We would like to thank Jean-Fran&#231;ois Papon, Bruno Louis, Estelle Escudier and all team members of PCD diagnostic center of Paris-Est for their availability and hearty welcome during the visit to their PCD diagnostic center, and the numerous exchanges. We also thank Robert Hirst and all team members at the PCD center of Leicester for their welcome and time, advice, and expertise.

Approval was obtained from the Liege hospital-faculty ethics committee and the University Department for Hygiene and Health Protection at Work.
1. Sampling respiratory ciliated epithelium
<ol>
	<li>Ensure that subjects are free of infection for at least 4-6 weeks, and free of nasal and inhaled medication, before sampling.</li>
	<li>Prepare supplemented M199 preparation: Supplement Cell Culture Medium 199 (M199) (500 mL) with antibiotic solution (5 mL of streptomycin/penicillin (50 &#956;g/mL...

This paper aims to provide a standard operating procedure for CFA using nasal brushing samples, with adjustments made for appropriate infection control considerations during the COVID-19 pandemic. PCD diagnosis is challenging, and currently requires a panel of different diagnostic tests, according to international recommendation, including nasal nitric oxide measurement, CFA using DHSV, ciliary ultrastructural analysis using transmission electron microscopy (TEM), labelling of ciliary proteins using immunofluorescence, a...

Primary ciliary dyskinesia (PCD) is an inherited heterogeneous motile ciliopathy, in which respiratory cilia are stationary, slow or dyskinetic, leading to impaired mucociliary clearance and chronic oto-sino-pulmonary disease1,2,3,4. The clinical manifestations of PCD are chronic wet cough and chronic nasal congestion starting in early infancy, recurrent or chronic upper and lower respiratory tract infections leading to bronchiectasis, and recurrent or chronic otitis media and sinusitis5,6,7. Approximately half of PCD patients present with organ laterality defects such as situs inversus or situs ambiguus. Some patients also present with infertility issues due to immotile sperm in men and immotile cilia in the Fallopian tubes in women1,2,8. PCD is rare, but the prevalence is difficult to define, and ranges from 1:10,000 to 1:20,0009,10. However, the real prevalence of PCD is thought to be higher due to difficulties in diagnosis and a lack of clinical suspicion. Symptoms of PCD mimic common respiratory manifestations of other acute or chronic respiratory conditions, and the diagnostic challenges of confirming the diagnosis are well known, leading to inadequate treatment and follow-up2,5,9,11.
Ciliary videomicroscopy, using Digital High-Speed Videomicroscopy (DHSV), is one of the diagnostic tools for PCD4,8,12,13. DHSV is considered the optimal method to perform ciliary functional analysis (CFA), comprising of ciliary beat frequency (CBF) and beat pattern (CBP) analysis2,14,15,16. DHSV uses living respiratory epithelium, usually obtained from nasal brushing13.
In view of the current COVID-19 outbreak, confirmation of a PCD diagnosis is now even more important as evidence suggests that underlying respiratory disease may lead to worse outcomes following COVID-19 infection17,18. A safe and efficient PCD diagnostic service during the current pandemic will also allow confirmed PCD patients to benefit from additional protective measures, compared with the general population19.
Transmission of COVID-19 occurs primarily through droplet spread20. High potential of transmission from asymptomatic (or minimally symptomatic) patients is suggested by the high viral load in nose sample20. Additionally, if viral particles become aerosolized, they stay in the air for at least 3 hours21. Therefore, respiratory healthcare workers are exposed to a high reservoir of viral load while performing clinical care and sample collection for diagnostic techniques22. Furthermore, manipulation of living respiratory samples exposes the technician to COVID-19 contamination. While best-practice recommendations for respiratory physicians and ENT surgeons caring for COVID-19 patients are being implemented23, there is a lack of recommendations for performing DHSV during the COVID-19 pandemic.
In order to continue providing a PCD diagnostic service, while ensuring the safety of the healthcare worker (performing sample collection) and technician (performing sample processing), the ciliary videomicroscopy protocol had to be adapted during the COVID-19 pandemic. The technique of ciliary videomicroscopy is currently limited to research service and specialized diagnostic centers, as CFA requires extensive training and experience. Furthermore, currently, there is a lack of standardization and precise operating procedure for processing and analyzing samples using DHSV4,13.
The aim of this paper is to describe standard operating procedures for DHSV, with particular reference to infection control measures and safety when sampling and processing living nasal epithelium. This will allow for high-quality PCD diagnosis and care to continue, despite the current COVID-19 outbreak.

