Name: label-free neutrophil enrichment from patient-derived airway secretion using closed-loop inertial microfluidics
Uploaded: 2018-06-07T14:30:00.000+00:00
Duration: 7 min 37 s
Description: Watch this Scientific Journal Video about Label-free Neutrophil Enrichment from Patient-derived Airway Secretion Using Closed-loop Inertial Microfluidics at JoVE.com

This work was supported by NIH/NIAID (R21AI119042) as well as NIH U24 Sample Sparing assay program (U24-AI118656).

The sample collection was approved by the University of Pittsburgh Institutional Review Board (IRB# PRO16060443, PRO10110387). All experiments are performed under a biosafety cabinet with the proper personal protective equipment.
1. Device Fabrication and Soft Lithography
NOTE: Standard soft lithography techniques15,16 were used to create the polydimethylsiloxane (PDMS) microchannel.
<ol>
	<li>Mix the PDMS precursor in a 10:1 ratio of base and curing agent.</li>
	<li>Pour 30 g of the PDMS precursor mixture on the micro-machined aluminum mold (see Figure 1 for dimensions) and 20 g of the PDMS precursor mixture on the 100 mm Petri dish. 
	NOTE: The Petri dish is used to make the thick film of PDMS (~3 mm) for the supporting layer. The supporting layer of PDMS provides uniform physical surface properties throughout the microfluidics. Alternatively, a thin PDMS film on the glass slide can be used. The master mold with specific channel dimensions was designed and fabricated by micro-milling on the aluminum sheet. The spiral channel used in this study was an 8-loop spiral microchannel with one inlet and two outlets, with the radius increasing from 8 mm to 24 mm for efficient sized-based separation.</li>
	<li>Place the mold and the Petri dish into a vacuum desiccator to degas until no bubbles are visible on the surface, typically 5 - 10 min. Use a house-vacuum for the desiccator.</li>
	<li>Release the vacuum and place the mold and the Petri dish in a 90 &#176;C oven for 1 h. 
	NOTE: A hotplate or other heating tools can be used instead of an oven.</li>
	<li>Remove the mold and the Petri dish from the oven and allow it to cool at room temperature for 10 min.</li>
	<li>Carefully cut out the outline and punch the fluid access holes on the device using a 2 mm puncher. 
	NOTE: The size of the punch can vary depending on the size of the tubing or fluid guide.</li>
	<li>Adhere a tape to the channel side of the device and thick film of PDMS, and peel carefully with forceps to remove dust.</li>
	<li>Treat both the channel side of the device and plain PDMS with air plasma and bond with the prepared supporting layer of plain PDMS (Figure 1A).</li>
	<li>Place the chip on the 50 mm x 70 mm glass slide and place it in a 70 &#176;C oven for at least 30 min to enhance the bonding strength.</li>
</ol>
2. Tracheal Secretion Collection from Mechanically Ventilated Patients
NOTE: Tracheal secretions can be obtained from mechanically ventilated patients during normal routine airway care by using a protocol modified from conventional methods to accommodate the standard, adult ventilated patient. Samples were de-identified and sent immediately for processing.
<ol>
	<li>If tracheal aspiration is indicated, use a catheter to extract airway secretions.</li>
	<li>Advance the catheter carefully half-way into the endotracheal tube and instill 5 mL of 0.9% sterile normal saline from a pre-filled 10 mL syringe.</li>
	<li>When the catheter is advanced fully, or resistance is met, aspirate the secretions and collect into a sterile sputum collection container.</li>
	<li>Collect 10 mL of tracheal secretions in a sterile sputum container and place it on ice. Send samples immediately for processing.</li>
</ol>
3. Tracheal Secretion Sample Preparation
NOTE: All experiments must be performed under a biosafety cabinet with the proper personal protective equipment.
<ol>
	<li>Disperse 1 mL of airway secretion samples in 9 mL of phosphate-buffered saline (PBS) using a plastic 10 mL syringe (for a 10x dilution). Use a blunt pipette to homogenize the mucus sample.</li>
	<li>Strain the diluent with a 40 &#181;m nylon cell strainer to remove large chunks or blood clots, which can block the microfluidics access hole or the channel. Place the sample on the ice after processing.</li>
