedded wt organoids and F508del/F508del CF organoids at indicated days of culture. Sample size as indicated on the graph; all obtained from three biological replicates. * P ≤ 0.003; ** P ≤ 0.002; *** P ≤ 0.001; ns: not significant. CF, cystic fibrosis; CFBE, cystic fibrosis bronchial epithelial; H&E, hematoxylin and eosin; HTBE, human tracheobronchial epithelial; RSPO2, R-spondin 2; wt, wild type.

Airway Epithelial Organoid Morphology and Polarity

Figure 1C shows the typical appearance of wild type (wt) organoids at weekly intervals to day 28, and Fig. 1, D and E, show CF organoids at day 28. One week after seeding, the organoids consisted of small spherical clusters of cells with some showing the beginnings of internal cavities. By 14 days, the organoids typically showed multiple small cavities with sparse cilia growth on their external organoid surface, and phase-contrast microscopy revealed beating cilia (not shown). At this stage the organoids are undergoing remodeling, and the small cavities contained dying or degenerating cells with solid areas showing apoptotic cells. Although the organoid walls were cellular and variably thick, larger cavities, often single, were evident by day 21. By day 28, organoid remodeling was generally complete. A majority (77%; Fig. 1F) of wt organoids had a single central cavity surrounded by thin walls. In contrast, organoids generated from F508del/F508del CFBE cells were more likely to have more than one cavity (Fig. 1, D and F). At 28 days, cilia were observed in 97% of wt organoids (n = 120, from three individuals) and 85% of CF organoids (n = 80, from three individuals), with the majority located exclusively on the external surface (Fig. 1G, outward). Cavitary cilia in the absence of external cilia were rarely seen in wt organoids (1%) but present slightly more frequently in CF (10%) (Fig. 1G, inward). In a substantial percentage of ciliated organoids (wt, 38%; CF, 18%), cilia were seen externally as well as on the luminal surfaces of the cavities. However, in these organoids, the cavitary cilia were generally sparse (Fig. 1E, 1G, both). Cilia were undetectable in 3% of wt and 15% of CF organoids. On average, throughout the culture period F508del/F508del organoids were larger than wt organoids (Fig. 1H). For example, at day 28, formalin-fixed, paraffin-embedded wt organoids ranged from ∼50 to 250 µm in diameter with average diameter of 113.8 ± 4.5 µm while the CF organoids had a diameter of 144.0 ± 6.0 µm (n = 120, from three individuals; P ≤ 0.001). The high density of differentiated ciliated cells as well as the emergence of polarized epithelia distinguished these organoids from cultures grown without the addition of RSPO2 and Noggin, which lacked signs of airway epithelial differentiation, ciliation, and epithelial polarization (Supplemental Fig. S2; see https://doi.org/10.6084/m9.figshare.15262320), as well as from cultures grown with RSPO2 or Noggin alone, each condition only showing sparse ciliation [Supplemental Fig. S3 (see https://doi.org/10.6084/m9.figshare.15262419) and Supplemental Fig. S4 (see https://doi.org/10.6084/m9.figshare.15262521)].

Airway Epithelial Organoid Fine Structure and Expression of Markers of Airway Differentiation

Electron microscopy of cilia at day 28 showed basal bodies docked to apical membranes (Fig. 2A), and axonemal cross sections demonstrated the usual microtubule arrangement of nine doublets surrounding a central pair, dynein arms and radial spokes (Fig. 2B). Confocal micrographs showed the cilia marker Ac.-α-tubulin at externally orientated apical membranes (Fig. 2C), or sometimes at the apical membranes of internal cavities displaying cilia (not shown). In terms of other differentiation markers, dual staining at day 28 revealed CFTR protein localized at the apical (outwardly directed) membrane and Na⁺/K⁺ ATPase along basolateral membranes (Fig. 2, D and E). Higher resolution of the apical location of CFTR is also shown in Fig. 2D along with cell-cell connections visualized by ZO-1. Prominent staining along organoid basolateral membranes for E-cadherin identified adherens junctions (Fig. 2F). Thus, the organoids developed a differentiated, polarized epithelium typically with outwardly facing apical membranes.

Figure 2.

