1
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Jaber N, Billet S. How to use an in vitro approach to characterize the toxicity of airborne compounds. Toxicol In Vitro 2024; 94:105718. [PMID: 37871865 DOI: 10.1016/j.tiv.2023.105718] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/22/2023] [Revised: 09/13/2023] [Accepted: 09/21/2023] [Indexed: 10/25/2023]
Abstract
As part of the development of new approach methodologies (NAMs), numerous in vitro methods are being developed to characterize the potential toxicity of inhalable xenobiotics (gases, volatile organic compounds, polycyclic aromatic hydrocarbons, particulate matter, nanoparticles). However, the materials and methods employed are extremely diverse, and no single method is currently in use. Method standardization and validation would raise trust in the results and enable them to be compared. This four-part review lists and compares biological models and exposure methodologies before describing measurable biomarkers of exposure or effect. The first section emphasizes the importance of developing alternative methods to reduce, if not replace, animal testing (3R principle). The biological models presented are mostly to cultures of epithelial cells from the respiratory system, as the lungs are the first organ to come into contact with air pollutants. Monocultures or cocultures of primary cells or cell lines, as well as 3D organotypic cultures such as organoids, spheroids and reconstituted tissues, but also the organ(s) model on a chip are examples. The exposure methods for these biological models applicable to airborne compounds are submerged, intermittent, continuous either static or dynamic. Finally, within the restrictions of these models (i.e. relative tiny quantities, adhering cells), the mechanisms of toxicity and the phenotypic markers most commonly examined in models exposed at the air-liquid interface (ALI) are outlined.
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Affiliation(s)
- Nour Jaber
- UR4492, Unité de Chimie Environnementale et Interactions sur le Vivant, Université du Littoral Côte d'Opale, Dunkerque, France
| | - Sylvain Billet
- UR4492, Unité de Chimie Environnementale et Interactions sur le Vivant, Université du Littoral Côte d'Opale, Dunkerque, France.
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2
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Keiser NW, Cant E, Sitaraman S, Shoemark A, Limberis MP. Restoring Ciliary Function: Gene Therapeutics for Primary Ciliary Dyskinesia. Hum Gene Ther 2023; 34:821-835. [PMID: 37624733 DOI: 10.1089/hum.2023.102] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 08/27/2023] Open
Abstract
Primary ciliary dyskinesia (PCD) is a genetic disease characterized by defects in motile cilia, which play an important role in several organ systems. Lung disease is a hallmark of PCD, given the essential role of cilia in airway surface defense. Diagnosis of PCD is complicated due to its reliance on complex tests that are not utilized by every clinic and also its phenotypic overlap with several other respiratory diseases. Nonetheless, PCD is increasingly being recognized as more common than once thought. The disease is genetically complex, with several genes reported to be associated with PCD. There is no cure for PCD, but gene therapy remains a promising therapeutic strategy. In this review, we provide an overview of the clinical symptoms, diagnosis, genetics, and current treatment regimens for PCD. We also describe PCD model systems and discuss the therapeutic potential of different gene therapeutics for targeting the intended cellular target, the ciliated cells of the airway.
