Analysis of neutrophil responses to biological exposures

Abstract

Neutrophils are an important part of the innate immune system and among the first cells to respond to infections and inflammation. Responses include chemotaxis towards stimuli, extravasation from the vasculature, and antimicrobial actions such as phagocytosis, granule release, reactive oxygen species (ROS) production, and neutrophil extracellular trap (NET) formation (NETosis). Studying how neutrophils respond to a variety of stimuli, from biomaterial interactions to microbial insults, is therefore an essential undertaking to fully comprehend the immune response. While there are some immortalized cell lines available that recapitulate many neutrophil responses, to fully understand the complete range of neutrophil phenotypes, ex vivo or in vivo studies are required. We describe here two protocols for neutrophil isolation for further ex vivo study: recovery of neutrophils from human peripheral blood and isolation of neutrophils from the oral cavity. We also discuss an in vivo model of general inflammation with the murine air pouch that can be used to assess numerous parameters of neutrophil and immune activation, including neutrophil recruitment and biological activity. In these protocols, the cells are isolated so as to allow for a high degree of experimental control. The protocols are also relatively straightforward and can be successfully used by labs with no prior primary cell experience.

INTRODUCTION:

Neutrophils, or polymorphonuclear leukocytes (PMNs), are the most abundant circulating leukocyte in the blood, following their production in bone marrow and subsequent trafficking and recruitment to tissues as part of protective immune surveillance or in response to infection or other inflammatory signals (Furze & Rankin, 2008). Neutrophils are the most promient immune cell transiting through the oral cavity and are prevalent in biological fluids such as saliva. The oral cavity provides an easily accesible resource and valuable measure of oral health, but also overall immune cell function and the effect of genetic deficiency, biolgical exposures and chronic conditions in the body. However, they have a relatively short lifespan and are consequently generated at a high rate (approximately 2×10¹¹ per day) in a healthy adult human (Borregaard, 2010). Neutrophils are highly dynamic cells; they perform chemotaxis towards stimuli and readily extravasate from the vasculature to sites of stimulation or infection. Once recruited, neutrophils undertake a multitude of antimicrobial actions including phagocytosis, granule release, reactive oxygen species (ROS) production, and NETosis (Lee & Dominguez, 2010; Roberts & Hallett, 2019) all of which are important for immune function, but which can cause tissue damage during extended or chronic activation. Often, the measurement of many of these neutrophil functions is more readily performed and quantified using ex vivo isolated cells rather than using immortalized cell lines that often do not recapitulate the full function of neutrophils (Blanter et al., 2021; Verdon et al., 2021). This is especially true for examining neutrophil phenotypic plasticity in responses to stimuli, which vary with local signals released during acute or chronic inflammatory conditions, injury, infection, cancer, and autoimmunity (Deniset & Kubes, 2018). We describe here two protocols for neutrophil isolation from readily accessible pools of neutrophils for further ex vivo study: recovery of neutrophils from human peripheral blood and enrichemnt of neutrophils from the oral cavity. With these two protocols, cells are isolated so as to allow for a high degree of experimental control to allow either direct neutrophil phenotypic and functional analysis in vivo or examine neutrophil responses to various biological exposures in vitro. We also describe an in vivo model of general inflammation using a murine air pouch that can be used to readily assess numerous parameters of neutrophil and immune activation, including neutrophil recruitment to tissue.

Considerations Prior to Isolation of Cells:

As isolated neutrophils have a brief lifespan in culture with high sensitivity to physical stress and lytic potential, it is essential to begin experiments promptly following isolation from blood or the oral cavity. Neutrophils are sensitive to any contamination of media and labware, therefore, all reagents and materials that come into contact with blood, salivary rinses or isolated cells should be clean, sterile and endotoxin-free. Disposable single-use plasticware should be used when possible, to maintain sterility and prevent erroneous activation. Unless otherwise indicated, maintain blood and isolated cells at physiological pH, room temperature and in buffers lacking divalent cations such as magnesium and calcium. Calcium and magnesium should be present at physiological levels during functional assays, but avoided during isolation to prevent activation. Neutrophils can be easily primed, thus avoid excessive shaking, vibration or vortexing of cells during preparation as this can enhance the intensity of responses and skew data, and strong activation can result in the formation of macroscopic clumps of cells.

Caution should be used when working with human blood and fluids, and best biosafety practices should be used to limit the risk of exposure to blood-borne pathogens. Personal protective gear (gloves, lab coats, eye protection), biosafety cabinets, and aerosol control devices should be used as required. Blood and oral rinses should be obtained under IRB approved protocols with documented informed consent. Appropriate IACUC approval protocols and animal handling procedures must be followed for all animal experiments.

