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Brahim Belhaouari D, Wurtz N, Grimaldier C, Lacoste A, Pires de Souza GA, Penant G, Hannat S, Baudoin JP, La Scola B. Microscopic Observation of SARS-Like Particles in RT-qPCR SARS-CoV-2 Positive Sewage Samples. Pathogens 2021; 10:516. [PMID: 33923138 PMCID: PMC8146039 DOI: 10.3390/pathogens10050516] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [What about the content of this article? (0)] [Affiliation(s)] [Abstract] [Key Words] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/02/2021] [Revised: 04/20/2021] [Accepted: 04/21/2021] [Indexed: 12/18/2022] Open
Abstract
The ongoing outbreak of novel coronavirus pneumonia (COVID-19) caused by SARS-CoV-2 infection has spread rapidly worldwide. The major transmission routes of SARS-CoV-2 are recognised as inhalation of aerosol/droplets and person-to-person contact. However, some studies have demonstrated that live SARS-CoV-2 can be isolated from the faeces and urine of infected patients, which can then enter the wastewater system. The currently available evidence indicates that the viral RNA present in wastewater may become a potential source of epidemiological data. However, to investigate whether wastewater may present a risk to humans such as sewage workers, we investigated whether intact particles of SARS-CoV-2 were observable and whether it was possible to isolate the virus in wastewater. Using a correlative strategy of light microscopy and electron microscopy (CLEM), we demonstrated the presence of intact and degraded SARS-like particles in RT-qPCR SARS-CoV-2-positive sewage sample collected in the city of Marseille. However, the viral infectivity assessment of SARS-CoV-2 in the wastewater was inconclusive, due to the presence of other viruses known to be highly resistant in the environment such as enteroviruses, rhinoviruses, and adenoviruses. Although the survival and the infectious risk of SARS-CoV-2 in wastewater cannot be excluded from our study, additional work may be required to investigate the stability, viability, fate, and decay mechanisms of SARS-CoV-2 thoroughly in wastewater.
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Affiliation(s)
- Djamal Brahim Belhaouari
- Microbes, Evolution, Phylogeny and Infection (MEPHI), UM63, Institut de Recherche pour le Développement (IRD), Aix- Marseille University, 13005 Marseille, France; (D.B.B.); (N.W.); (G.A.P.d.S.); (S.H.)
- IHU Méditerranée Infection, 13005 Marseille, France
| | - Nathalie Wurtz
- Microbes, Evolution, Phylogeny and Infection (MEPHI), UM63, Institut de Recherche pour le Développement (IRD), Aix- Marseille University, 13005 Marseille, France; (D.B.B.); (N.W.); (G.A.P.d.S.); (S.H.)
- IHU Méditerranée Infection, 13005 Marseille, France
| | - Clio Grimaldier
- Assistance Publique—Hôpitaux de Marseille, 13005 Marseille, France; (C.G.); (G.P.)
| | - Alexandre Lacoste
- Bataillon des Marins Pompiers de Marseille, 13003 Marseille, France;
| | - Gabriel Augusto Pires de Souza
- Microbes, Evolution, Phylogeny and Infection (MEPHI), UM63, Institut de Recherche pour le Développement (IRD), Aix- Marseille University, 13005 Marseille, France; (D.B.B.); (N.W.); (G.A.P.d.S.); (S.H.)
- IHU Méditerranée Infection, 13005 Marseille, France
| | - Gwilherm Penant
- Assistance Publique—Hôpitaux de Marseille, 13005 Marseille, France; (C.G.); (G.P.)
| | - Sihem Hannat
- Microbes, Evolution, Phylogeny and Infection (MEPHI), UM63, Institut de Recherche pour le Développement (IRD), Aix- Marseille University, 13005 Marseille, France; (D.B.B.); (N.W.); (G.A.P.d.S.); (S.H.)
- IHU Méditerranée Infection, 13005 Marseille, France
| | - Jean-Pierre Baudoin
- Assistance Publique—Hôpitaux de Marseille, 13005 Marseille, France; (C.G.); (G.P.)
| | - Bernard La Scola
- Microbes, Evolution, Phylogeny and Infection (MEPHI), UM63, Institut de Recherche pour le Développement (IRD), Aix- Marseille University, 13005 Marseille, France; (D.B.B.); (N.W.); (G.A.P.d.S.); (S.H.)
