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...

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&#8221; x 3&#8221; 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 &#38; preparation</td>
		</tr>
		<tr>
			<td>Phosphate-Buffered Saline, 1X Without Calcium and Magnesium</td>
			<td>Corning</td>
			<td>45000-446&#160;</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&#8482; 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&#160; Blunt Fill Needle, with BD Luer-Lok&#160; 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&#181;m nylon cell strainer&#160;</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&#181;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...

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

Research

JoVE Journal

Immunology and Infection

6.3K Views.  Massachusetts Institute of Technology. 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.

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

Peptide:MHC Tetramer-based Enrichment of Epitope-specific T cells

A basic necessity for researchers studying adaptive immunity with in vivo experimental models is an ability to identify T cells based on their T cell antigen receptor (TCR) specificity. Many indirect methods are available in which a bulk population of T cells is stimulated in vitro with a specific antigen and epitope-specific T cells are identified through the measurement of a functional response such as proliferation, cytokine production, or expression of activation markers1. However, these methods only identify epitope-specific T cells exhibiting one of many possible functions, and they are not sensitive enough to detect epitope-specific T cells at naive precursor frequencies. A popular alternative is the TCR transgenic adoptive transfer model, in which monoclonal T cells from a TCR transgenic mouse are seeded into histocompatible hosts to create a large precursor population of epitope-specific T cells that can be easily tracked with the use of a congenic marker antibody2,3. While powerful, this method suffers from experimental artifacts associated with the unphysiological frequency of T cells with specificity for a single epitope4,5. Moreover, this system cannot be used to investigate the functional heterogeneity of epitope-specific T cell clones within a polyclonal population.
	The ideal way to study adaptive immunity should involve the direct detection of epitope-specific T cells from the endogenous T cell repertoire using a method that distinguishes TCR specificity solely by its binding to cognate peptide:MHC (pMHC) complexes. The use of pMHC tetramers and flow cytometry accomplishes this6, but is limited to the detection of high frequency populations of epitope-specific T cells only found following antigen-induced clonal expansion. In this protocol, we describe a method that coordinates the use of pMHC tetramers and magnetic cell enrichment technology to enable detection of extremely low frequency epitope-specific T cells from mouse lymphoid tissues3,7. With this technique, one can comprehensively track entire epitope-specific populations of endogenous T cells in mice at all stages of the immune response.

This protocol describes the use of peptide:MHC tetramers and magnetic microbeads to isolate low frequency populations of epitope-specific T cells and analyze them by flow cytometry. This method enables the direct study of endogenous T cell populations of interest from in vivo experimental systems.

A  basic necessity for researchers studying adaptive immunity with in vivo experimental models is an  ability to identify T cells based on their T cell ...

Long Term Chronic Pseudomonas aeruginosa Airway Infection in Mice

A mouse model of chronic airway infection is a key asset in cystic fibrosis (CF) research, although there are a number of concerns regarding the model itself. Early phases of inflammation and infection have been widely studied by using the Pseudomonas aeruginosa agar-beads mouse model, while only few reports have focused on the long-term chronic infection in vivo. The main challenge for long term chronic infection remains the low bacterial burden by P. aeruginosa and the low percentage of infected mice weeks after challenge, indicating that bacterial cells are progressively cleared by the host.
This paper presents a method for obtaining efficient long-term chronic infection in mice. This method is based on the embedding of the P. aeruginosa clinical strains in the agar-beads in vitro, followed by intratracheal instillation in C57Bl/6NCrl mice. Bilateral lung infection is associated with several measurable read-outs including weight loss, mortality, chronic infection, and inflammatory response. The P. aeruginosa RP73 clinical strain was preferred over the PAO1 reference laboratory strain since it resulted in a comparatively lower mortality, more severe lesions, and higher chronic infection. P. aeruginosa colonization may persist in the lung for over three months. Murine lung pathology resembles that of CF patients with advanced chronic pulmonary disease.
This murine model most closely mimics the course of the human disease and can be used both for studies on the pathogenesis and for the evaluation of novel therapies.