<table>
	<tbody>
		<tr>
			<td>15 mL conical tubes</td>
			<td>FisherScientific</td>
			<td>352096</td>
			<td>15 ml High-Clarity Polypropylene Conical Tube with lid</td>
		</tr>
		<tr>
			<td>Amphotericin B</td>
			<td>LONZA</td>
			<td>17-836E</td>
			<td>Antifungal solution</td>
		</tr>
		<tr>
			<td>Blakesley-weil nasal forceps</td>
			<td>NOVO SURGICAL</td>
			<td>E7739-12</td>
			<td>Used to hold the brush to perform the nasal brushing</td>
		</tr>
		<tr>
			<td>Bronchial cytology brush</td>
			<td>CONMED</td>
			<td>129</td>
			<td>Used for nasal brushing</td>
		</tr>
		<tr>
			<td>Cotton swab</td>
			<td>NUOVA APTACA</td>
			<td>2150/SG</td>
			<td>Used for COVID-19 testing</td>
		</tr>
		<tr>
			<td>Digitial high-speed videomicroscopy camera</td>
			<td>IDTeu Innovation in motion</td>
			<td>CrashCam Mini 1510</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass slide</td>
			<td>ThermoScientific</td>
			<td>12372098</td>
			<td>Microscope slides used to create the visualization chamber</td>
		</tr>
		<tr>
			<td>Heated Box</td>
			<td>IBIDI cells in focus</td>
			<td>10918</td>
			<td>Used to heat the sample</td>
		</tr>
		<tr>
			<td>Inverted Light microscope</td>
			<td>Zeiss</td>
			<td>AXIO Vert.A1</td>
			<td></td>
		</tr>
		<tr>
			<td>Lens Heater</td>
			<td>TOKAI HIT</td>
			<td>TPiE-LH</td>
			<td>Used to heat the oil immersion lens</td>
		</tr>
		<tr>
			<td>Medium 199 (M199), HEPES</td>
			<td>TermoFisher Scientific</td>
			<td>12340030</td>
			<td>Cell Culture Medium</td>
		</tr>
		<tr>
			<td>Motion Studio X64</td>
			<td>IDT Motion</td>
			<td>version 2.14.01</td>
			<td>Software</td>
		</tr>
		<tr>
			<td>Oil</td>
			<td>FischerScientific, Carl Zeiss</td>
			<td>11825153</td>
			<td></td>
		</tr>
		<tr>
			<td>Rectangular cover slip</td>
			<td>VWR</td>
			<td>631-0145</td>
			<td>Used to cover the visualization chamber</td>
		</tr>
		<tr>
			<td>Spacer (Ispacer) 0.25 mm</td>
			<td>Sunjinlab</td>
			<td>IS203</td>
			<td>Used for the creation of the hermetic closed visualization chamber</td>
		</tr>
		<tr>
			<td>Square cover slip</td>
			<td>VWR</td>
			<td>631-0122</td>
			<td>Used for the creation of lab-built open visualization chamber</td>
		</tr>
		<tr>
			<td>Streptomycin/Penicillin</td>
			<td>FisherScientific, Gibco</td>
			<td>11548876</td>
			<td>Antiobiotics solution</td>
		</tr>
	</tbody>
</table>

nasal brushing sampling and processing using digital high speed ciliary videomicroscopy &ndash; adaptation for the covid-19 pandemic