BASIC PROTOCOL 1

Basic protocol title:

Neutrophil isolation from human blood

Introductory paragraph:

This protocol isolates human peripheral blood neutrophils via a density gradient using Accurate Chemical 1-Step Polymorphs and includes steps for washing cells and removing residual red blood cells (RBCs) via hypotonic lysis. When performed carefully, this method results in minimal neutrophil activation and the isolated cells are acceptable for most assays. It is important to obtain signed consent forms from each blood donor and to ensure all appropriate human subject guidelines are followed.

Materials:

1-Step Polymorphs (Accurate Chemical cat. no. AN221725)

HBSS without calcium and magnesium (Corning, cat. no. 21S-022-CV)

RBC Lysis Buffer (Invitrogen, cat. no. 00-4333-57)

HBSS with calcium and magnesium (Corning, cat. no. 21-023-CM)

Trypan Blue (VWR, cat. no. 97063-702)

Blood Collection Set (BD, cat. no.368656)

-Includes 21-guage winged needle, collection tubing and pre-attached tube holder

Tourniquet (Heathrow Scientific, cat. no. HEA-120108)

Vacuette blood collection tubes with EDTA (Greiner bio-one, cat. no. 456038)

15 ml conical tubes (Greiner bio-one, cat. no. 188261)

Transfer pipets, sterile (Fisherbrand, cat. no. 13-711-20)

Centrifuge (Eppendorf, Centrifuge 5804/5804 R or equivalent)

50 ml conical tubes (Greiner bio-one, cat. no. 227261)

Microcentrifuge tubes (Thermo Fisher, cat. no. 3451)

Single channel pipettes (2 μl through 1 ml)

5 ml Serological Pipette (Corning, cat. no. 4487)

10 ml Serological Pipette (Corning, cat. no. 4488)

Hemacytometer (Fisher Scientific, cat. no. 02-671-10)

Brightfield Microscope

Biological safety cabinet

Protocol steps with step annotations:

1. Using a blood collection set and tourniquet, collect 36 ml of peripheral blood total (non-fasting venous sample) into 6 vacuette tubes with EDTA (~ 6 mL per tube).

   Keep blood and isolation media at room temperature.

   EDTA or another anti-coagulant is necessary to prevent clotting, but heparin is not recommended for use with Polymorphprep.

2. Invert tubes well to mix blood well with anti-coagulant.

3. Fill six 15 ml conical tubes with 5 ml of 1-step Polymorphs.

   Polymorphprep solution should be stored at room-temperature and protected from light.

4. Using a transfer pipet, gently layer 5 ml of blood on top of the 5 ml of 1-step Polymorphs.

   Tilt tube containing 1-step Polymorphs at a roughly 45-degree angle and drip the blood down the side of the tube to lay atop of the Polymorphs. Blood should settle on top of the Polymorphs, and not break through the surface (Figure 1A).

5. Centrifuge for 35 minutes, 730 x g at room temperature with no brake and minimum acceleration.

   After centrifugation, the tube will appear stratified. The yellow-colored plasma will be at the top of the tube, a dense layer of debris and red blood cells will be at the bottom. In the clear middle section, there will be two white bands. The upper band is composed mononuclear cells (monocytes, lymphocytes), while the lower band is mature neutrophils (Figure 1B).

6. Use a transfer pipet to remove and discard the upper layers and top band, until there is a minimal amount of fluid above the neutrophil band.

   Neutrophil layers can be removed directly by bypassing the plasma and mononuclear cell layer with a transfer pipet.

7. Use a new transfer pipet to collect lower band and transfer to a new 15 ml conical tube. Pool together neutrophil bands from all gradient tubes into 1 or 2 (as required) 50 ml conical tubes.

   Neutrophil layers can alternatively be removed directly by bypassing the plasma and mononuclear cell layer with a transfer pipet.

8. Wash cells: fill tube with 20 ml HBSS (without Ca or Mg) and centrifuge 5 minutes, 400 x g at room temperature, acceleration & deceleration 4.

   A red tinged pellet will form at the bottom of the conical tube, consisting of neutrophils and some remaining RBCs (Figure 1C).

9. Remove supernatant and resuspend pellet in 10 ml RBC Lysis Buffer to lyse remaining red blood cells for 7 minutes at room temp. Immediately follow with the next step to prevent neutrophil lysis.

10. Wash cells: add 10 ml HBSS (without Ca or Mg) and centrifuge 5 minutes, 400 x g at room temperature, acceleration & deceleration 4 (Figure 1D).