	<li>Disperse the diluent of each sample using a 100x volume of PBS buffer, resulting in a 1,000x diluted suspension. Place the sample on the ice during the entire dissociation operation.</li>
</ol>
4. Experimental Setup
NOTE: All experiments must be performed under a biosafety cabinet with the proper personal protective equipment.
<ol>
	<li>Assemble the PDMS chip with the fluid guide to apply uniform flow to each of the four spiral microchannels (Figure 1B). 
	NOTE: Four channels were used to increase the throughput by parallelization, as well as optimization of the volume and cell density of the resulting suspension. The fluid guide is designed and made with a stereolithography type 3D printer with clear resin. The inlet and the outlet port of the fluid guide use female Luer connectors for ease of connection and stable sealing during the operation.</li>
	<li>Connect a 1/16-inch male Luer connector to the inlet, the inner wall (IW) outlet, and the outer wall (OW) outlet port of the fluid guide and connect a silicone tubing to the sample suspension.</li>
	<li>Insert blunt tips to each end of the inlet and outlets to reach the bottom of the sample reservoir.</li>
	<li>Connect the peristaltic pump with the inlet tubing.</li>
	<li>Place the end of the inlet tubing and IW outlet tubing in the sample reservoir and place the end of the OW outlet tubing in the waste reservoir (Figure 1C, D).</li>
	<li>Place the 50 mL tube of filtered PBS without calcium and magnesium at the sample reservoir.</li>
	<li>Start pumping at a low flow rate (~1 mL/min) to prime the device. 
	NOTE: When bubbles are captured in the channel, push the top of the channel with the tweezer to destabilize and eliminate air bubbles. When the device is fully filled with buffer solution, change the PBS tube to prepare the airway secretion sample tube.</li>
	<li>When the sample is placed in the sample reservoir, switch on the peristaltic pump and set the flow rate at 4 mL/min (Figure 1C, D).</li>
	<li>When the sample volume reaches the designated volume (~1 mL), stop the operation and disconnect the silicone tubing. 
	NOTE: The proposed method is more efficient at reducing a large volume of diluent (&#62;50 mL) into a micro-centrifuge tube volume (~1 mL) (Figure 2).</li>
</ol>
5. Flow Cytometry Analysis
NOTE: To compare the dissociation methods (microfluidics and lysis), flow cytometry with immunofluorescence was used.
<ol>
	<li>Add fluorescein isothiocyanate (FITC)-conjugated mouse anti-human CD66b monoclonal antibody and allophycocyanin (APC)-conjugated mouse anti-human CD45 monoclonal antibody to the initial suspensions (step 3.3) and resulting suspensions (step 4.9) (200 &#181;L of each antibody) in the ratio of 1:25 v/v.</li>
	<li>Incubate the samples at 4 &#176;C in the dark for 30 min.</li>
	<li>Analyze both samples on a flow cytometer to quantify the PMNs. 
	NOTE: The population that is FITC-APC double positive, CD66b positive and CD45 positive was considered to be neutrophils. Recovery was calculated by the ratio of the total cell numbers of the input and the resulting suspension.</li>
</ol>
6. NE Release Analysis
NOTE: To compare the functionality of the isolated neutrophil by the microfluidics and lysis method, an NE assay kit was used.
<ol>
	<li>Add PMA (provided in the assay kit) to each resulting suspension (obtained at step 4.9) to a final concentration of 50 nM.</li>
	<li>Incubate the samples at 37 &#176;C for 2 h.</li>
	<li>Centrifuge the suspensions at 300 x g for 10 min.</li>
	<li>Transfer 10 &#181;L of each supernatant to a 96-well plate (provided in the assay kit). 
	NOTE: A 96-well microplate with a flat bottom and black polystyrene can be used as well.</li>
	<li>Add 90 &#181;L of diluted assay buffer (provided in the assay kit) to each well.</li>
	<li>Add 10 &#181;L of the elastase substrate (Z-Ala-Ala-Ala-Ala 2Rh110, provided in the assay kit).</li>
	<li>Incubate for 1.5 h at 37 &#176;C.</li>
	<li>Read the 96-well plate using a fluorometer at an excitation wavelength of 485 nm and an emission wavelength of 525 nm. 
	NOTE: The level of NE was divided by the PMN count from the flow cytometry analysis.</li>
</ol>