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Affiliation(s)
| | - Erin Cant
- Division of Molecular and Clinical Medicine, University of Dundee, Ninewells Hospital and Medical School, Dundee, United Kingdom
| | | | - Amelia Shoemark
- Division of Molecular and Clinical Medicine, University of Dundee, Ninewells Hospital and Medical School, Dundee, United Kingdom
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3
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Humbert MV, Spalluto CM, Bell J, Blume C, Conforti F, Davies ER, Dean LSN, Elkington P, Haitchi HM, Jackson C, Jones MG, Loxham M, Lucas JS, Morgan H, Polak M, Staples KJ, Swindle EJ, Tezera L, Watson A, Wilkinson TMA. Towards an artificial human lung: modelling organ-like complexity to aid mechanistic understanding. Eur Respir J 2022; 60:2200455. [PMID: 35777774 DOI: 10.1183/13993003.00455-2022] [Citation(s) in RCA: 4] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/01/2022] [Accepted: 06/11/2022] [Indexed: 11/05/2022]
Abstract
Respiratory diseases account for over 5 million deaths yearly and are a huge burden to healthcare systems worldwide. Murine models have been of paramount importance to decode human lung biology in vivo, but their genetic, anatomical, physiological and immunological differences with humans significantly hamper successful translation of research into clinical practice. Thus, to clearly understand human lung physiology, development, homeostasis and mechanistic dysregulation that may lead to disease, it is essential to develop models that accurately recreate the extraordinary complexity of the human pulmonary architecture and biology. Recent advances in micro-engineering technology and tissue engineering have allowed the development of more sophisticated models intending to bridge the gap between the native lung and its replicates in vitro Alongside advanced culture techniques, remarkable technological growth in downstream analyses has significantly increased the predictive power of human biology-based in vitro models by allowing capture and quantification of complex signals. Refined integrated multi-omics readouts could lead to an acceleration of the translational pipeline from in vitro experimental settings to drug development and clinical testing in the future. This review highlights the range and complexity of state-of-the-art lung models for different areas of the respiratory system, from nasal to large airways, small airways and alveoli, with consideration of various aspects of disease states and their potential applications, including pre-clinical drug testing. We explore how development of optimised physiologically relevant in vitro human lung models could accelerate the identification of novel therapeutics with increased potential to translate successfully from the bench to the patient's bedside.
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Affiliation(s)
- Maria Victoria Humbert
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
- Department of Medicine, University of Cambridge, Cambridge, UK
| | - Cosma Mirella Spalluto
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
- M.V. Humbert and C.M. Spalluto are co-first authors and contributed equally to this work
| | - Joseph Bell
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
| | - Cornelia Blume
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
- Institute for Life Sciences, University of Southampton, Southampton, UK
- School of Human Development and Health, Faculty of Medicine, University of Southampton, Southampton, UK
| | - Franco Conforti
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
| | - Elizabeth R Davies
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- Biological Sciences, Faculty of Environmental and Life Sciences, University of Southampton, Southampton, UK
| | - Lareb S N Dean
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
| | - Paul Elkington
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
- Institute for Life Sciences, University of Southampton, Southampton, UK
| | - Hans Michael Haitchi
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
- Institute for Life Sciences, University of Southampton, Southampton, UK
| | - Claire Jackson
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
| | - Mark G Jones
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
| | - Matthew Loxham
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
- Institute for Life Sciences, University of Southampton, Southampton, UK
| | - Jane S Lucas
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
| | - Hywel Morgan
- Institute for Life Sciences, University of Southampton, Southampton, UK
- Electronics and Computer Science, Faculty of Physical Sciences and Engineering, University of Southampton, Southampton, UK
| | - Marta Polak
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
- Institute for Life Sciences, University of Southampton, Southampton, UK
| | - Karl J Staples
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
| | - Emily J Swindle
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
- Institute for Life Sciences, University of Southampton, Southampton, UK
| | - Liku Tezera
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- Department of Infection and Immunity, Faculty of Medicine, University College London, London, UK
| | - Alastair Watson
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
- College of Medical and Dental Sciences, University of Birmingham, Birmingham, UK
- School of Clinical Medicine, University of Cambridge, Cambridge, UK
- Department of Medicine, University of Cambridge, Cambridge, UK
| | - Tom M A Wilkinson
- School of Clinical and Experimental Sciences, Faculty of Medicine, University of Southampton, Southampton, UK
- NIHR Southampton Biomedical Research Centre, University Hospital Southampton, Southampton, UK
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4
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Jackson CL, Bottier M. Methods for the assessment of human airway ciliary function. Eur Respir J 2022; 60:13993003.02300-2021. [PMID: 35595315 DOI: 10.1183/13993003.02300-2021] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/20/2021] [Accepted: 04/19/2022] [Indexed: 11/05/2022]
Affiliation(s)
- Claire L Jackson
- Primary Ciliary Dyskinesia Centre, NIHR Biomedical Research Centre, University Hospital Southampton NHS Foundation Trust, Southampton, UK .,School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton, UK
| | - Mathieu Bottier
- Division of Molecular and Clinical Medicine, Ninewells Hospital and Medical School, University of Dundee, Dundee, UK
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5
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Boecking CA, Walentek P, Zlock LT, Sun DI, Wolters PJ, Ishikawa H, Jin BJ, Haggie PM, Marshall WF, Verkman AS, Finkbeiner WE. A simple method to generate human airway epithelial organoids with externally orientated apical membranes. Am J Physiol Lung Cell Mol Physiol 2022; 322:L420-L437. [PMID: 35080188 PMCID: PMC8917940 DOI: 10.1152/ajplung.00536.2020] [Citation(s) in RCA: 10] [Impact Index Per Article: 5.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/08/2023] Open
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.