Long Term Chronic Pseudomonas aeruginosa Airway Infection in Mice

We describe the agar-beads method to establish persistent long-term chronic Pseudomonas aeruginosa airway infection in the mouse model.

A mouse model of chronic airway infection is a key asset in cystic fibrosis (CF) research, although there are a number of concerns regarding the model ...

Human Neutrophil Flow Chamber Adhesion Assay

Neutrophil firm adhesion to endothelial cells plays a critical role in inflammation in both health and disease. The process of neutrophil firm adhesion involves many different adhesion molecules including members of the &beta;2 integrin family and their counter-receptors of the ICAM family. Recently, naturally occurring genetic variants in both &beta;2 integrins and ICAMs are reported to be associated with autoimmune disease. Thus, the quantitative adhesive capacity of neutrophils from individuals with varying allelic forms of these adhesion molecules is important to study in relation to mechanisms underlying development of autoimmunity. Adhesion studies in flow chamber systems can create an environment with fluid shear stress similar to that observed in the blood vessel environment in vivo. Here, we present a method using a flow chamber assay system to study the quantitative adhesive properties of human peripheral blood neutrophils to human umbilical vein endothelial cell (HUVEC) and to purified ligand substrates. With this method, the neutrophil adhesive capacities from donors with different allelic variants in adhesion receptors can be assessed and compared. This method can also be modified to assess adhesion of other primary cell types or cell lines.

A method of quantitating neutrophil adhesion is reported. This method creates a dynamic flow environment similar to that encountered in a blood vessel. It allows the investigation of neutrophil adhesion to either purified adhesion molecules (ligand) or endothelial cell substrate (HUVEC) in a context similar to the in vivo environment with sheer stress.

Neutrophil firm adhesion to endothelial cells plays a critical role in inflammation in both health and disease. The process of neutrophil firm adhesion ...

Detection of Human Leukocyte Antigen Biomarkers in Breast Cancer Utilizing Label-free Biosensor Technology

According to the American Cancer Society, more than 200,000 women will be diagnosed with invasive breast cancer each year and approximately 40,000 will die from the disease. The human leukocyte antigen (HLA) class I samples peptides derived from proteasomal degradation of cellular proteins and presents these fragments on the cell surface for interrogation by circulating cytotoxic T lymphocytes (CTL). Generation of T-cell receptor mimic (TCRm) monoclonal antibodies (mAbs) which recognize breast cancer specific peptide/HLA-A*02:01 complexes such as those derived from macrophage migration inhibitory factor (MIF19-27) and NY-ESO-1157-165 enable detection and destruction of breast cancer cells in the absence of an effective anti-tumor CTL response. Intact class I HLA/peptide complexes are shed by breast cancer cells and represent potentially relevant cancer biomarkers. In this work, a breakthrough biomarker screening system for cancer diagnostics incorporating T-cell receptor mimic monoclonal antibodies combined with a novel, label-free biosensor utilizing guided-mode resonance (GMR) sensor technology is presented. Detection of shed MIF/HLA-A*02:01 complexes in MDA-MB-231 cell supernatants, spiked human serum, and patient plasma is demonstrated. The impact of this work could revolutionize personalized medicine through development of companion disease diagnostics for targeted immunotherapies.

Intact class I HLA/peptide complexes are shed by cancer cells, representing a potential relevant cancer biomarker. Utilizing label-free sensor technology and T-cell receptor mimicking monoclonal antibodies, detection of shed MIF/HLA-A*02:01 complexes in MDA-MB-231 cell supernatants, spiked human serum, and patient plasma is demonstrated, enabling development of a novel cancer diagnostic platform.

According to the American Cancer Society, more than 200,000 women will be diagnosed with invasive breast cancer each year and approximately 40,000 will ...