Fine structure of organoid cilia and markers of airway differentiation examined with electron microscopy and immunomicroscopy. A: ultrastructure of apical membrane of a 28-day organoid shows basal bodies docked to apical membranes. B: axonemal cross sections demonstrate the usual microtubule arrangement of nine doublets surrounding a central pair, dynein arms, and radial spokes. C: low and high power (indicated by yellow box) confocal micrographs of single cavity wt organoid show strong immunostaining for the cilia marker Ac.-α-tubulin (green) at externally orientated apical membranes. Cell-cell connections are visualized by ZO-1 (magenta). D: similarly, CFTR is present at the outwardly directed apical membrane. CFTR (green), ZO-1 (magenta), and nuclei (DAPI, blue) with higher power view indicated by yellow box. E: dual staining reveals CFTR protein localized at the apical membrane and Na⁺/K⁺ ATPase detected along basolateral membranes. CFTR (green), Na⁺/K⁺ATPase (magenta), and nuclei (DAPI, blue). F: prominent staining along the organoid basolateral membranes for E-cadherin identified adherens junctions. E-cadherin (magenta) and nuclei (DAPI, blue). wt, wild type; Ac, acetylated; CFTR, cystic fibrosis transmembrane conductance regulator; DAPI, 4′,6-diamidino-2-phenylindole; ZO-1, zonula occludens-1.

Comparison of Cilia in Airway Epithelial Organoid and ALI Cultures

Using high-speed video imaging, we compared ciliary beat frequency (CBF) and ciliary length in primary (P0) organoid and ALI cultures generated from matched HTBE cells (Table 3). Cilia length of P0 organoids (6.1 µm) and ALI cultures (5.7 µm) was not significantly different. However, CBF in primary ALI cultures was significantly reduced (P < 0.005) when compared to that in the P0 organoids. Conditionally reprogrammed HBTE cells (CRCs) generated from the same individuals studied at P0 were used to generate organoids and ALI cultures at P2–4. Like the P0 organoids, organoids derived from CRCs demonstrated externally orientated, ciliated apical membranes (Supplemental Videos S1–S4; see https://doi.org/10.6084/m9.figshare.14489208; https://doi.org/10.6084/m9.figshare.14489211; https://doi.org/10.6084/m9.figshare.14489214; https://doi.org/10.6084/m9.figshare.14489220). Cilia length and CBF of organoid and ALI cultures were compared between their respective primary and passaged cultures and between organoid and ALI cultures grown at the same passages. Data are summarized in Table 3. Cilia length of organoid cultures was decreased at P2, and cilia shortening continued with subsequent passaging. Cilia length of ALI cultures was maintained at P2, but progressively decreased at P3 and P4. Comparing cilia length between the CRC organoid and planar ALI cultures at identical passages revealed that organoid cilia were significantly shorter (P < 0.005). For organoid cultures, CBF increased after each passage. Conditionally reprogrammed ALI cultures showed a similar but less pronounced pattern with CBF increasing from 5.3 ± 0.8 Hz at P0 (n = 15), to 8.8 ± 1.7 Hz (n = 15) at P4 (P < 0.005). Overall, CBF was significantly higher (P < 0.005) in organoid cultures when compared to ALI cultures at P0 and P2–P4 (P < 0.005).

Table 3.

Cilia length and beat frequency

	Organoids		Planar ALI Cultures
Passage Number	Ciliary Beat Frequency (Hz)1,3	Cilia Length (µm)2,4	Ciliary Beat Frequency (Hz)1,5	Cilia Length (µm)2,6
0	7.7 ± 0.3 (n = 30)	6.1 ± 0.2 (n = 30)	5.3 ± 0.8 (n =15)	5.7 ± 0.1 (n = 14)
2	9.7 ± 0.4 (n = 29)	4.4 ± 0.1 (n = 29)	5.7 ± 1.5 (n = 15)	5.6 ± 0.2 (n = 15)
3	11.8 ± 0.7 (n = 17)	3.8 ± 0.1 (n = 17)	6.4 ± 0.5 (n = 14)	5.0 ± 0.1 (n = 14)
4	14.3 ± 0.6 (n = 11)	3.5 ± 0.2 (n = 13)	8.8 ± 0.5 (n = 15)	4.2 ± 0.1 (n = 14)

Values are means ± SE from cultures obtained from three individuals. Passage number 0 refers to primary cultures. All subsequent passaged cultures were generated by conditional reprogramming. ALI, air-liquid interface.