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Affiliation(s)
- Carolin A. Boecking
- 1Department of Pathology, University of California, San Francisco, California
| | - Peter Walentek
- 2Genomics and Development Division, Molecular and Cell Biology Department, University of California, Berkeley, California,3Renal Division, Department of Medicine, University Hospital Freiburg, Freiburg University Faculty of Medicine, Freiburg, Germany,4CIBSS – Centre for Integrative Biological Signalling Studies, Albert Ludwigs University Freiburg, Freiburg, Germany
| | - Lorna T. Zlock
- 1Department of Pathology, University of California, San Francisco, California
| | - Dingyuan I. Sun
- 1Department of Pathology, University of California, San Francisco, California
| | - Paul J. Wolters
- 5Department of Medicine, University of California, San Francisco, California
| | - Hiroaki Ishikawa
- 6Department of Biochemistry and Biophysics, University of California, San Francisco, California
| | - Byung-Ju Jin
- 5Department of Medicine, University of California, San Francisco, California
| | - Peter M. Haggie
- 5Department of Medicine, University of California, San Francisco, California
| | - Wallace F. Marshall
- 6Department of Biochemistry and Biophysics, University of California, San Francisco, California
| | - Alan S. Verkman
- 5Department of Medicine, University of California, San Francisco, California,7Department of Physiology, University of California, San Francisco, California
| | - Walter E. Finkbeiner
- 1Department of Pathology, University of California, San Francisco, California,8Innovative Genomics Institute, University of California, Berkeley, California
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6
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Niziolek M, Bicka M, Osinka A, Samsel Z, Sekretarska J, Poprzeczko M, Bazan R, Fabczak H, Joachimiak E, Wloga D. PCD Genes-From Patients to Model Organisms and Back to Humans. Int J Mol Sci 2022; 23:ijms23031749. [PMID: 35163666 PMCID: PMC8836003 DOI: 10.3390/ijms23031749] [Citation(s) in RCA: 7] [Impact Index Per Article: 3.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/30/2021] [Revised: 01/25/2022] [Accepted: 01/31/2022] [Indexed: 01/27/2023] Open
Abstract
Primary ciliary dyskinesia (PCD) is a hereditary genetic disorder caused by the lack of motile cilia or the assembxly of dysfunctional ones. This rare human disease affects 1 out of 10,000-20,000 individuals and is caused by mutations in at least 50 genes. The past twenty years brought significant progress in the identification of PCD-causative genes and in our understanding of the connections between causative mutations and ciliary defects observed in affected individuals. These scientific advances have been achieved, among others, due to the extensive motile cilia-related research conducted using several model organisms, ranging from protists to mammals. These are unicellular organisms such as the green alga Chlamydomonas, the parasitic protist Trypanosoma, and free-living ciliates, Tetrahymena and Paramecium, the invertebrate Schmidtea, and vertebrates such as zebrafish, Xenopus, and mouse. Establishing such evolutionarily distant experimental models with different levels of cell or body complexity was possible because both basic motile cilia ultrastructure and protein composition are highly conserved throughout evolution. Here, we characterize model organisms commonly used to study PCD-related genes, highlight their pros and cons, and summarize experimental data collected using these models.