Automated Cell Enrichment of Cytomegalovirus-specific T cells for Clinical Applications using the Cytokine-capture System

The adoptive transfer of pathogen-specific T cells can be used to prevent and treat opportunistic infections such as cytomegalovirus (CMV) infection occurring after allogeneic hematopoietic stem-cell transplantation. Viral-specific T cells from allogeneic donors, including third party donors, can be propagated ex vivo in compliance with current good manufacturing practice (cGMP), employing repeated rounds of antigen-driven stimulation to selectively propagate desired T cells. The identification and isolation of antigen-specific T cells can also be undertaken based upon the cytokine capture system of T cells that have been activated to secrete gamma-interferon (IFN-&#947;). However, widespread human application of the cytokine capture system (CCS) to help restore immunity has been limited as the production process is time-consuming and requires a skilled operator. The development of a second-generation cell enrichment device such as CliniMACS Prodigy now enables investigators to generate viral-specific T cells using an automated, less labor-intensive system. This device separates magnetically labeled cells from unlabeled cells using magnetic activated cell sorting technology to generate clinical-grade products, is engineered as a closed system and can be accessed and operated on the benchtop. We demonstrate the operation of this new automated cell enrichment device to manufacture CMV pp65-specific T cells obtained from a steady-state apheresis product obtained from a CMV seropositive donor. These isolated T cells can then be directly infused into a patient under institutional and federal regulatory supervision. All the bio-processing steps including removal of red blood cells, stimulation of T cells, separation of antigen-specific T cells, purification, and washing are fully automated. Devices such as this raise the possibility that T cells for human application can be manufactured outside of dedicated good manufacturing practice (GMP) facilities and instead be produced in blood banking facilities where staff can supervise automated protocols to produce multiple products.

The goal of this protocol is to manufacture pathogen-specific clinical-grade T cells using a bench-top, automated, second generation cell enrichment device that incorporates a closed cytokine capture system and does not require dedicated staff or use of a GMP facility. The cytomegalovirus pp65-specific-T cells generated can be directly administered to patients.

The adoptive transfer of pathogen-specific T cells can be used to prevent and treat opportunistic infections such as cytomegalovirus (CMV) infection ...

Intravital Imaging of Neutrophil Priming Using IL-1&#946; Promoter-driven DsRed Reporter Mice

Neutrophils are the most abundant leukocytes in human blood circulation and are quickly recruited to inflammatory sites. Priming is a critical event that enhances the phagocytic functionality of neutrophils. Although extensive studies have unveiled the existence and importance of neutrophil priming during infection and injury, means of visualizing this process in vivo have been unavailable. The protocol provided enables monitoring of the dynamic process of neutrophil priming in living animals by combining three methodologies: 1) DsRed reporter signal &#8212; used as a measure of priming 2) in vivo neutrophil labeling &#8212; achieved by injection of fluorescence-conjugated anti-lymphocyte antigen 6G (Ly6G) monoclonal antibody (mAb) and 3) intravital confocal imaging. Several critical steps are involved in this protocol: oxazolone-induced mouse ear skin inflammation, appropriate sedation of animals, repeated injections of anti-Ly6G mAb, and prevention of focus drift during imaging. Although a few limitations have been observed, such as the limit of continuous imaging time (~ 8 hr) in one mouse and the leakage of fluorescein isothiocyanate-dextran from blood vessels in the inflammatory state, this protocol provides a fundamental framework for intravital imaging of primed neutrophil behavior and function, which can easily be expanded to examination of other immune cells in mouse inflammation models.

This current protocol employs fluorescent reporters, in vivo labeling, and intravital imaging techniques to enable monitoring of the dynamic process of neutrophil priming in living animals.

Neutrophils are the most abundant leukocytes in human blood circulation and are quickly recruited to inflammatory sites. Priming is a critical event that ...