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The authors have filed a patent application on the technology described here.

In inertial microfluidics, particle and cells localize at a certain lateral position in a micro-channel based on their size5,18,19,20. Due to the combined effect of the Dean drag force and the inertial lift force in the curved microchannel, large particles or neutrophils (&#62;10 &#181;m) are located inside the channel and small particles, mucus aggregates, and debris smaller than 6 &#181;m are positioned outside the channel at certain flow velocity conditions10,11,13,14.
However, there is a trade-off between the concentration and recovery of the separation when using microfluidics, especially when the separation size range of the particle or the cell of interest overlaps with other cells/particles. To achieve a high concentration of sorted particles and cells, the separation range needs to be narrow and requires a fine-tuning with microscopic observation for each sample run; this limits the usage of inertial microfluidics on heterogeneous biometrics, such as airway secretions.
Here, we introduce a clinical airway secretions sample preparation method for in vitro assays using closed-loop inertial microfluidics (Figure 1B, C). The closed-loop operation by recirculation of the particle/cell through a focusing outlet achieved both high concentration factors and high recovery rates. The OW outlet flows into the waste tube, and the IW outlet is continuously fed to the original sample tube. Cells larger than 10 &#181;m in diameter remain in the initial sample tube, thus, increasing the cell density of the sample, while the background fluid is eliminated to the waste tube. The high dilution rate allows clearance of the background fluid, including debris and mucus. Also, regardless of the condition of patient sample, the proposed method can uniformly dissociate and isolate immune cells.
DTT is a widely used mucin lysis buffer that cleaves disulfide bonds in proteoglycan aggregation to separate cell content from the mucilage1,21,22. However, DTT reportedly interferes with the white blood cell surface antigens, which can affect leukocyte function2. In our study, the cellular contents recovered by DTT dissociation were inconsistent across samples, because of non-uniform mucolytic efficiency and clogging of the filter. This becomes more critical for the preparation of initial samples with small volumes and relatively thick airway secretion samples. On the other hand, the method using microfluidics enabled the consistent recovery of cell contents in heterogeneous patient samples (Figure 3C). As shown in Figure 3B, we could consistently isolate the cell contents of the airway secretion in a clean buffer regardless of the original sample state, i.e., bloody, tenacious, or watery. Compared to the conventional method, the proposed method has less debris in the recovered sample, which minimizes possible interference of debris in downstream biochemical analyses.
NE release was examined to evaluate the functional integrity of captured neutrophils by both enrichment methods (Figure 3D). Neutrophils remove foreign organic molecules through NE release23,24,25. PMA is commonly used to activate neutrophils in vitro26. Cells isolated by the microfluidics and the mucolytic (DTT) methods were stimulated with 50 nM of PMA. We tested each method with human airway secretion samples (labeled as P173, P174, P176, P177, P181, and P182). Samples concentrated by microfluidics showed an increase in NE by PMA stimulation for all 6 samples. In contrast, only 3 of the DTT treated samples could induce increased NE activity after PMA stimulation. In addition, the neutrophils isolated by the DTT mucolytic method showed much higher baseline activity than the neutrophils isolated by microfluidics. These results showed possible chemical disturbance of the NE machinery, i.e., disruption of the surface bound antigen2,3, which suggests that the microfluidics method is an improved method in terms of preserving the neutrophil function than DTT homogenization. The proposed dissociation method also operates in an automated and fully contained manner, minimizing the exposure of patient samples with potential hazards of the external environment.
However, the proposed method requires a high flow rate and, therefore, a stable fluidic connection. Also, due to the inherent fluctuation of the peristaltic circulation pump, the IW outlet dimension must be greater than the OW outlet dimension, otherwise throughput may be compromised. We expect that the proposed method will provide higher throughput if there is a circulation pump that can provide a more stable flow rate.
By providing a dissociation protocol to capture neutrophils from clinical airway secretions, the method presented in this study is expected to expand the scope of clinical research on respiratory diseases, such as pneumonia, asthma, cystic fibrosis, and chronic obstructive pulmonary disease.

Since cells are encapsulated in a large amount of mucus in airway secretions, the functional assessment of leukocytes by an in vitro assay has been hindered. Dithiothreitol (DTT) is the most common lysis buffer to dissociate and homogenize the sputum for cytological analysis and detection of mediators while providing tolerable viability of isolated cells1,2. However, DTT can interfere with surface-bound antigens of airway neutrophils, resulting in the disruption of neutrophil function such as elastase and myeloperoxidase (MPO) release2,3.Therefore, few studies of human airway neutrophil function have been conducted with peripheral blood neutrophils, which may not reveal the physiological characteristics of pulmonary4. Meanwhile, inertial microfluidics has made advances in isolating cells from various patient biomatrices5,6.The equilibrium between inertial lift forces and Dean drag aligns the particle/cell according to their size, which allows label-free particle separation7. Our group previously introduced a sample preparation method for circulating tumor cells8,9, pathogens in blood8, cells from a suspension culture10,11,12, and polymorphonuclear leukocytes (PMNs) from blood13,14.
Here, we introduce a protocol to prepare immune cells from a patient&#39;s airway secretions using closed-loop inertial microfluidics for a downstream in vitro assay, such as the neutrophil elastase (NE) assay. This method provides both high concentration and recovery, especially when there is a significant overlap in the lateral direction of the cell/particle from which the cell/particle-of-interest is to be removed, which is commonly observed in clinical samples. By recirculating the Inner wall (IW)-focused large particles or cells back to the input sample tube, the particle or cell-of-interest concentrates in the original reservoir, while background fluids with small mucin aggregates pass through the waste reservoir. Despite the heterogeneous fluidic properties of clinical samples, the proposed method recovers consistently above 95% of neutrophils from airway secretion samples that are diluted 1,000-fold with a clean saline solution (~1 mL). By contrast, the lysis method presents a wide range of PMNs recovery rates depending on the sample condition. The proposed protocol captures leukocytes in a label-free manner with no physical or chemical disruption, which provides the possibility to harvest delicate cells from clinically challenging biometrics with minimally invasive procedures.