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Affiliation(s)
- Michal Niziolek
- Laboratory of Cytoskeleton and Cilia Biology, Nencki Institute of Experimental Biology, Polish Academy of Sciences, 3 Pasteur Street, 02-093 Warsaw, Poland; (M.N.); (M.B.); (A.O.); (Z.S.); (J.S.); (M.P.); (R.B.); (H.F.)
| | - Marta Bicka
- Laboratory of Cytoskeleton and Cilia Biology, Nencki Institute of Experimental Biology, Polish Academy of Sciences, 3 Pasteur Street, 02-093 Warsaw, Poland; (M.N.); (M.B.); (A.O.); (Z.S.); (J.S.); (M.P.); (R.B.); (H.F.)
- Faculty of Chemistry, University of Warsaw, 1 Pasteur Street, 02-093 Warsaw, Poland
| | - Anna Osinka
- Laboratory of Cytoskeleton and Cilia Biology, Nencki Institute of Experimental Biology, Polish Academy of Sciences, 3 Pasteur Street, 02-093 Warsaw, Poland; (M.N.); (M.B.); (A.O.); (Z.S.); (J.S.); (M.P.); (R.B.); (H.F.)
| | - Zuzanna Samsel
- Laboratory of Cytoskeleton and Cilia Biology, Nencki Institute of Experimental Biology, Polish Academy of Sciences, 3 Pasteur Street, 02-093 Warsaw, Poland; (M.N.); (M.B.); (A.O.); (Z.S.); (J.S.); (M.P.); (R.B.); (H.F.)
| | - Justyna Sekretarska
- Laboratory of Cytoskeleton and Cilia Biology, Nencki Institute of Experimental Biology, Polish Academy of Sciences, 3 Pasteur Street, 02-093 Warsaw, Poland; (M.N.); (M.B.); (A.O.); (Z.S.); (J.S.); (M.P.); (R.B.); (H.F.)
| | - Martyna Poprzeczko
- Laboratory of Cytoskeleton and Cilia Biology, Nencki Institute of Experimental Biology, Polish Academy of Sciences, 3 Pasteur Street, 02-093 Warsaw, Poland; (M.N.); (M.B.); (A.O.); (Z.S.); (J.S.); (M.P.); (R.B.); (H.F.)
- Laboratory of Immunology, Mossakowski Medical Research Institute, Polish Academy of Sciences, 5 Pawinskiego Street, 02-106 Warsaw, Poland
| | - Rafal Bazan
- Laboratory of Cytoskeleton and Cilia Biology, Nencki Institute of Experimental Biology, Polish Academy of Sciences, 3 Pasteur Street, 02-093 Warsaw, Poland; (M.N.); (M.B.); (A.O.); (Z.S.); (J.S.); (M.P.); (R.B.); (H.F.)
| | - Hanna Fabczak
- Laboratory of Cytoskeleton and Cilia Biology, Nencki Institute of Experimental Biology, Polish Academy of Sciences, 3 Pasteur Street, 02-093 Warsaw, Poland; (M.N.); (M.B.); (A.O.); (Z.S.); (J.S.); (M.P.); (R.B.); (H.F.)
| | - Ewa Joachimiak
- Laboratory of Cytoskeleton and Cilia Biology, Nencki Institute of Experimental Biology, Polish Academy of Sciences, 3 Pasteur Street, 02-093 Warsaw, Poland; (M.N.); (M.B.); (A.O.); (Z.S.); (J.S.); (M.P.); (R.B.); (H.F.)
- Correspondence: (E.J.); (D.W.); Tel.: +48-22-58-92-338 (E.J. & D.W.)
| | - Dorota Wloga
- Laboratory of Cytoskeleton and Cilia Biology, Nencki Institute of Experimental Biology, Polish Academy of Sciences, 3 Pasteur Street, 02-093 Warsaw, Poland; (M.N.); (M.B.); (A.O.); (Z.S.); (J.S.); (M.P.); (R.B.); (H.F.)
- Correspondence: (E.J.); (D.W.); Tel.: +48-22-58-92-338 (E.J. & D.W.)