¹Ciliary beat frequency of organoids was statistically different from ciliary beat frequency of planar ALI cultures comparing all corresponding passages (P < 0.005).

²Cilia length of organoids was statistically different from cilia length of planar ALI cultures at corresponding passages 2 through 4 (P < 0.005).

³Quickening of ciliary beat frequency of organoids was statistically different in passages 2, 3, and 4 compared to P0 (P < 0.005).

⁴Shortening of ciliary length was statistically different in passages 2, 3, and 4 compared to P0 (P < 0.005).

⁵Quickening of ciliary beat frequency was statistically different in P4 compared to P0, P2, or P3 (P < 0.005).

⁶Shortening of ciliary length was statistically significant between P0 compared to P3 or P4 (P < 0.005).

Response of Airway Epithelial Organoid Volume to Osmotic Gradients

As an example of a physiological application involving measurement of organoid function, organoid volume response to osmotic challenge was measured to determine their water permeability. Organoids were immobilized in a pinball-like array of micropillars in a microchannel device with continuous fluid superfusion and rapid solution exchange (Fig. 3A and Supplemental Video S5; see https://doi.org/10.6084/m9.figshare.14489226). Organoid swelling and shrinking were seen in response to superfusion with hypoosmolar and hyperosmolar solutions, respectively. Figure 3B shows the deduced time course of relative organoid volume in response to osmotic challenges that drive water influx into the luminal space resulting in organoid swelling, and, conversely, water efflux and organoid shrinking. These experiments demonstrate the osmotic intactness of the organoid preparation and give a transepithelial osmotic water permeability coefficient (P_f) of ∼0.002 cm/s as computed from the time course data and organoid size. This low P_f value suggested the absence of aquaporins in the cell luminal and/or basolateral membranes; for comparison, P_f was estimated to be 0.02 cm/s in nasopharyngeal epithelium in mice in vivo (53) and 0.018 cm/s in human spheroid-shaped human nasal epithelial explant cultures (54).

Figure 3.

Organoid volume response to osmotic gradients. A: transmitted light micrographs of immobilized organoids during superfusion with PBS (300 mOsm) and after perfusion with hypo-osmolar PBS (150 mOsm) and then hyperosmolar PBS (500 or 1,300 mOsm). Three organoids are shown, with four micropillars surrounding each organoid and perfusion from right to left. See Supplemental Movie 1 for full time course. B: representative organoid volume responses to changes in perfusate osmolality from 300 to 150 mOsm and from 300 to 450 mOsm.

Gene Transcript Expression in Airway Epithelial Organoids, ALI Cultures, and Native Airway Epithelium

Comparison of mucociliary gene expression of organoids was compared to that in native human bronchial epithelium and HTBE ALI cultures (Fig. 4). Using qPCR, we studied genes related to tracheobronchial epithelial cell differentiation including FOXJ1 (multiciliated cells), TP63 and KRT5 (basal cells), SCGB1A1 (encoding CC10), as well as genes encoding airway mucins (MUC1, MUC5AC, and MUC5B). FOXJ1 transcript was expressed in both culture conditions at comparable levels to that seen in the native epithelium. Expression of the basal cell marker TP63 was similar in ALI cultures and native tissue, which was greater than the expression in organoids. KRT5 was highest in the ALI cultures but similar in organoid cultures and native tissue. Transcript expression of SCGB1A1, though robust in both culture conditions, was even greater in the native tissue. Transcript expression of the cell surface-associated Mucin 1, MUC1, was greater in organoids than in native epithelium and ALI cultures, while transcripts of the secreted mucins MUC5AC and MUC5B were greatest in native tissues. Also, we measured transcript expression of relevant ion channels and pumps, including CFTR, calcium-activated chloride channel TMEM16A, the alpha subunit of the epithelial sodium channel (ENaC, SCNN1A), sodium-potassium pump (NaKATPase, ATP1A1), and the alpha subunit of the hydrogen-potassium ATPase (ATP12A). CFTR transcript expression was greatest in the native epithelium. However, organoids showed robust expression of CFTR and other ion transporters, similar to or exceeding their expression in ALI cultures, with the exception of ATP12A, which was expressed more in ALI cultures than in organoids or in airway tissue.