Imaging the Neutrophil Phagosome and Cytoplasm Using a Ratiometric pH Indicator

Neutrophils are crucial to host innate defense and, consequently, constitute an important area of medical research. The phagosome, the intracellular compartment where the killing and digestion of engulfed particles take place, is the main arena for neutrophil pathogen killing that requires tight regulation. Phagosomal pH is one aspect that is carefully controlled, in turn regulating antimicrobial protease activity. Many fluorescent pH-sensitive dyes have been used to visualize the phagosomal environment. S-1 has several advantages over other pH-sensitive dyes, including its dual emission spectra, its resistance to photo-bleaching, and its high pKa. Using this method, we have demonstrated that the neutrophil phagosome is unusually alkaline in comparison to other phagocytes. By using different biochemical conjugations of the dye, the phagosome can be delineated from the cytoplasm so that changes in the size and shape of the phagosome can be assessed. This allows for further monitoring of ionic movement.

This manuscript describes a simple method to measure the phagosomal pH and area as well as the cytoplasmic pH of human and mouse neutrophils using the ratiometric indicator seminaphthorhodafluor (SNARF)-1, or S-1. This is achieved using live-cell confocal fluorescence microscopy and image analysis.

Neutrophils are crucial to host innate defense and, consequently, constitute an important area of medical research. The phagosome, the intracellular ...

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 ...

Label-Free Identification of Lymphocyte Subtypes Using Three-Dimensional Quantitative Phase Imaging and Machine Learning

We describe here a protocol for the label-free identification of lymphocyte subtypes using quantitative phase imaging and machine learning. Identification of lymphocyte subtypes is important for the study of immunology as well as diagnosis and treatment of various diseases. Currently, standard methods for classifying lymphocyte types rely on labeling specific membrane proteins via antigen-antibody reactions. However, these labeling techniques carry the potential risks of altering cellular functions. The protocol described here overcomes these challenges by exploiting intrinsic optical contrasts measured by 3D quantitative phase imaging and a machine learning algorithm. Measurement of 3D refractive index (RI) tomograms of lymphocytes provides quantitative information about 3D morphology and phenotypes of individual cells. The biophysical parameters extracted from the measured 3D RI tomograms are then quantitatively analyzed with a machine learning algorithm, enabling label-free identification of lymphocyte types at a single-cell level. We measure the 3D RI tomograms of B, CD4+ T, and CD8+ T lymphocytes and identified their cell types with over 80% accuracy. In this protocol, we describe the detailed steps for lymphocyte isolation, 3D quantitative phase imaging, and machine learning for identifying lymphocyte types.

We describe a protocol for the label-free identification of lymphocyte subtypes using quantitative phase imaging and a machine learning algorithm. Measurements of 3D refractive index tomograms of lymphocytes present 3D morphological and biochemical information for individual cells, which is then analyzed with a machine-learning algorithm for identification of cell types.

We describe here a protocol for the label-free identification of lymphocyte subtypes using quantitative phase imaging and machine learning. Identification ...

Automated Image-Based Quantification of Neutrophil Extracellular Traps Using NETQUANT

Neutrophil extracellular traps (NETs) are web-like antimicrobial structures consisting of DNA and granule derived antimicrobial proteins. Immunofluorescence microscopy and image-based quantification methods remain important tools to quantitate neutrophil extracellular trap formation. However, there are key limitations to the immunofluorescence-based methods that are currently available for quantifying NETs. Manual methods of image-based NET quantification are often subjective, prone to error and tedious for users, especially non-experienced users. Also, presently available software options for quantification are either semi-automatic or require training prior to operation. Here, we demonstrate the implementation of an automated immunofluorescence-based image quantification method to evaluate NET formation called NETQUANT. The software is easy to use and has a user-friendly graphical user interface (GUI). It considers biologically relevant parameters such as an increase in the surface area and DNA:NET marker protein ratio, and nuclear deformation to define NET formation. Furthermore, this tool is built as a freely available app, and allows for single-cell resolution quantification and analysis.

Here, we present a protocol for generating neutrophil extracellular traps (NETs) and operating NETQUANT, a fully automatic software option for quantification of NETs in immunofluorescence images.