<table><tbody><tr><td>PDMS precursor</td><td>Dow corning</td><td>184 SIL ELAST KIT 3.9KG</td><td>10:1 ratio of base and curing agent</td></tr><tr><td>VWR gravity convection oven</td><td>VWR</td><td>414005-128</td><td>PDMS precursor to be cured in 90 deg.</td></tr><tr><td>100mm petri dish</td><td>VWR</td><td>89000-324</td><td>Fabrication of PDMS Supporting layer</td></tr><tr><td>Harris Uni-core puncher</td><td>Sigma-aldrich</td><td>WHAWB100076</td><td>2mm diameter or other depending on the tubing size</td></tr><tr><td>Air plasma machine</td><td>Femto Science</td><td>Cute</td><td>Surface plasma treatment for PDMS device to bottom base.</td></tr><tr><td>2&amp;rdquo; x 3&amp;rdquo; glass slide</td><td>TED PELLA, INC.</td><td>2195</td><td>To support PDMS device</td></tr><tr><td>Masterflex spooled platinum-cured silicone tubing, L/S 14</td><td>Cole-Parmer</td><td>EW-96410-14</td><td>Tubing for microfluidics and peristlatic pump</td></tr><tr><td>1/16 inch Luer connector, male</td><td>Harvard apparatus</td><td>PC2 72-1443</td><td>Connector for fluid guide</td></tr><tr><td>50mL Falcon tube</td><td>Corning</td><td>21008-940</td><td>sample collection &amp;amp; preparation</td></tr><tr><td>Phosphate-Buffered Saline, 1X Without Calcium and Magnesium</td><td>Corning</td><td>45000-446&amp;nbsp;</td><td>Buffer solution to dilute sample</td></tr><tr><td>Halyard Closed suction Catheter, Elbow, 14F/ channel 4.67mm</td><td>HALYARD HEALTH</td><td>22113</td><td>Tracheal seceation suction catheter</td></tr><tr><td>0.9% Sterile Normal saline, 10mL pre-filled syringe</td><td>BD PosiFlush</td><td>NHRIC: 8290-306547</td><td>For tracheal seceation collection from the patients</td></tr><tr><td>SecurTainer&amp;trade; III Specimen Containers, 20mL</td><td>Simport</td><td>1176R36</td><td>Sterile sputum (airway secretion) collection container</td></tr><tr><td>Syringe with Luer-Lok Tip, 10mL</td><td>BD</td><td>BD309604</td><td>To pipette homogenize the mucus sample and reach the bottom of sample tube</td></tr><tr><td>BD&amp;nbsp; Blunt Fill Needle, with BD Luer-Lok&amp;nbsp; Tip</td><td>BD</td><td></td><td>To pipette homogenize the mucus sample and reach the bottom of sample tube</td></tr><tr><td>40&amp;micro;m nylon cell strainer&amp;nbsp;</td><td>Falcon</td><td>21008-949</td><td>To remove large chunk or blood clots, which can block the microfluidics access hole or the channel.</td></tr><tr><td>Peristaltic pump (Masterflex L/S Digital Drive)</td><td>Cole-Parmer</td><td>HV-07522-30</td><td>operation of microfluidics</td></tr><tr><td>BD LSR II flow cytometer</td><td>BD Bioscience</td><td>LSR II flow cytometer</td><td>Quantification of cell recovery ratio</td></tr><tr><td>Fluorescein isothiocyanate (FITC)-conjugated mouse anti-human CD66b monoclonal antibody</td><td>BD Bioscience</td><td>561927</td><td>Immunostaining of neutrophils for Flow cytometer analysis</td></tr><tr><td>Allophycocyanin (APC)-conjugated mouse anti-human CD45 monoclonal antibody</td><td>BD Bioscience</td><td>561864</td><td>Immunostaining of neutrophils for Flow cytometer analysis</td></tr><tr><td>Plate reader</td><td>Thermo Fisher scientific</td><td>Varioskan</td><td>Plate reader for neutrophil elastase assay, ex485/em525</td></tr><tr><td>Neutrophil elastase assay kit</td><td>Cayman Chemical</td><td>600610</td><td>Neutrophil functionality assessment</td></tr><tr><td>Fluoresbrite YG Microspheres 10.0&amp;micro;m</td><td>PolyScience, Inc.</td><td>18140-2</td><td>Fluorescent particles to express white blood cell trajectory in microfluidics</td></tr></tbody></table>

label-free neutrophil enrichment from patient-derived airway secretion using closed-loop inertial microfluidics

Airway secretions contain a large number of immune-related cells, e.g., neutrophils, macrophages, and lymphocytes, which can be used as a major resource to evaluate a variety of pulmonary diseases, both for research and clinical purposes. However, due to the heterogeneous and viscous nature of patient mucus, there is currently no reliable dissociation method that does not damage the host immune cells in the patient airway secretion. In this research, we introduce a sample preparation method that uses inertial microfluidics for the patient&#39;s immune assessment. Regardless of the heterogeneous fluidic properties of the clinical samples, the proposed method recovers more than 95% of neutrophils from airway secretion samples that are diluted 1,000-fold with milliliters of clean saline. By recirculating the concentrated output stream to the initial sample reservoir, a high concentration, recovery, and purity of the immune cells are provided; recirculation is considered a trade-off to the single-run syringe-based operation of inertial microfluidics. The closed-loop operation of spiral microfluidics provides leukocytes without physical or chemical disturbance, as demonstrated by the phorbol 12-myristate 13-acetate (PMA)-induced elastase release of sorted neutrophils.