Figure 4.

Comparison of gene expression in organoids (at day 28 of culture), ALI cultures (at day 28 of culture), and airway tissue. Quantitative RT-PCR of gene expression comparing to human airway tissue. Boxplots were generated in R. The line represents the median; 50% of values are represented by the box; 95% of values are presented within the whiskers; values beyond 95% are depicted as outliers. (n = 3 biological replicates per condition). ALI, air-liquid interface.

DISCUSSION

Various culture techniques utilizing scaffold-based and scaffold-free methods have been used to generate 3-D cultures (55). Stem cells, either human embryonic stem cells (hPSCs) or induced pluripotent stem cells (iPSC), can be promoted to develop into organoids when placed in a suitable culture environment containing extracellular matrix (ECM) scaffolds (9). Tractional forces created by cell/ECM interactions facilitate cell self-assembly into organotypic cell clusters that support growth, proliferation and differentiation (56–58). Once fully matured, organoids recapitulate certain structural characteristics of native tissue (59, 60). The ability to generate complex tissue structures or populations of specialized cell types in vitro has motivated increasing use of organoids for basic and biomedical research, drug discovery, and tissue engineering (55, 61).

The mucociliary surface epithelium of the human nose, trachea and bronchi is largely recapitulated in planar ALI cultures, making these the most frequently used cell models for studying the proximal airway epithelium. Airway organoids have also shown great promise as models for studying human airway development, cell biology, and disease (8, 62, 63). Additionally, airway epithelial organoids offer potential advantages in that they can be generated from minimal starting material making them useful for diagnosis of diseases such as ciliopathies and precision medicine such as testing individual responses to drugs. Airway organoid cultures can also provide large numbers of disease-relevant, testable units for drug discovery. Airway cell models with externally orientated polarized epithelium provide some useful advantages. Ready access to the apical membrane permits testing uptake and effectiveness of drugs designed for inhalational delivery. Delivery of RNA- and DNA-based therapies directed at the surface epithelial cells can be tested in experiments that better mimic aerosolized delivery. Of particular current importance, airway organoids represent a useful tool to study infectivity of aerosolized microorganisms including respiratory viruses such as coronavirus, influenza virus and others, especially in light of findings suggesting an important role for ciliated cells in viral infection and spreading (39, 64–66).

In this report, we describe a method for producing airway epithelial organoids either directly from isolated tracheobronchial cells without pre-culture manipulations, induction of pluripotency, isolation of stem cells, or addition of mesenchymal stromal cells. We also show that organoids could be produced from small numbers of tracheobronchial epithelial cells that underwent up to four rounds of conditional reprogramming and serial expansion. A comparison of the methods and findings herein with those described in previous reports of airway organoids originating from airway tissues and cells is summarized in Table 4. Remarkably, each of these previous studies describe organoid type cultures that develop inwardly orientated apical membranes lining their cystic cavities. In contrast to previous approaches, our organoid cultures primarily display externally orientated polarity. Otherwise, our technique produces airway organoids that share features with previously described organoid cell models, including the expression of airway epithelial markers of differentiation.

Table 4.

Comparison of airway organoid-type cultures derived from human nasal, tracheal, or bronchial cells or tissues from studies that include detailed characterization of the organoids¹

