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

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.