- IHU Méditerranée Infection, 13005 Marseille, France
- Assistance Publique—Hôpitaux de Marseille, 13005 Marseille, France; (C.G.); (G.P.)
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Heiligenstein X, de Beer M, Heiligenstein J, Eyraud F, Manet L, Schmitt F, Lamers E, Lindenau J, Kea-Te Lindert M, Salamero J, Raposo G, Sommerdijk N, Belle M, Akiva A. HPM live μ for a full CLEM workflow. Methods Cell Biol 2021; 162:115-149. [PMID: 33707009 DOI: 10.1016/bs.mcb.2020.10.022] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.7] [Reference Citation Analysis] [What about the content of this article? (0)] [Affiliation(s)] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/08/2023]
Abstract
With the development of advanced imaging methods that took place in the last decade, the spatial correlation of microscopic and spectroscopic information-known as multimodal imaging or correlative microscopy (CM)-has become a broadly applied technique to explore biological and biomedical materials at different length scales. Among the many different combinations of techniques, Correlative Light and Electron Microscopy (CLEM) has become the flagship of this revolution. Where light (mainly fluorescence) microscopy can be used directly for the live imaging of cells and tissues, for almost all applications, electron microscopy (EM) requires fixation of the biological materials. Although sample preparation for EM is traditionally done by chemical fixation and embedding in a resin, rapid cryogenic fixation (vitrification) has become a popular way to avoid the formation of artifacts related to the chemical fixation/embedding procedures. During vitrification, the water in the sample transforms into an amorphous ice, keeping the ultrastructure of the biological sample as close as possible to the native state. One immediate benefit of this cryo-arrest is the preservation of protein fluorescence, allowing multi-step multi-modal imaging techniques for CLEM. To minimize the delay separating live imaging from cryo-arrest, we developed a high-pressure freezing (HPF) system directly coupled to a light microscope. We address the optimization of sample preservation and the time needed to capture a biological event, going from live imaging to cryo-arrest using HPF. To further explore the potential of cryo-fixation related to the forthcoming transition from imaging 2D (cell monolayers) to imaging 3D samples (tissue) and the associated importance of homogeneous deep vitrification, the HPF core technology has been revisited to allow easy modification of the environmental parameters during vitrification. Lastly, we will discuss the potential of our HPM within CLEM protocols especially for correlating live imaging using the Zeiss LSM900 with electron microscopy.
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Affiliation(s)
| | - Marit de Beer
- Electron Microscopy Center, Radboudumc Technology Center Microscopy, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands; Department of Cell Biology, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands; Department of Biochemistry, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands
| | | | | | | | | | | | | | - Mariska Kea-Te Lindert
- Electron Microscopy Center, Radboudumc Technology Center Microscopy, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands; Department of Cell Biology, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands
| | - Jean Salamero
- SERPICO Inria Team/UMR 144 CNRS & National Biology and Health Infrastructure "France Bioimaging", Institut Curie, Paris, France
| | - Graça Raposo
- Institut Curie, PSL Research University, CNRS, UMR144, Cell and Tissue Imaging Facility (PICT-IBiSA), Paris, France; Institut Curie, PSL Research University, CNRS, UMR144, Structure and Membrane Compartments, Paris, France
| | - Nico Sommerdijk
- Electron Microscopy Center, Radboudumc Technology Center Microscopy, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands; Department of Biochemistry, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands
| | | | - Anat Akiva
- Electron Microscopy Center, Radboudumc Technology Center Microscopy, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands; Department of Cell Biology, Radboud Institute of Molecular Life Sciences, Radboud University Medical Center, Nijmegen, The Netherlands.
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Betteridge KB, Arkill KP, Neal CR, Harper SJ, Foster RR, Satchell SC, Bates DO, Salmon AHJ. Sialic acids regulate microvessel permeability, revealed by novel in vivo studies of endothelial glycocalyx structure and function. J Physiol 2018; 595:5015-5035. [PMID: 28524373 PMCID: PMC5538239 DOI: 10.1113/jp274167] [Citation(s) in RCA: 84] [Impact Index Per Article: 14.0] [Reference Citation Analysis] [What about the content of this article? (0)] [Affiliation(s)] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/10/2017] [Accepted: 05/08/2017] [Indexed: 12/15/2022] Open
Abstract
Key points We have developed novel techniques for paired, direct, real‐time in vivo quantification of endothelial glycocalyx structure and associated microvessel permeability. Commonly used imaging and analysis techniques yield measurements of endothelial glycocalyx depth that vary by over an order of magnitude within the same vessel. The anatomical distance between maximal glycocalyx label and maximal endothelial cell plasma membrane label provides the most sensitive and reliable measure of endothelial glycocalyx depth. Sialic acid residues of the endothelial glycocalyx regulate glycocalyx structure and microvessel permeability to both water and albumin.