References	Starting Material	Cells Used to Create Organoids	Organoid Culture Basal Media2	Undefined Media Com-ponents	Organoid Culture Additives2	Matrix	Seeding Density	Mean Organoid Yield	Mean Organoid Diameter(µm)	Organoid Surface Area(µm2)	Polarity
Boecking et al. (this publication)	Isolated HTBE and CFBE cells (wt and CF)	1) Isolated HTBE and CFBE cells (P0) 2) HTBE CRCs (P2–P4)	UNC ALI medium (18)3	BPE	R-spondin 2 and Noggin beginning at day 14	1:1 mixture of PureCol and GFR Matrigel4	2.5 × 105 cells/cm2	1,720 organoids/cm2	HTBE: 129 CFBE: 163 (corrected for processing shrinkage)	HTBE: 52,279 CFBE: 83,467 (calculated)	Primarily external but some internal
Danahay et al. (32) [also see Hild and Jaffe (69)]	HTBE cells, P1	HTBE cells, P2	BEGM growth medium5	BPE	5% (v/v) GFR Matrigel in culture media	25% solution of GFR Matrigel in culture media	1.714 × 104 cells/cm2	Not provided Note: at ≤75 cells/0.056/cm2 40% cloning efficiency	Not provided	Not provided	Internal
Gao et al. (33)	Isolated HBE cells	HBE cells, P1	UNC ALI medium	BPE; MRC5 human lung fibroblasts (7.2 × 104 cells/cm2)	50% UNC ALI; 50% GFR Matrigel	100% GFR Matrigel coating	3.6 × 103 cells/cm2	CFE ∼11%	∼183	Not provided	Internal
Butler et al. (35) [also see Hynds et al. (72)]	Explant cultures of airway brushings or biopsies expanded as CRCs or in BEGM	1) HBE CRCs at P1 and P4 2) HBE cells propagated in BEGM at P1 and P4	Tracheosphere Medium (1:1 mixture of DMEM and BEBM with BEGM supplements5 except all-trans RA added separately at time of use; triiodothyronine and amphotericin B excluded)	BPE	5% GFR Matrigel Y-27632 (5 µM) added at seeding but not feeding	25% GFR Matrigel precoat of plates	7.812 × 103/cm2	Not provided	P1: CRCs: ∼90–100 BEGM: ∼85–90 P4: CRC: ∼105–115 BEGM: ∼45–90 Note: obtained by measuring 30 largest tracheospheres/well	Not provided	Internal
Zhou et al. (39)	Lung tissue [method of Sachs, et al. (59)]	Airway stem cells	AO medium: Advanced DMEM/F12 PD medium:PneumaCult-ALI6	PD medium: PneumaCultALI4 (Proprietary formulation)	AO medium: R-spondin 1 Noggin B27 Y-27632 A83-01 SB202190 FGF 7FGF10 Heregulin β-1 PD medium: DAPT (10 µM)	500 µL 100% GFR Matrigel coating/6-well plate	See method of Sachs et al. (59) below	Not provided	∼190 (AO medium) ∼∼110 (PD medium)	Not provided	AO medium: organoids show sparse cilia, mostly internalPD medium: Internal
Tanaka et al. (34)	Isolated HBE cells cultured for 13 days on mitomycin-C-treated NIH 3T3 feeder cells in BEGM	HBE P1 cells	CnT-PR-AD medium7 with 1 mM CaCl2	FGF (100 ng/mL) DAPT 20 µM	1% GFR Matrigel FGF (100 ng/mL) DAPT 20 µM	100% GFR Matrigel coating	1.05 × 105 cells/cm2	Not provided	Not provided	Not provided	Internal
Brewington et al. (37)	Human nasal epithelial (HNE) cells and F508del/F508del CFNE cells obtained by curettage	CRC P1 and P2	Vertex differentiation media (DF12 with supplements8)	2% USG 2% Fetal Clone II0. 25% bovine brain extract		100% Matrigel	5 × 105 cells/mL of 100% Matrigel (type not specified) seeded as Matrigel drop at 2.5 × 103 cells/cm2)	50,000 cells yielded 50–100 organoids	50 to 400+	Not provided	Internal
Dobzanski et al. (36)	HSNE cells obtained by brushing	HSNE cells expanded in Pneumacult-Ex Plus6	1:1 PneumaCult-ALI and GFR Matrigel	PneumaCult ALI Basal transwell compartment with: a) media only, b) media and control fibroblasts c) media and nasal polyp fibroblasts		100% GFR Matrigel	3 × 103 cells/cm2	CFE ∼6% (control) ∼12% (control fibroblast coculture) ∼21% (nasal polyp fibroblast coculture	Calculated from Figure 4: a) control ≤300 (∼69% 50–150; ∼31% 50–150) b) fibroblast co-culture ≤500 (∼29% 50–150; ∼67%; 150–300; ∼4% 300–500) c) nasal polyp fibroblast coculture (∼12.5% 50–150; ∼52%; 150–300; ∼33% 300–500; ∼3% > 500)	Not provided	Not described but phase micrographs do not reveal external cilia
Sachs et al. (40); see also Hui et al. (70)	Lung tissue; Broncho-alveolar lavage fluids (wt and CF)	Airway stem cells	Advanced DMEM/F-12		R-spondin 1 FGF 7 FGF10 Noggin A83-01 Y-27632 SB202190 B27 supplement	Cultrex BME4	Initial seeding density not provided. Organoids collected, mechanically sheared and fragments passaged every 2 weeks at 1:1-1:6 reseeding ratios	Not provided	Not provided	Not provided	Internal
Liu et al. (38)	HNE and CFNE (various genotypes) obtained by brush biopsies	CRC ≤ P3	Vertex Differentiation Media	2% USG 2% Fetal Clone II0. 25% Bovine brain extract	20% Matrigel	100% Matrigel	Angiogenesis slides: 4 × 103/cm2 Transwells: ∼9.1 × 104 cells/cm2	31 wt (248/cm2) 37 CF (296/cm2) on angiogenesis slides	Not provided	wt: ∼52,000 CF: ∼28,000 to ∼30,000	Internal