We achieved transparent immune-cell suspensions with both DTT mucolysis and microfluidics dissociation (Figure 3A). Microfluidics dissociation collected 4.40 x 105 PMNs on average (2.1 x 104 to 5.60 x 105 PMNs, n = 6) from airway secretion samples diluted 1,000-fold (50 mL total volume) in 1 mL of clean suspension. Compared to the initial diluent, 94.0% PMNs (CD66b+/CD45+) were recovered in a small volume, consistently over 6 clinical samples. However, the DTT mucolysis method showed a 53.5% recovery on average with significant sample-to-sample variations (30.8&#160;- 96.0%) (Figure 3B).
Next, the release of elastase was examined with a commercial NE assay kit as a proof-of-concept. Without external stimulation, the sample prepared by the mucolytic method showed a higher NE level than the sample from the microfluidics method. PMA was added to the resulting suspensions from both the microfluidics and mucolytic dissociation methods to trigger the NE release. We tested each method with airway secretion samples from 6 different patients. Immune cells separated with microfluidics presented intact functionality, showing an increased amount of NE release by PMA stimulation. On the other hand, cells prepared by the mucolytic dissociation method showed an inconsistent increase of NE release (Figure 3C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57673/57673fig1.jpg" /> 
Figure 1: Spiral microfluidic device and experimental settings. Photograph images of (A) spiral microfluidics and (B) the device assembled with the fluidic adaptor. (C) Schematic diagram of closed-loop separation using spiral inertial microfluidics. By recirculating the particle/cell concentrated stream (IW outlet) back into the original sample reservoir, immune-related cells were concentrated in a small volume while the background fluid containing mucin aggregates and minor RBCs were continuously removed through the waste tube connected to the OW outlet. (D) Experimental setup and the fluorescent image of a 10 &#181;m particle, the trajectory (green) and the outlet of the spiral channel (upper right). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57673/57673fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57673/57673fig2.jpg" /> 
Figure 2: Schematic diagram of the airway secretion sample preparation using closed-loop inertial microfluidics. Clinically induced airway secretion dispersed mechanically in 1,000x volume of buffer solution. The diluent is concentrated by microfluidics, resulting in ~1 mL of cell suspension with the clear background. Adapted with permission from Ryu et al., Copyright (2017) American Chemical Society17. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57673/57673fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57673/57673fig3.jpg" /> 
Figure 3: Representative comparisons of the DTT mucolytic method to the microfluidic dissociation method. (A) Tracheal secretions sample (left) and microscopic image after the microfluidic dissociation. (B) Photographic image of the original and resulting suspensions. (C) Comparison of PMN recovery rates between the mucolytic and microfluidics methods. (D) Comparison of NE release of separated neutrophils by the microfluidics and DTT mucolytic methods before and after PMA stimulation. NE release was significantly triggered by PMA on the sample from microfluidic separation (p &#60;0.0001). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57673/57673fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Label-free Neutrophil Enrichment from Patient-derived Airway Secretion Using Closed-loop Inertial Microfluidics at JoVE.com

Label-free Neutrophil Enrichment from Patient-derived Airway Secretion Using Closed-loop Inertial Microfluidics

In this research, we demonstrate a label-free neutrophil separation method from clinical airway secretions using closed-loop operation of spiral inertial microfluidics. The proposed method would expand the clinical in vitro assays for various respiratory diseases.

Airway secretions contain a large number of immune-related cells, e.g., neutrophils, macrophages, and lymphocytes, which can be used as a major resource ...

label-free-neutrophil-enrichment-from-patient-derived-airway

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

6.3K Views.  Massachusetts Institute of Technology. In dit onderzoek tonen we een label-vrije neutrofiele scheiding methode van klinische airway afscheidingen met kringloopsysteem bewerking van spiraal inertial microfluidics. De voorgestelde methode zou het uitbreiden van de klinische in vitro tests voor verschillende aandoeningen van de luchtwegen.

Video: Label-free Neutrophil Enrichment from Patient-derived Airway Secretion Using Closed-loop Inertial Microfluidics

Microfluidic Platform for Measuring Neutrophil Chemotaxis from Unprocessed Whole Blood

Neutrophils play an essential role in protection against infections and their numbers in the blood are frequently measured in the clinic. Higher neutrophil counts in the blood are usually an indicator of ongoing infections, while low neutrophil counts are a warning sign for higher risks for infections. To accomplish their functions, neutrophils also have to be able to move effectively from the blood where they spend most of their life, into tissues, where infections occur. Consequently, any defects in the ability of neutrophils to migrate can increase the risks for infections, even when neutrophils are present in appropriate numbers in the blood. However, measuring neutrophil migration ability in the clinic is a challenging task, which is time consuming, requires large volume of blood, and expert knowledge. To address these limitations, we designed a robust microfluidic assays for neutrophil migration, which requires a single droplet of unprocessed blood, circumvents the need for neutrophil separation, and is easy to quantify on a simple microscope. In this assay, neutrophils migrate directly from the blood droplet, through small channels, towards the source of chemoattractant. To prevent the granular flow of red blood cells through the same channels, we implemented mechanical filters with right angle turns that selectively block the advance of red blood cells. We validated the assay by comparing neutrophil migration from blood droplets collected from finger prick and venous blood. We also compared these whole blood (WB) sources with neutrophil migration from samples of purified neutrophils and found consistent speed and directionality between the three sources. This microfluidic platform will enable the study of human neutrophil migration in the clinic and the research setting to help advance our understanding of neutrophil functions in health and disease.

This protocol details an assay designed to measure human neutrophil chemotaxis from one droplet of whole blood with robust reproducibility. This approach circumvents the need for neutrophil separation and requires only a few minutes of assay preparation time. The microfluidic chip enables the repeated measure of neutrophil chemotaxis over time in infants or small mammals, where sample volume is limited.