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7
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Three-Dimensional Airway Spheroids and Organoids for Cystic Fibrosis Research. JOURNAL OF RESPIRATION 2021. [DOI: 10.3390/jor1040022] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/13/2022] Open
Abstract
Cystic fibrosis (CF) is an autosomal recessive multi-organ disease caused by mutations in the CF Transmembrane Conductance Regulator (CFTR) gene, with morbidity and mortality primacy related to the lung disease. The CFTR protein, a chloride/bicarbonate channel, is expressed at the apical side of airway epithelial cells and is mainly involved in appropriate ion and fluid transport across the epithelium. Although many animal and cellular models have been developed to study the pathophysiological consequences of the lack/dysfunction of CFTR, only the three-dimensional (3D) structures termed “spheroids” and “organoids” can enable the reconstruction of airway mucosa to model organ development, disease pathophysiology, and drug screening. Airway spheroids and organoids can be derived from different sources, including adult lungs and induced pluripotent stem cells (iPSCs), each with its advantages and limits. Here, we review the major features of airway spheroids and organoids, anticipating that their potential in the CF field has not been fully shown. Further work is mandatory to understand whether they can accomplish better outcomes than other culture conditions of airway epithelial cells for CF personalized therapies and tissue engineering aims.
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8
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Nasal Epithelial Cell-Based Models for Individualized Study in Cystic Fibrosis. Int J Mol Sci 2021; 22:ijms22094448. [PMID: 33923202 PMCID: PMC8123210 DOI: 10.3390/ijms22094448] [Citation(s) in RCA: 13] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Key Words] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/09/2021] [Revised: 04/22/2021] [Accepted: 04/23/2021] [Indexed: 12/11/2022] Open
Abstract
The emergence of highly effective CFTR modulator therapy has led to significant improvements in health care for most patients with cystic fibrosis (CF). For some, however, these therapies remain inaccessible due to the rarity of their individual CFTR variants, or due to a lack of biologic activity of the available therapies for certain variants. One proposed method of addressing this gap is the use of primary human cell-based models, which allow preclinical therapeutic testing and physiologic assessment of relevant tissue at the individual level. Nasal cells represent one such tissue source and have emerged as a powerful model for individual disease study. The ex vivo culture of nasal cells has evolved over time, and modern nasal cell models are beginning to be utilized to predict patient outcomes. This review will discuss both historical and current state-of-the art use of nasal cells for study in CF, with a particular focus on the use of such models to inform personalized patient care.
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9
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Coles JL, Thompson J, Horton KL, Hirst RA, Griffin P, Williams GM, Goggin P, Doherty R, Lackie PM, Harris A, Walker WT, O’Callaghan C, Hogg C, Lucas JS, Blume C, Jackson CL. A Revised Protocol for Culture of Airway Epithelial Cells as a Diagnostic Tool for Primary Ciliary Dyskinesia. J Clin Med 2020; 9:E3753. [PMID: 33233428 PMCID: PMC7700393 DOI: 10.3390/jcm9113753] [Citation(s) in RCA: 21] [Impact Index Per Article: 5.3] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/19/2020] [Revised: 11/17/2020] [Accepted: 11/19/2020] [Indexed: 12/14/2022] Open
Abstract
Air-liquid interface (ALI) culture of nasal epithelial cells is a valuable tool in the diagnosis and research of primary ciliary dyskinesia (PCD). Ex vivo samples often display secondary dyskinesia from cell damage during sampling, infection or inflammation confounding PCD diagnostic results. ALI culture enables regeneration of healthy cilia facilitating differentiation of primary from secondary ciliary dyskinesia. We describe a revised ALI culture method adopted from April 2018 across three collaborating PCD diagnostic sites, including current University Hospital Southampton COVID-19 risk mitigation measures, and present results. Two hundred and forty nasal epithelial cell samples were seeded for ALI culture and 199 (82.9%) were ciliated. Fifty-four of 83 (63.9%) ex vivo samples which were originally equivocal or insufficient provided diagnostic information following in vitro culture. Surplus basal epithelial cells from 181 nasal brushing samples were frozen in liquid nitrogen; 39 samples were ALI-cultured after cryostorage and all ciliated. The ciliary beat patterns of ex vivo samples (by high-speed video microscopy) were recapitulated, scanning electron microscopy demonstrated excellent ciliation, and cilia could be immuno-fluorescently labelled (anti-alpha-tubulin and anti-RSPH4a) in representative cases that were ALI-cultured after cryostorage. In summary, our ALI culture protocol provides high ciliation rates across three centres, minimising patient recall for repeat brushing biopsies and improving diagnostic certainty. Cryostorage of surplus diagnostic samples was successful, facilitating PCD research.