Abstract The endothelial glycocalyx forms a continuous coat over the luminal surface of all vessels, and regulates multiple vascular functions. The contribution of individual components of the endothelial glycocalyx to one critical vascular function, microvascular permeability, remains unclear. We developed novel, real‐time, paired methodologies to study the contribution of sialic acids within the endothelial glycocalyx to the structural and functional permeability properties of the same microvessel in vivo. Single perfused rat mesenteric microvessels were perfused with fluorescent endothelial cell membrane and glycocalyx labels, and imaged with confocal microscopy. A broad range of glycocalyx depth measurements (0.17–3.02 μm) were obtained with different labels, imaging techniques and analysis methods. The distance between peak cell membrane and peak glycocalyx label provided the most reliable measure of endothelial glycocalyx anatomy, correlating with paired, numerically smaller values of endothelial glycocalyx depth (0.078 ± 0.016 μm) from electron micrographs of the same portion of the same vessel. Disruption of sialic acid residues within the endothelial glycocalyx using neuraminidase perfusion decreased endothelial glycocalyx depth and increased apparent solute permeability to albumin in the same vessels in a time‐dependent manner, with changes in all three true vessel wall permeability coefficients (hydraulic conductivity, reflection coefficient and diffusive solute permeability). These novel technologies expand the range of techniques that permit direct studies of the structure of the endothelial glycocalyx and dependent microvascular functions in vivo, and demonstrate that sialic acid residues within the endothelial glycocalyx are critical regulators of microvascular permeability to both water and albumin. We have developed novel techniques for paired, direct, real‐time in vivo quantification of endothelial glycocalyx structure and associated microvessel permeability. Commonly used imaging and analysis techniques yield measurements of endothelial glycocalyx depth that vary by over an order of magnitude within the same vessel. The anatomical distance between maximal glycocalyx label and maximal endothelial cell plasma membrane label provides the most sensitive and reliable measure of endothelial glycocalyx depth. Sialic acid residues of the endothelial glycocalyx regulate glycocalyx structure and microvessel permeability to both water and albumin.
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Affiliation(s)
- Kai B Betteridge
- Bristol Renal, Schools of Clinical Sciences and Physiology & Pharmacology, Dorothy Hodgkin Building, University of Bristol, Bristol, BS1 3NY, UK
| | - Kenton P Arkill
- School of Medicine, Faculty of Medicine and Health Sciences, University of Nottingham Medical School, Nottingham, NG7 2UH, UK.,Biofisika Institute (CSIC UPV/EHU) and Research Centre for Experimental Marine Biology and Biotechnology (PiE), University of the Basque Country, Spain
| | - Christopher R Neal
- Bristol Renal, Schools of Clinical Sciences and Physiology & Pharmacology, Dorothy Hodgkin Building, University of Bristol, Bristol, BS1 3NY, UK
| | - Steven J Harper
- Bristol Renal, Schools of Clinical Sciences and Physiology & Pharmacology, Dorothy Hodgkin Building, University of Bristol, Bristol, BS1 3NY, UK
| | - Rebecca R Foster
- Bristol Renal, Schools of Clinical Sciences and Physiology & Pharmacology, Dorothy Hodgkin Building, University of Bristol, Bristol, BS1 3NY, UK
| | - Simon C Satchell
- Bristol Renal, Schools of Clinical Sciences and Physiology & Pharmacology, Dorothy Hodgkin Building, University of Bristol, Bristol, BS1 3NY, UK
| | - David O Bates
- School of Medicine, Faculty of Medicine and Health Sciences, University of Nottingham Medical School, Nottingham, NG7 2UH, UK
| | - Andrew H J Salmon
- Bristol Renal, Schools of Clinical Sciences and Physiology & Pharmacology, Dorothy Hodgkin Building, University of Bristol, Bristol, BS1 3NY, UK.,Renal Service, Specialist Medicine and Health of Older People, Waitemata DHB, Auckland, New Zealand
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