Neutrophils play an essential role in protection against infections and their numbers in the blood are frequently measured in the clinic. Higher ...

Bioengineering

A Method for Generating Pulmonary Neutrophilia Using Aerosolized Lipopolysaccharide

Acute lung injury (ALI) is a severe disease characterized by alveolar neutrophilia, with limited treatment options and high mortality. Experimental models of ALI are key in enhancing our understanding of disease pathogenesis. Lipopolysaccharide (LPS) derived from gram positive bacteria induces neutrophilic inflammation in the airways and lung parenchyma of mice. Efficient pulmonary delivery of compounds such as LPS is, however, difficult to achieve. In the approach described here, pulmonary delivery in mice is achieved by challenge to aerosolized Pseudomonas aeruginosa LPS. Dissolved LPS was aerosolized by a nebulizer connected to compressed air. Mice were exposed to a continuous flow of LPS aerosol in a Plexiglas box for 10 min, followed by 2 min conditioning after the aerosol was discontinued. Tracheal intubation and subsequent bronchoalveolar lavage, followed by formalin perfusion was next performed, which allows for characterization of the sterile pulmonary inflammation. Aerosolized LPS generates a pulmonary inflammation characterized by alveolar neutrophilia, detected in bronchoalveolar lavage and by histological assessment. This technique can be set up at a small cost with few appliances, and requires minimal training and expertise. The exposure system can thus be routinely performed at any laboratory, with the potential to enhance our understanding of lung pathology.

We describe a method for inducing neutrophilic pulmonary inflammation by challenge to aerosolized lipopolysaccharide by nebulization, to model acute lung injury. In addition, basic surgical techniques for lung isolation, tracheal intubation and bronchoalveolar lavage are also described.

Acute lung injury (ALI) is a severe disease characterized by alveolar neutrophilia, with limited treatment options and high mortality. Experimental models ...

Microfluidic Buffer Exchange for Interference-free Micro/Nanoparticle Cell Engineering

Engineering cells with active-ingredient-loaded micro/nanoparticles (NPs) is becoming an increasingly popular method to enhance native therapeutic properties, enable bio imaging and control cell phenotype. A critical yet inadequately addressed issue is the significant number of particles that remain unbound after cell labeling which cannot be readily removed by conventional centrifugation. This leads to an increase in bio imaging background noise and can impart transformative effects onto neighboring non-target cells. In this protocol, we present an inertial microfluidics-based buffer exchange strategy termed as Dean Flow Fractionation (DFF) to efficiently separate labeled cells from free NPs in a high throughput manner. The developed spiral microdevice facilitates continuous collection (&#62;90% cell recovery) of purified cells (THP-1 and MSCs) suspended in new buffer solution, while achieving &#62;95% depletion of unbound fluorescent dye or dye-loaded NPs (silica or PLGA). This single-step, size-based cell purification strategy enables high cell processing throughput (106 cells/min) and is highly useful for large-volume cell purification of micro/nanoparticle engineered cells to achieve interference-free clinical application.

This protocol describes the use of an inertial microfluidics-based buffer exchange strategy to purify micro/nanoparticle engineered cells with efficient depletion of unbound particles.

Engineering cells with active-ingredient-loaded micro/nanoparticles (NPs) is becoming an increasingly popular method to enhance native therapeutic ...

An All-on-chip Method for Rapid Neutrophil Chemotaxis Analysis Directly from a Drop of Blood

Neutrophil migration and chemotaxis are critical for our body's immune system. Microfluidic devices are increasingly used for investigating neutrophil migration and chemotaxis owing to their advantages in real-time visualization, precise control of chemical concentration gradient generation, and reduced reagent and sample consumption. Recently, a growing effort has been made by the microfluidic researchers toward developing integrated and easily operated microfluidic chemotaxis analysis systems, directly from whole blood. In this direction, the first all-on-chip method was developed for integrating the magnetic negative purification of neutrophils and the chemotaxis assay from small blood volume samples. This new method permits a rapid sample-to-result neutrophil chemotaxis test in 25 min. In this paper, we provide detailed construction, operation and data analysis method for this all-on-chip chemotaxis assay with a discussion on troubleshooting strategies, limitations and future directions. Representative results of the neutrophil chemotaxis assay testing a defined chemoattractant, N-Formyl-Met-Leu-Phe (fMLP), and sputum from a chronic obstructive pulmonary disease (COPD) patient, using this all-on-chip method are shown. This method is applicable to many cell migration-related investigations and clinical applications.

This article provides the detailed method of performing a rapid neutrophil chemotaxis assay by integrating the on-chip neutrophil isolation from whole blood and the chemotaxis test on a single microfluidic chip.

Neutrophil migration and chemotaxis are critical for our body's immune system. Microfluidic devices are increasingly used for investigating neutrophil ...