11. Resuspend cells in 1 ml HBSS (with Ca and Mg) or other medium as required for further experiments.

12. Quantitate isolated neutrophils using a hemacytometer, automated cell counter or differential staining.

13. To quantitate neutrophils and assess viability using a hemacytometer: Dilute 1-2 μl of cells 1:100 with HBSS in a microcentrifuge tube, mix 10 μl diluted cells in a new microcentrifuge tube and mix with 10 μl trypan blue.

14. Place 10 μl into a hemocytometer and count cells under the microscope. Calculate how many total cells have been isolated and how many will be needed for downstream experiments and analysis.

Figure 1. Separation of blood during neutrophil isolation.

A) Peripheral blood is delicately layered over 1-step Polymorph solution prior to centrifugation. B) Stratified layers of plasma, mononuclear, neutrophils and blood cells after centrifugation. C) Collected neutrophils form a red pellet after centrifugation indicating the presence of red blood cells (RBCs). D) Collected neutrophils after centrifugation following RBC lysis.

BASIC PROTOCOL 2:

Basic protocol title:

Neutrophil Isolation from the oral cavity

Introductory paragraph:

This protocol describes a method for isolating oral neutrophils with minimal contamination of oral debris and epithelial cells. This method uses oral rinses passed through a series of size exclusion filters to obtain an isolated population of neutrophils. Due to the proteolytic properties and the fragile nature of oral neutrophils, it is necessary to perform the all steps on ice or at 4C with minimal delay. Institutional review board approval and consent forms are required from each donor, as are appropriate infection control procedures.

Materials:

0.9% sodium chloride solution for irrigation (VWR, cat. no. 68100-026 or equivalent)

Dulbecco’s phosphate-buffered saline without Ca or Mg (Corning, cat. no. 21-030-CV)

50 ml Conical tubes (Greiner bio-one, cat. no. 227261)

15 ml conical tubes (Greiner bio-one, cat. no. 188261)

Timer

Ice and ice bucket

40 μm cell strainer (Griener bio-one, cat. no. 542040)

20 μm nylon filter (Millipore, cat. no. NY2002500)

11 μm nylon filter (Millipore, cat. no. NY1102500)

Swinnex 25 filter holders (Millipore, cat. no. SX0002500)

20 ml syringes, sterile (Becton Dickinson, cat. no. 302831)

Centrifuge (Eppendorf, Centrifuge 5804/5804 R or equivalent)

Protocol steps with step annotations:

1. Collect oral rinses prior to oral instrumentation and exam to avoid initiating bleeding.

2. Have the donor rinse their oral cavity with 5 ml isotonic 0.9% sodium chloride solution for 30 seconds and expectorate into a sterile 50 ml tube, on ice.

3. Repeat rinse step 6 times with 3-minute intervals between each rinse sample for a total collection volume of 30 ml.

   You may need to adjust the number of rinses collected to isolate the desired number of cells based on the number of neutrophils populating the oral cavity (for example: a subject with good oral health = 20 to 30 times, a subject with periodontal disease = 6 to 10 times).

4. Mount 20 and 10 μm filters in Swinnex 25 filter holders. Connect each filter/ filter holder to a 20 ml syringe and remove the plunger. Place the filter holder/ syringe set-up over a 15 ml conical tube on ice.

5. Filter pooled rinse samples sequentially through a 40 μm cell strainer, then the 20 μm and 11 μm nylon mesh filters. Use gravity flow only to pass the samples through the filters.

   Do not push the sample through the syringe using the plunger. Keep samples on ice during filtration process.

6. Centrifuge collected effluent at 500 x g for 10 minutes, 4°C

7. Remove supernatant and resuspend cell pellet in 1 ml cold phosphate buffered saline.

8. Count the number of neutrophils using a hemocytometer (see step 14 in Neutrophil Isolation from Blood protocol above) or automated cell counter. Purity of cell population can be assessed by differential staining of cytopsin preparations (i.e., DiffQuick or Geimsa stain) or flow cytometry of appropriate surface markers and viability assessed by trypan blue exclusion assay.

BASIC PROTOCOL 3:

Basic protocol title:

Murine air pouch model of general inflammation

Introductory paragraph:

The murine air pouch model, described here, is a simple in vivo model that allows for study of induction and/or resolution of the inflammatory response, and responses to test compounds or biological exposures. Cellular processes can be measured by quantifying the volume and cellular composition of exudate, levels of secreted inflammatory mediators or other molecules, and biological or molecular responses of recruited cells. Air pouch-lining tissue can also be collected for histological or downstream measures such as protein and gene expression analysis. This method provides an easily-accessible subcutaneous environment for experimental manipulation and investigation. Multiple subcutaneous injections of sterile air in the dorsal intrascapular region over time results in the formation of a cell-lined membrane around the air-filled space, separating the dermis and epidermis from the subcutaneous tissue (Figure 2).