¹Abbreviations: ALI, air-liquid interface; BEBM, bronchial epithelial basal media; BEGM, basal epithelial growth medium; BME, basement membrane extract; BPE, bovine pituitary extract; CF, cystic fibrosis; CFBE, cystic fibrosis bronchial epithelial; CFE, colony forming efficiency; CFNE, cystic fibrosis nasal epithelial; CRCs, conditional reprogrammed cells; DAPT, (N-[N-(3,5-difluorophenacetyl)-l-alanyl]-S-phenylglycine t-butyl ester (a γ-secretase inhibitor blocker of Notch pathway); DF12, (1:1 mixture of DMEM and Ham’s F12 nutrient mixture); DMEM, Dulbecco’s modified Eagle’s medium; FGF, fibroblast growth factor; GFR, growth factor reduced; F12, Ham’s F12 nutrient mixture; HBE, human bronchial epithelium; HTBE, human tracheobronchial epithelial; HSNE, human sinonasal epithelial; P0, Passage number; RA, retinoic acid; USG, Ultroser G; wt, wild type.

²See specific references for complete listing of media components, their respective concentrations, and contact information for commercial suppliers.

³See also Supplemental Table 1, herein.

⁴Matrigel (Corning Life Sciences) and Cultrex BME (basement membrane extract; R&D Systems, Inc.) are basement membrane substrates produced by Engelbreth-Holm-Swarm mouse sarcoma cells.

⁵BEGM growth medium: BEBM supplemented with single Quots supplements and growth factors (Lonza) containing human recombinant epidermal growth factor (0.5 ng/mL), insulin (5 μg/mL), transferrin (10 μg/mL), hydrocortisone (0.5 μg/mL), triiodothyronine (6.5 μg/mL), epinephrine (0.5 μg/mL), retinoic acid (50 nM), gentamycin and amphotericin-B (50 μg/mL), and bovine pituitary extract (35 mg/mL).

⁶STEMCELL Technologies.

⁷Cellntec.

⁸Vertex Differentiation Media: 2% Ultroser-G (Pal Corporation), 2% Fetal Clone II (VWR), insulin (2.5 μg/mL), 0.25% bovine brain extract (MilliporeSigma), hydrocortisone (20 nM), triiodothyronine (500 nM), transferrin (2.5 μg/mL), ethanolamine (250 nM), epinephrine (1.5 μM), phosphoethanolamine (250 nM), and RA (10 nM). See also Van Goor et al. (73) and Galietta et al. (74).

We can opine on potential reason(s) that the method reported here produces organoids with external polarity whereas prior methods do not. Our protocol employs a mixture of Pure Col and Matrigel, in contrast to the use of Matrigel alone in other reports. We found that the addition of PureCol, a collagen mixture composed primarily of Type I collagen, greatly increased the number of organoid units. However, using Matrigel alone while keeping other conditions constant also resulted in externally orientated organoids though fewer in number and of inferior quality. Methods for producing airway organoids starting with ESCs or iPSCs typically use mediators of Wnt and bone morphogenic protein (BMP) signaling throughout the organoid culture period. Uniformly, these studies report internally orientated organoids (26–28, 67). Similarly, other studies in which organoids are produced directly from human airway cells yielded only internally orientated organoids do not generally use mediators of Wnt or BMP signaling pathways. Sachs et al. (40) reported generation of organoids from lung tissue or bronchoalveolar lavage fluids in which R-spondin 1 and Noggin were added from the onset of culture. Though they noted that R-spondin 1 is critical for long-term, serial propagation of cultures, their organoids also develop internally orientated polarity. In a related study using the same media components, Zhou et al. (39) found rare organoids with sparse externally orientated cilia among mostly internally orientated organoids. Our method includes an initial 14-day period in which organoids begin development, followed by culture in the presence of RSPO2 and Noggin. During the initial 14-day phase of differentiation, our organoids behave similarly to airway spheroids which are also externally oriented. Our choice of RSPO2 rather than RSPO1 reflects its importance in laryngotracheal development (68). This and the 14-day lag in adding RSPO2 and Noggin offer possible reasons for the polarity of our organoids, though additional mechanistic studies are needed to resolve this question.