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Affiliation(s)
- Janice L. Coles
- Primary Ciliary Dyskinesia Centre, NIHR Biomedical Research Centre, University Hospital Southampton NHS Foundation Trust, Southampton SO16 6YD, UK; (J.L.C.); (J.T.); (A.H.); (W.T.W.)
- School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton SO16 6YD, UK; (K.L.H.); (P.M.L.)
| | - James Thompson
- Primary Ciliary Dyskinesia Centre, NIHR Biomedical Research Centre, University Hospital Southampton NHS Foundation Trust, Southampton SO16 6YD, UK; (J.L.C.); (J.T.); (A.H.); (W.T.W.)
- School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton SO16 6YD, UK; (K.L.H.); (P.M.L.)
| | - Katie L. Horton
- School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton SO16 6YD, UK; (K.L.H.); (P.M.L.)
| | - Robert A. Hirst
- Centre for PCD Diagnosis and Research, Department of Respiratory Sciences, University of Leicester, Robert Kilpatrick Clinical Sciences Building, Leicester LE2 7LX, UK; (R.A.H.); (G.M.W.); (C.O.)
| | - Paul Griffin
- Paediatric Respiratory department, Royal Brompton and Harefield NHS Foundation Trust, Sydney Street, London SW3 6NP, UK; (P.G.); (C.H.)
| | - Gwyneth M. Williams
- Centre for PCD Diagnosis and Research, Department of Respiratory Sciences, University of Leicester, Robert Kilpatrick Clinical Sciences Building, Leicester LE2 7LX, UK; (R.A.H.); (G.M.W.); (C.O.)
| | - Patricia Goggin
- Biomedical Imaging Unit, University Hospital Southampton NHS Foundation Trust, Southampton SO16 6YD, UK; (P.G.); (R.D.)
| | - Regan Doherty
- Biomedical Imaging Unit, University Hospital Southampton NHS Foundation Trust, Southampton SO16 6YD, UK; (P.G.); (R.D.)
| | - Peter M. Lackie
- School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton SO16 6YD, UK; (K.L.H.); (P.M.L.)
- Biomedical Imaging Unit, University Hospital Southampton NHS Foundation Trust, Southampton SO16 6YD, UK; (P.G.); (R.D.)
| | - Amanda Harris
- Primary Ciliary Dyskinesia Centre, NIHR Biomedical Research Centre, University Hospital Southampton NHS Foundation Trust, Southampton SO16 6YD, UK; (J.L.C.); (J.T.); (A.H.); (W.T.W.)
- School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton SO16 6YD, UK; (K.L.H.); (P.M.L.)
| | - Woolf T. Walker
- Primary Ciliary Dyskinesia Centre, NIHR Biomedical Research Centre, University Hospital Southampton NHS Foundation Trust, Southampton SO16 6YD, UK; (J.L.C.); (J.T.); (A.H.); (W.T.W.)
| | - Christopher O’Callaghan
- Centre for PCD Diagnosis and Research, Department of Respiratory Sciences, University of Leicester, Robert Kilpatrick Clinical Sciences Building, Leicester LE2 7LX, UK; (R.A.H.); (G.M.W.); (C.O.)
- Respiratory, Critical Care and Anaesthesia, UCL Great Ormond Street Institute of Child Health, 30 Guilford Street, London WC1N 1EH, UK
| | - Claire Hogg
- Paediatric Respiratory department, Royal Brompton and Harefield NHS Foundation Trust, Sydney Street, London SW3 6NP, UK; (P.G.); (C.H.)
| | - Jane S. Lucas
- Primary Ciliary Dyskinesia Centre, NIHR Biomedical Research Centre, University Hospital Southampton NHS Foundation Trust, Southampton SO16 6YD, UK; (J.L.C.); (J.T.); (A.H.); (W.T.W.)