Primary Ciliary Dyskinesia (PCD) is a genetic motile ciliopathy, leading to significant otosinopulmonary disease. PCD diagnosis is often missed or delayed due to challenges with different diagnostic modalities. Ciliary videomicroscopy, using Digital High-Speed Videomicroscopy (DHSV), one of the diagnostic tools for PCD, is considered the optimal method to perform ciliary functional analysis (CFA), comprising of ciliary beat frequency (CBF) and beat pattern (CBP) analysis. However, DHSV lacks standardized, published operating procedure for processing and analyzing samples. It also uses living respiratory epithelium, a significant infection control issue during the COVID-19 pandemic. To continue providing a diagnostic service during this health crisis, the ciliary videomicroscopy protocol has been adapted to include adequate infection control measures.
Here, we describe a revised protocol for sampling and laboratory processing of ciliated respiratory samples, highlighting adaptations made to comply with COVID-19 infection control measures. Representative results of CFA from nasal brushing samples obtained from 16 healthy subjects, processed and analyzed according to this protocol, are described. We also illustrate the importance of obtaining and processing optimal quality epithelial ciliated strips, as samples not meeting quality selection criteria do now allow for CFA, potentially decreasing the diagnostic reliability and the efficiency of this technique.

To illustrate the efficiency of the technique, we present the results of the CFA in a series of 16 healthy adult volunteers (5 males, age range 22-54 years).
Nasal brushing samples from 14 (4 males, age range 24-54 years) out of the total of 16 volunteers provided enough appropriate epithelial edges that satisfied the selection criteria needed to perform CFA. From these 14 nasal brushing samples, a total of 242 ciliated edges were recorded, and 212 edges met the defined inclusion criteria and ...

Watch this Scientific Journal Video about Nasal Brushing Sampling and Processing Using Digital High Speed Ciliary Videomicroscopy â€“ Adaptation for the COVID-19 Pandemic at JoVE.com

Nasal Brushing Sampling and Processing Using Digital High Speed Ciliary Videomicroscopy &#8211; Adaptation for the COVID-19 Pandemic

To guarantee a successful and high-quality ciliary functional analysis for PCD diagnosis, a precise and careful method for respiratory epithelium sampling and processing is essential. To continue providing PCD diagnostic service during the COVID-19 pandemic, the ciliary videomicroscopy protocol has been updated to include appropriate infection control measures.

Nasal Brushing Sampling and Processing Using Digital High Speed Ciliary Videomicroscopy &ndash; Adaptation for the COVID-19 Pandemic

Primary Ciliary Dyskinesia (PCD) is a genetic motile ciliopathy, leading to significant otosinopulmonary disease. PCD diagnosis is often missed or delayed ...