Airway epithelial organoids (nasospheres, bronchospheres, tracheospheres) develop by first forming clusters of cells that proliferate eventually forming single central cavities that expand or several cavities that typically coalesce to form a single cavity (32, 69). While other studies have not quantified mature organoids with multiple cavities, images of such are seen in a number of studies (34, 38, 40, 70). The final yield of organoids expressed as a ratio of organoids per number of initial cells seeded using our protocol (0.006) is greater than that reported by Brewington et al. (0.001–0.002) (37) but 10-fold less than that reported by Liu et al. (0.062) (38). In our protocol, a substantial loss of organoid units occurs with the complete media change at day 14. Modifying our protocol by adding RSPO2 and Noggin directly to the cultures at day 14 rather than harvesting and replating would likely increase the final organoid yield; however, its effect on polarity needs to be determined. The average size of organoids varies slightly among reported studies. Adjusted for tissue processing shrinkage (71), the calculated average diameter of our P0 wt HTBE organoids (129 µm) is slightly larger than reported by Butler et al. for passage 1 CRC bronchial organoids (90–100 µm) (35), but generally slightly smaller than that those produced by others (Table 4) (72–74). For organoids generated from nasal epithelial CRCs, Liu et al. (38) measured the surface area of wt organoids as ∼52,000 µm² while organoids from cells with minimal function CFTR genotypes were calculated to be notably smaller (∼29,000 µm²). The calculated surface area of our wt organoids at 52,279 µm² is nearly identical while our F508del/F508del CF organoids are larger at 83,467 µm², which is likely due to their predominately externally orientated polarization (Table 4). Except herein, the percentage of airway organoids lacking central cavities or containing multiple cavities has not been quantified, though published images of mature organoids depict some with more than one cavity (34, 35, 38, 40, 70).

Our study also compared airway organoid cultures with planar ALI cultures, and in some studies, with the native epithelium. Mean cilia length of organoid and ALI cultures were at the low end of the normally expected length of 6.5 to 7 µm (75). However, this was not the case following conditional reprogramming, as increasing passage resulted in progressive decreases in cilia length with organoids showing greater shortening than ALI cultures, an effect of conditional reprogramming that to our knowledge has not been previously reported. Organoid cilia beat frequency was greater than that in ALI cultures, and though CBF increased progressively with successive CRC passages for each airway cell model, organoid CBF was higher at comparable cell passages suggesting that the increased CBF was related to the decrease in ciliary length. In contrast to our data, Konishi et al. (27) compared CBF in organoids and ALI cultures derived from hPSC and serially passaged HBE cells, reporting that the cilia of ALI cultures beat slightly faster than organoid cultures. The authors suggested that two factors may have contributed to the differences, including trapped, viscous mucoid secretions within the organoid lumen and use of minced organoids for measurement of CBF. Liu et al. (38) studied nasal CRC organoids prepared from 10 individuals, reporting CBF ranged from 11 to 17 Hz. Although passage number was not reported, none of their organoids were greater than passage 3. Finally, we note that normal values obtained for CBF of human respiratory MCC vary widely from 7.0 to 20 Hz, which may be due to lack of a standard measurement technique and to true variations based on the anatomical location of the ciliated cells (76–79). Such factors, as well variable culture conditions and cellular changes following conditional reprogramming, are relevant considerations when analyzing CBF in both planar and organoid airway cell cultures (80).