Noninvasive Sampling of Mucosal Lining Fluid for the Quantification of In Vivo Upper Airway Immune-mediator Levels

This protocol describes noninvasive sampling of undisturbed upper airway mucosal lining fluid. It also details the extraction procedure used prior to the analysis of immune mediators in fluid eluates for the study of the airway topical immune signature, without the need for stimulation procedures (often used by other techniques). The mucosal lining fluid is sampled on a strip of filter paper placed at the anterior part of the inferior turbinate and left for 2 min of absorption. Analytes are eluted from the filter papers, and the extracted protein-based eluates are analyzed by an electrochemiluminescence-based immunoassay, allowing for the high-sensitivity quantification of low- and high-level analytes in the same sample. We measured the in vivo levels of 20 preselected immune mediators related to specific immune signaling pathways in the upper airway mucosa, but the technique is not limited to that specific panel or sampling site. The technique was first implemented in 7-year-old children from the Copenhagen Prospective Studies on Asthma in Childhood2000 (COPSAC2000) cohort with allergic rhinitis. It was thereafter used in the longitudinal COPSAC2010 birth cohort, sampled at 1 month, 2 years, and 6 years of age and at instances of acute respiratory symptoms. We successfully obtained and analyzed samples from 620 (89%) of 700 1-month-old children; a few samples were below the assay detection limit (reported as the median (Inter-Quartile Range (IQR)). The number of samples below the detection limit (i.e.&#160;from 0 to the set point for the lower limit of detection) for each mediator was 29 (7.25 - 119.5). This technique enables the quantification of the in vivo airway mucosal immune profile from birth, can be applied longitudinally, and can be applied to studies on the effect of genetics and early-life environmental exposures, pathophysiology, endotyping, and monitoring of respiratory diseases, and development and evaluation of novel therapeutics.

Noninvasive Sampling of Mucosal Lining Fluid for the Quantification of In Vivo Upper Airway Immune-mediator Levels

This protocol describes a noninvasive technique for the sampling of undisturbed mucosal lining fluid from the upper airways. It can be used to perform the quantification of in vivo levels of protein mediators, such as cytokines and chemokines, in subjects of all ages.

This protocol describes noninvasive sampling of undisturbed upper airway mucosal lining fluid. It also details the extraction procedure used prior to the ...

Absorption of Nasal and Bronchial Fluids: Precision Sampling of the Human Respiratory Mucosa and Laboratory Processing of Samples

The methods of nasal absorption (NA) and bronchial absorption (BA) use synthetic absorptive matrices (SAM) to absorb the mucosal lining fluid (MLF) of the human respiratory tract. NA is a non-invasive technique which absorbs fluid from the inferior turbinate, and causes minimal discomfort. NA has yielded reproducible results with the ability to frequently repeat sampling of the upper airway. 
By comparison, alternative methods of sampling the respiratory mucosa, such as nasopharyngeal aspiration (NPA) and conventional swabbing, are more invasive and may result in greater data variability. Other methods have limitations, for instance, biopsies and bronchial procedures are invasive, sputum contains many dead and dying cells and requires liquefaction, exhaled breath condensate (EBC) contains water and saliva, and lavage samples are dilute and variable.
BA can be performed through the working channel of a bronchoscope in clinic. Sampling is well tolerated and can be conducted at multiple sites in the airway. BA results in MLF samples being less dilute than bronchoalveolar lavage (BAL) samples.
This article demonstrates the techniques of NA and BA, as well as the laboratory processing of the resulting samples, which can be tailored to the desired downstream biomarker being measured. These absorption techniques are useful alternatives to the conventional sampling techniques used in clinical respiratory research.

This manuscript describes the use of nasal and bronchial absorption techniques, specifically using synthetic absorptive matrices (SAM) to sample the mucosal lining fluid (MLF) of the upper and lower airway. These methods provide better standardization and tolerability than existing respiratory sampling techniques.

The methods of nasal absorption (NA) and bronchial absorption (BA) use synthetic absorptive matrices (SAM) to absorb the mucosal lining fluid (MLF) of the ...

Medicine

A Microphysiological System to Study Leukocyte-Endothelial Cell Interaction during Inflammation

Leukocyte-endothelial cell interactions play an important role in inflammatory diseases such as sepsis. During inflammation, excessive migration of activated leukocytes across the vascular endothelium into key organs can lead to organ failure. A physiologically relevant biomimetic microfluidic assay (bMFA) has been developed and validated using several experimental and computational techniques, which can reproduce the entire leukocyte rolling/adhesion/migration cascade to study leukocyte-endothelial cell interactions. Microvascular networks obtained from in vivo images in rodents were digitized using a Geographic Information System (GIS) approach and microfabricated with polydimethylsiloxane (PDMS) on a microscope slide. To study the effect of shear rate and vascular topology on leukocyte-endothelial cell interactions, a Computational Fluid Dynamics (CFD) model was developed to generate a corresponding map of shear rates and velocities throughout the network. The bMFA enables the quantification of leukocyte-endothelial cells interactions, including rolling velocity, number of adhered leukocytes in response to different shear rates, number of migrated leukocytes, endothelial cell permeability, adhesion molecule expression and other important variables. Furthermore, by using human-related samples, such as human endothelial cells and leukocytes, bMFA provides a tool for rapid screening of potential therapeutics to increase their clinical translatability.