- School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton SO16 6YD, UK; (K.L.H.); (P.M.L.)
| | - Cornelia Blume
- School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton SO16 6YD, UK; (K.L.H.); (P.M.L.)
| | - Claire L. Jackson
- Primary Ciliary Dyskinesia Centre, NIHR Biomedical Research Centre, University Hospital Southampton NHS Foundation Trust, Southampton SO16 6YD, UK; (J.L.C.); (J.T.); (A.H.); (W.T.W.)
- School of Clinical and Experimental Sciences, University of Southampton Faculty of Medicine, Southampton SO16 6YD, UK; (K.L.H.); (P.M.L.)
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10
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Mianné J, Ahmed E, Bourguignon C, Fieldes M, Vachier I, Bourdin A, Assou S, De Vos J. Induced Pluripotent Stem Cells for Primary Ciliary Dyskinesia Modeling and Personalized Medicine. Am J Respir Cell Mol Biol 2019; 59:672-683. [PMID: 30230352 DOI: 10.1165/rcmb.2018-0213tr] [Citation(s) in RCA: 14] [Impact Index Per Article: 2.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/31/2022] Open
Abstract
Primary ciliary dyskinesia (PCD) is a rare and heterogeneous genetic disorder that affects the structure and function of motile cilia. In the airway epithelium, impaired ciliary motion results in reduced or absent mucociliary clearance that leads to the appearance of chronic airway infection, sinusitis, and bronchiectasis. Currently, there is no effective treatment for PCD, and research is limited by the lack of convenient models to study this disease and investigate innovative therapies. Furthermore, the high heterogeneity of PCD genotypes is likely to hinder the development of a single therapy for all patients. The generation of patient-derived, induced pluripotent stem cells, and their differentiation into airway epithelium, as well as genome editing technologies, could represent major tools for in vitro PCD modeling and for developing personalized therapies. Here, we review PCD pathogenesis and then discuss how human induced pluripotent stem cells could be used to model this disease for the development of innovative, patient-specific biotherapies.
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Affiliation(s)
- Joffrey Mianné
- 1 Institute for Regenerative Medicine and Biotherapy, University of Montpellier, Institut National de la Santé et de la Recherche Médicale (INSERM), Centre Hospitalier Universitaire Montpellier, Montpellier, France
| | - Engi Ahmed
- 1 Institute for Regenerative Medicine and Biotherapy, University of Montpellier, Institut National de la Santé et de la Recherche Médicale (INSERM), Centre Hospitalier Universitaire Montpellier, Montpellier, France
| | - Chloé Bourguignon
- 1 Institute for Regenerative Medicine and Biotherapy, University of Montpellier, Institut National de la Santé et de la Recherche Médicale (INSERM), Centre Hospitalier Universitaire Montpellier, Montpellier, France
| | - Mathieu Fieldes
- 1 Institute for Regenerative Medicine and Biotherapy, University of Montpellier, Institut National de la Santé et de la Recherche Médicale (INSERM), Centre Hospitalier Universitaire Montpellier, Montpellier, France
| | - Isabelle Vachier
- 2 PhyMedExp, University of Montpellier, INSERM, Centre Hospitalier Universitaire Montpellier, Montpellier, France; and
| | - Arnaud Bourdin
- 2 PhyMedExp, University of Montpellier, INSERM, Centre Hospitalier Universitaire Montpellier, Montpellier, France; and
| | - Said Assou
- 1 Institute for Regenerative Medicine and Biotherapy, University of Montpellier, Institut National de la Santé et de la Recherche Médicale (INSERM), Centre Hospitalier Universitaire Montpellier, Montpellier, France
| | - John De Vos
- 1 Institute for Regenerative Medicine and Biotherapy, University of Montpellier, Institut National de la Santé et de la Recherche Médicale (INSERM), Centre Hospitalier Universitaire Montpellier, Montpellier, France.,3 Centre Hospitalier Universitaire Montpellier, Department of Cell and Tissue Engineering, Hospital Saint-Eloi, Montpellier, France
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11