Although our method does produce differentiated organoids, several limitations to the current study are noted. Some of the externally orientated organoids contain more than one internal cavity though the majority of these internal cavities lack cilia or obvious apical differentiation. A small subset of organoids shows only inward polarity. Variability in polarity could hinder some assays for functional testing of CFTR and other epithelial ion channels without additional analysis of orientation at the individual organoid level. As our organoids lack differentiated goblet cells, studies of mucin production and secretion are currently limited. However, since components of the culture medium have significant effects on basal cell differentiation, conditions that promote goblet cell differentiation, such as activators of Notch signaling, may skew differentiation towards secretory cell types (32, 81).

Conclusions

As a model of the human airway epithelium, organoids complement planar ALI cultures and may offer several potential advantages as described in the introduction. Herein, we describe an uncomplicated protocol for culturing differentiated airway organoids with externally orientated apical membranes from small numbers of bronchial epithelial cells, either freshly isolated or expanded in numbers via conditional reprogramming. This method circumvents the generation of human ESCs or iPSCs, isolation of stem cells, coculture with mesenchymal stem cells, or other preculture manipulations that are often used for tissue-specific organoid research. The culture media is well-defined and does not rely on proprietary formulations, containing only one undefined supplement (bovine pituitary extract) that may have batch-to-batch variability. Future modifications of soluble and insoluble culture conditions can be readily tested for their effect on organoid growth, differentiation and function, as well as cilia formation and function. Since only a small amount of starting cellular material is required, the method is applicable to producing organoids from nasal or tracheobronchial brushings to supplement diagnostic studies or individualized theratyping. In addition, its potential for producing a large number of testable organoid units make this model potentially suitable for high-throughput screening.

DATA AVAILABILITY

Data will be made available upon reasonable request.

SUPPLEMENTAL DATA

Supplemental Table S1: https://doi.org/10.6084/m9.figshare.14489352.v2.

Supplemental Figure Legends: https://doi.org/10.6084/m9.figshare.14489352.

Supplemental Fig. S1: https://doi.org/10.6084/m9.figshare.14489169.

Supplemental Fig. S2: https://doi.org/10.6084/m9.figshare.15262320.

Supplemental Fig. S3: https://doi.org/10.6084/m9.figshare.15262419.

Supplemental Fig. S4: https://doi.org/10.6084/m9.figshare.15262521.

Supplemental Video S1: https://doi.org/10.6084/m9.figshare.14489208.

Supplemental Video S2: https://doi.org/10.6084/m9.figshare.14489211.

Supplemental Video S3: https://doi.org/10.6084/m9.figshare.14489214.

Supplemental Video S4: https://doi.org/10.6084/m9.figshare.14489220.

Supplemental Video S5: https://doi.org/10.6084/m9.figshare.14489226.

GRANTS

This work was supported by NIH Grant DK72517, a Research Development grant from the Cystic Fibrosis Foundation (A.S.V., P.M.H., W.E.F), and Cystic Fibrosis Foundation Collaborative Research Grant URNOV19XX0 (W.E.F). C.A.B. was supported in part by an Elizabeth Nash Fellowship from Cystic Fibrosis Research, Inc. P.W. was funded by a Pathway to Independence Award by the National Heart, Lung, and Blood Institute (K99HL127275), and work in the Walentek Lab was supported by the German Research Foundation (DFG) under the Emmy Noether Program (Grant WA3365/2-1) and under Germany’s Excellence Strategy (CIBSS–EXC-2189–Project ID 390939984). Work in the Marshall laboratory was supported by NIH Grant R35 GM130327. P.J.W. was supported in part by the Nina Ireland Program for Lung Health.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

C.A.B., A.S.V., W.E.F., and P.W. conceived and designed research; C.A.B., P.W., L.T.Z., D.I.S., H.I., and B.-J.J. performed experiments; C.A.B., A.S.V., W.E.F., P.W., H.I., P.M.H., and W.F.M. analyzed data; C.A.B., A.S.V., W.E.F., P.W., and B.-J.J. interpreted results of experiments; C.A.B., A.S.V., W.E.F., P.W., and B.-J.J. prepared figures; C.A.B., A.S.V., W.E.F., and P.W. drafted manuscript; C.A.B., A.S.V., W.E.F., P.W., L.T.Z., D.I.S., P.J.W., H.I., P.M.H., and W.F.M. edited and revised manuscript; C.A.B., A.S.V., W.E.F., and P.W. approved final version of manuscript.