Figure 2. Schematic of Murine Air Pouch Model Procedure.

Representation of steps of the air pouch model of inflammation: A) preparation, B) formation and maintenance of the air pouch, C) treatment and D) sample collection.

STRATEGIC PLANNING:

Experimental plans should be made well in advance of performing the actual procedures. Experimenters should carefully plan how many conditions to be tested, how many time points the experiment should include, and how many mice per condition to achieve biological significance. All conditions should have a minimum of 3 mice, with 6 approaching a practical maximum for most experiments, though exact numbers should be determined by performing an appropriate statistical power analysis. The study design should include a vehicle control group, a positive control group (a known inflammatory stimulus), and exposure test groups for different substances, doses, and/or time points as appropriate.

To ensure the timely processing of mice, there should be a minimum of two experimenters for every stage of this protocol, though three is ideal. Experimenters should plan endpoint analyses and biological samples to be collected, as this determines whether the mice need to be shaved (optional step) and the number of collection tubes required. To simplify workflow and maintain efficiency, ensure you have all required paperwork available for surgical procedures and animal monitoring per IACUC and institutional guidelines.

Materials:

Mice (weighing ~20 to 25 grams. C57BL/6, BALB/c or strain of choice)

Anesthetic (e.g., 5% (v/v) isoflurane for inhalation; VWR cat. no. AAAL17315-14)

Ethanol, 70% (v/v) (Decon Labs, cat. no. 2701)

Dulbecco’s phosphate-buffered saline without Ca or Mg (Corning, cat. no. 21-030-CV)

Low density carboxymethlycelluose (CMC) (Sigma cat. no. C5678) (if required)

Positive control stimulus of choice (if required). Examples include:

• N-formyl-methionine-leucyl-phenlalanine (fMLP, fMLF). Sigma cat. no. F3506)

• E. coli 055:B5 lipopolysaccharide (LPS. Sigma cat. no. L6529)

Biological stimuli to be tested

Liquid nitrogen

1- , 3- and 5-ml syringes, sterile (Becton Dickinson, cat. no. 309659, 309656, and 309646)

Covered plastic storage box (plastic Rubbermaid storage container or similar item)

Biological safety cabinet

18- and 26-gauge needles, sterile (Becton Dickinson, BD PrecisionGlide^™ needles, cat. no. 305195 and 302887)

Anesthesia machine

Anesthetic chamber

(Optional) Animal electric shaver

Cotton balls

Sharpie

Scale

50 ml Conical tubes (Greiner bio-one, cat. no. 227261)

15 ml conical tubes (Greiner bio-one, cat. no. 188261)

2 ml and 1.5 ml Microcentrifuge tubes (VWR cat. no. 20901-505, Avant cat. no. SRS-2925)

Scissors and forceps, sterile

Ice and ice bucket

Transfer pipets, sterile (Fisherbrand, cat. no. 13-711-20)

CO₂ Chamber

Cell counter or hemacytometer

Benchtop microcentrifuge

Protocol steps with step annotations:

Preparation (Figure 2A)

1. Follow best practices for animal use as per institutional guidelines. Prior to beginning the experiment, acclimate the mice for one week under standard lighting and temperature conditions, with appropriate food and water available ad lib.

2. Prepare syringes filled with sterile air.

   1. Spray 5 ml syringes and needles with 70% ethanol and move them to a tissue culture hood.
   2. Turn on the UV light for 10–15 minutes.
   3. Fill syringes with 3 ml of air from inside the tissue culture hood and cap with 26-gauge needles.
   4. Store filled syringes in a covered storage box until needed.

   Prepare 2n+5 syringes, where n is your number of mice to ensure you have enough prepared needles for both air pouch inflation days with extra syringes as needed.

3. Prepare all required paperwork for animal surgery and monitoring procedures per IACUC and institutional guidelines.

Creation of the air pouch (Day 0) (Figure 2B)

• 4. Anesthetize mice with isoflurane according to equipment and institution guidelines.

  Inhalation anesthesia is recommended for this protocol, with isoflurane as the anesthetic of choice. Isoflurane causes minimal depression of the cardiovascular and respiratory system; thus, a heating pad is usually not required for this protocol.

  Prior to injection, confirm the level of anesthesia by pinching a paw to verify the lack of a nociceptive response.