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Guimbellot JS, Leach JM, Chaudhry IG, Quinney NL, Boyles SE, Chua M, Aban I, Jaspers I, Gentzsch M. Nasospheroids permit measurements of CFTR-dependent fluid transport. JCI Insight 2017; 2:95734. [PMID: 29202459 DOI: 10.1172/jci.insight.95734] [Citation(s) in RCA: 34] [Impact Index Per Article: 4.9] [Reference Citation Analysis] [Abstract] [Key Words] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/15/2017] [Accepted: 10/11/2017] [Indexed: 12/19/2022] Open
Abstract
Expansion of novel therapeutics to all patients with cystic fibrosis (CF) requires personalized CFTR modulator therapy. We have developed nasospheroids, a primary cell culture-based model derived from individual CF patients and healthy subjects by a minimally invasive nasal biopsy. Confocal microscopy was utilized to measure CFTR activity by analyzing changes in cross-sectional area over time that resulted from CFTR-mediated ion and fluid movement. Both the rate of change over time and AUC were calculated. Non-CF nasospheroids with active CFTR-mediated ion and fluid movement showed a reduction in cross-sectional area, whereas no changes were observed in CF spheroids. Non-CF spheroids treated with CFTR inhibitor lost responsiveness for CFTR activation. However, nasospheroids from F508del CF homozygotes that were treated with lumacaftor and ivacaftor showed a significant reduction in cross-sectional area, indicating pharmacologic rescue of CFTR function. This model employs a simple measurement of size corresponding to changes in CFTR activity and is applicable for detection of small changes in CFTR activity from individual patients in vitro. Advancements of this technique will provide a robust model for individualized prediction of CFTR modulator efficacy.
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Affiliation(s)
| | - Justin M Leach
- Department of Biostatistics, University of Alabama at Birmingham, Birmingham, Alabama, USA
| | | | | | | | - Michael Chua
- Marsico Lung Institute/Cystic Fibrosis Research Center
| | - Inmaculada Aban
- Department of Biostatistics, University of Alabama at Birmingham, Birmingham, Alabama, USA
| | - Ilona Jaspers
- Center for Environmental Medicine, Asthma, and Lung Biology, and
| | - Martina Gentzsch
- Marsico Lung Institute/Cystic Fibrosis Research Center.,Department of Cell Biology and Physiology, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina, USA
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12
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Mitchison HM, Shoemark A. Motile cilia defects in diseases other than primary ciliary dyskinesia: The contemporary diagnostic and research role for transmission electron microscopy. Ultrastruct Pathol 2017; 41:415-427. [PMID: 28925789 DOI: 10.1080/01913123.2017.1370050] [Citation(s) in RCA: 10] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/12/2022]
Abstract
Ultrastructural studies have underpinned the cell biological and clinical investigations of the varied roles of motile cilia in health and disease, with a long history since the 1950s. Recent developments from transmission electron microscopy (TEM; cryo-electron microscopy, electron tomography) have yielded higher resolution and fresh insights into the structure and function of these complex organelles. Microscopy in ciliated organisms, disease models, and in patients with ciliopathy diseases has dramatically expanded our understanding of the ubiquity, multisystem involvement, and importance of cilia in normal human development. Here, we review the importance of motile cilia ultrastructural studies in understanding the basis of diseases other than primary ciliary dyskinesia.
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Affiliation(s)
- Hannah M Mitchison
- a Newlife Birth Defects Research Centre, Experimental and Personalised Medicine, Genetics and Genomic Medicine Programme, UCL Great Ormond Street Institute of Child Health , University College London , London , UK
| | - Amelia Shoemark
- b Department of Paediatric Respiratory Medicine , Royal Brompton & Harefield NHS Trust , London , UK.,c Division of Molecular & Clinical Medicine, School of Medicine , Ninewells Hospital and Medical School , Dundee , UK
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