In this protocol, a biomimetic microfluid assay, which can reproduce a physiologically relevant microvascular environment and reproduce the entire leukocyte adhesion/migration cascade, is employed to study leukocyte-endothelial cell interactions in inflammatory disease.

Leukocyte-endothelial cell interactions play an important role in inflammatory diseases such as sepsis. During inflammation, excessive migration of ...

Isolation and Enrichment of Human Lung Epithelial Progenitor Cells for Organoid Culture

Epithelial organoid models serve as valuable tools to study the basic biology of an organ system and for disease modeling. When grown as organoids, epithelial progenitor cells can self-renew and generate differentiating progeny that exhibit cellular functions similar to those of their in vivo counterparts. Herein we describe a step-by-step protocol to isolate region-specific progenitors from human lung and generate 3D organoid cultures as an experimental and validation tool. We define proximal and distal regions of the lung with the goal of isolating region-specific progenitor cells. We utilized a combination of enzymatic and mechanical dissociation to isolate total cells from the lung and trachea. Specific progenitor cells were then fractionated from the proximal or distal origin cells using fluorescence associated cell sorting (FACS) based on cell type-specific surface markers, such as NGFR&#160;for sorting basal cells and HTII-280 for sorting alveolar type II cells. Isolated basal or alveolar type II progenitors were used to generate 3D organoid cultures. Both distal and proximal progenitors formed organoids with a colony forming efficiency of 9-13% in distal region and 7-10% in proximal region when plated 5000 cell/well on day 30. Distal organoids maintained HTII-280+ alveolar type II cells in culture whereas proximal organoids differentiated into ciliated and secretory cells by day 30. These 3D organoid cultures can be used as an experimental tool for studying the cell biology of lung epithelium and epithelial mesenchymal interactions, as well as for the development and validation of therapeutic strategies targeting epithelial dysfunction in a disease.

This article provides a detailed methodology for tissue dissociation and cellular fractionation approaches allowing enrichment of viable epithelial cells from proximal and distal regions of the human lung. Herein these approaches are applied for the functional analysis of lung epithelial progenitor cells through the use of 3D organoids culture models.

Epithelial organoid models serve as valuable tools to study the basic biology of an organ system and for disease modeling. When grown as organoids, ...

Biology

Label-Free Neutrophil Separation from Tracheal Secretion Using Spiral Microfluidics Channel

Source: Ryu, H., et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/57673">Label-free Neutrophil Enrichment from Patient-derived Airway Secretion Using Closed-loop Inertial Microfluidics.</a>&#160;J. Vis. Exp.&#160;(2018).
This video demonstrates separating neutrophils from tracheal secretions using a microfluidic device with spiral channels. The device achieves efficient separation based on cell size by utilizing varying thicknesses and trapezoidal-shaped channels. The interplay between forces results in large neutrophils accumulating near the inner wall while erythrocytes and mucin aggregates remain near the outer wall, ensuring effective neutrophil separation.

Source: Ryu, H., et al., Label-free Neutrophil Enrichment from Patient-derived Airway Secretion Using Closed-loop Inertial Microfluidics.&#160;J. Vis. ...

JoVE Encyclopedia of Experiments

Immunology

Immune Systems and Components

Assays and Functional Profiling

Peritoneal Low-density Neutrophil Isolation: A Technique to Obtain Low-density Neutrophils from Peritoneal Lavage Fluid Using Magnetic Activated Cell Sorting

To acquire the neutrophils, first, infuse 1 liter of normal sterile saline immediately before wound closure directly into the abdominal cavity of a patient who has just undergone abdominal surgery due to gastrointestinal malignancy, and lavage the abdominal cavity extensively for at least 1 minute. Then, stir the saline within the cavity and recover 200 milliliters of lavage fluid with four 50-milliliter syringes.
In the lab, transfer the peritoneal lavage fluid through a 100-micrometer nylon filter into individual 50-milliliter tubes for centrifugation. Resuspend the pellets in 5 milliliters of PBS supplemented with 0.02% EDTA and carefully overlay each cell suspension onto 3 milliliters of density gradient solution. After density gradient separation, harvest 2 to 3 milliliters of the intermediate layer from each tube and wash the peritoneal fluid samples in 10 milliliters of fresh PBS plus EDTA per tube. Pour the pellets in 10 milliliters of fresh PBS plus EDTA for an additional wash and dilute the cells to a 1 times 10 to the seventh cells per 60 micrometers of magnetic activated cell sorting, or MACS, buffer concentration.
Block any non-specific binding with 20 microliters of Fc block for 10 minutes at 4 degrees Celsius, followed by the addition of 20 microliters of anti-CD66b magnetic beads. After 10 minutes at 4 degrees Celsius, wash and resuspend the cells in 500 microliters of MACS buffer and add the cells to an appropriately sized magnetic column within the magnetic field of a suitable magnetic separator. When all of the CD66b negative cells have run through the column, add 15 milliliters of fresh MACS buffer to the column reservoir and transfer the column from the magnetic separator into a new conical tube. Then, immediately plunge the column to flush the magnetically labeled CD66b positive into the tube.

Label-free Neutrophil Enrichment from Patient-derived Airway Secretion Using Closed-loop Inertial Microfluidics

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