The lack of published formal operating procedures and standardization prevents digital high-speed videomicroscopy from being considered as a confirmatory diagnostic test for primary ciliary dyskinesia. Furthermore, to continue providing a diagnostic service during this COVID-19 crisis, the ciliary videomicroscopy protocol had been adapted to include adequate infection control measures. The ciliary functional evaluation using digital high-speed videomicroscopy is highly sensitive and specific for primary ciliary dyskinesia diagnosis.Compared with other primary ciliary dyskinesia diagnostic tests, digital high-speed videomicroscopy is relatively easy to perform, inexpensive, and results may be available within the day. Digital high-speed videomicroscopy has a higher sensitivity and specificity for primary ciliary dyskinesia diagnosis. Currently, only electron microscopy and genetic testing are recognized as confirmatory tests for primary ciliary dyskinesia, but these tests will miss primary ciliary dyskinesia diagnosis in 15 to 30%of patients.Demonstrating the nasal brushing procedure will be Lionel Benchimol, an ENT resident from my laboratory. Before collecting the nasal sample, have the patient blow their nose. After clearing the nasal passages, have the patient lie or sit comfortably with the head resting backwards.Shake the brush in the supplemented Medium 199 to moisten it, and place an endoscope at the entrance of the nose to visualize the inferior nasal turbinate. Insert the cytology brush into the nose, and move the brush posteriorly and anteriorly several times over the posterior part of the inferior nasal turbinate before withdrawing. Place the brush into a tube of M199.Cut the wire so that it can fit completely inside the tube, and immediately close the tube. Then, place the sample in an airtight double bag. Within nine hours of sample collection, open the sample tubes in a microbiological safety cabinet, and use Weil-Blakesley nasal forceps to agitate the brush to dislodge the collected epithelial strips.Stick a double-sided spacer onto a glass slide, and remove the protection from the spacer. Gently shake the tube to allow the cilia to spread throughout the tube, and use a pipette to transfer approximately 60 microliters of ciliated epithelium from the middle of the tube into the spacer. Place a 22-by-4-millimeter rectangular cover slip onto the spacer to close the chamber, and disinfect the slide with 70%ethanol before removing it from the microbiological safety cabinet.After changing gloves, place this slide on the plate of a heated box, and close the lid. Add oil to the oil immersion objective, and place the box onto the stage of an upright or an inverted light microscope. Turn on the box and the lens heater, and adjust the temperature settings on the box and lens heater controllers.After five minutes, position the objective until it is just touching the cover slip with the tip of the lens. To visualize the respiratory ciliated edges of the sample, open the camera software, then use the microscope objective to manually locate the cells within the sample and to project the cells onto the monitor. Check that the image is visible on the monitor, and adjust the condenser and lens focus as necessary to improve the quality of the image.When the strips of ciliated epithelium have been located, select only intact, undisrupted ciliated epithelial edges that measure at least 50 microns in length. Be sure to use only normal edges or edges with minor projections for ciliary functional analysis and to exclude ciliated edges with major projections and isolated ciliated cells or single cells with no contact between themselves and any other cell type. When a good edge has been selected, use a camera frame rate of 500 frames per second to record the beating cilia edge for approximately two seconds.After recording a minimum of six good edges in side view, remove the slide from the heated box, and place the slide, cover slip, and spacer into an airtight bag. Then, place the gloves and mask into the bag, and place the bag into the appropriate hazardous medical waste container. To analyze the ciliary beat frequency, open the recording in an appropriate video analysis software program, and divide the ciliated edges into approximately five adjacent approximately 10-micron areas.Identify and visualize the cilia or groups of cilia at a reduced frame rate and a maximum of two ciliary beat frequency measurements per area to obtain a maximum of 10 ciliary beat frequency measurements per edge. Record the number of frames required for a group of cilia to complete five beat cycles, and use the formula to calculate the ciliary beat frequency. To determine the percentage of each distinct ciliary beat pattern within a sample, for each cilia or group of cilia, compare the precise path taken by the cilia during a full beat cycle with the normal ciliary beat pattern observed on the digital high-speed videomicroscopy analysis.Attribute a distinct ciliary beat pattern, such as normal, stiff, immotile, asynchronous or dyskinetic, and circular, to each cilia or group of cilia analyzed. Then, calculate the percentage of each distinct ciliary beat pattern within each sample. The ciliary beat pattern attributed to the sample is the predominant ciliary beat pattern observed.Here, we observe detailed ciliary functional analysis results from the nasal brushing samples of 14 adult volunteers. From these 14 nasal brushing samples, 242 ciliated edges were recorded, and 212 met the defined inclusion criteria for analysis. A total of 807 ciliary beat frequency measurements and ciliary beat pattern evaluations were obtained from the cilia or groups of cilia analyzed.The mean immotility index were similar to the previously reported values observed in healthy volunteers. The dyskinesia score and the ciliary beat frequency were similar to the previously reported values obtained when selecting only normal edges or edges with minor projection. When low-quality edges are used, lower ciliary beat frequencies and higher dyskinesia scores are reported.Blood cells and mucus acquired from a too robust brushing can block free cilia beating or hide cilia from the observer. Conversely, if the brush does not press firmly against the inferior nasal turbinate, the sample may not contain enough high-quality ciliated epithelial strips. In addition, if the nasal brushing is performed on the anterior part of the nasal cavity, only transitional non-ciliated epithelial cells will be obtained.The two most important aspects of the protocol are the quality of nasal brushing and the choice of high-quality epithelial fragments to avoid a full positive result or a failure of the technique.

Nasal Brushing Sampling and Processing Using Digital High Speed Ciliary Videomicroscopy – Adaptation for the COVID-19 Pandemic

Abstract