  The anesthesia will wear off quickly, so the injection of air should be rapid. With familiarity, speed will increase.

  it is possible, especially when first practicing this protocol, that mice might wake in the anesthesia box and need to be re-treated with isoflurane or for experimenters to anesthetize only one mouse at a time.

  As the anesthesia will wear off quickly, the injection of air should be rapid, but constant.

• 5. (Optional) If you will collect tissue sections from the air pouch, shave the dorsal cervical and thoracic regions of the mice at this step.

• 6. Injection of sterile air.

  1. Wipe the entire back of the mouse at the injection site with 70% ethanol on a cotton ball.
  2. Pull up scruff around the top of the scapulae and inject 3 ml of prepared sterile air between your fingers.

  The air bubble should remain over the mouse’s shoulders, giving it a ‘hunchbacked’ appearance. The bubble should not spread down along the spine, towards the head, or down over the shoulder. If necessary, you can use your fingers to guide the bubble while injecting.

  Experimenter 1 should hold the isoflurane tube/nose cone and mouse’s tail while Experimenter 2 performs the injection.

• 7. Mark and weigh mice.

  1. Using a sharpie, mark mouse with a tail stripe for identification purposes.
  2. Weigh the mouse, record its initial weight as required, and return mouse to home cage.

  It is recommended to have a third experimenter weigh and record weights of the mice. Mice will likely begin recovering from anesthesia during or soon after weighing.

  Individual housing of mice during the air pouch formation and experimental period is generally not required.

Animal monitoring (Day 1 and 2)

• 8. As appropriate per institutional guidelines, check mice for overall health, weight and visible dorsal air pouches each day.

  Mice should display no significant loss of weight or signs of stress.

  Common signs of stress and pain in mice can include behaviors such as head pressing, a hunched posture, awkward walking or stumbling, excessive vocalizing or aggression, lethargy, shallow breathing, decreased grooming and greasy or scruffy fur.

Re-inflation of air pouch (Day 3)

• 9. Prepare for injections and anesthetize mice as listed in step 4.

• 10. Inject 3 ml sterile air to reinflate/ maintain inflation of air pouch.

  Inject in the same spot as on day 1, or as close as possible. It is important to inject within the initial air pouch created.

• 11. Weigh mice, fill out surgical and monitoring paperwork, and refresh tail markings as in Step 8.

  Mice should display no significant changes in weight and should not display any signs of stress or pain.

Animal monitoring (Day 4 and 5) and preparation for experimental exposure

• 12. As appropriate per institutional guidelines, check mice for overall health, weight and visible dorsal air pouches each day.

  Mice should display no significant changes in weight and should not display any signs of stress or pain.

• 13. Prepare tubes, plates, solutions and equipment required for experimental exposure and sample collection. For all materials required, we suggest including 1 or 2 extra tubes and syringes, etc.

  1. Label collection tubes with treatment and mouse number.

     1. Possible tubes needed per mouse depending on analysis to be formed:

     1. 2 ml microcentrifuge tubes for collection of air pouch lavage fluid.
     2. 1.5 ml centrifuge tubes for preparation of lavage fluid aliquots for analysis (2 to 3 aliquots per mouse).
     3. 50 ml conical tubes for collection of air pouch tissue for snap freezing.
     4. 50 ml conical tubes containing 10 ml of 10% formalin for air pouch tissue collection for histological analysis.
  2. 2 ml PBS in 3 ml syringe with 18-gauge needle – 1 syringe per mouse for lavage collection from air pouch.
  3. Sterile scissors and forceps.
  4. Sterile transfer pipets.

Experimental Exposure (Day 6 or later; See Background) (Figure 2C)

• 14. Prepare 1 ml syringes containing 1 ml of positive control substances to be tested and cap with 26-gauge needle (one syringe per mouse).

  We recommend always including a positive control stimulus such as a known chemoattractant or bacterial product to induce inflammation and leukocyte recruitment. The choice of stimulus will vary depending on length of exposure, cell type and pathway activation of interest (for example: neutrophil versus monocyte).

  We routinely use the bacterial peptide fMLP or LPS as a stimulus, but other immunogens are possible.

  1. Some different stimuli that can be used include:

     1. 1 ug/ml fMLP in 0.5% carboxymethylcellulose (CMC).
        See reagents section for preparation of CMC solution
     2. 1 ug/ml Zymosan in PBS.
     3. 0.5 ug/ml IL-8 in 0.5% CMC.
     4. 1 ug/ml E. coli 055:B55 LPS in PBS.
        Immunogens can be prepared in PBS, saline or low-density carboxymethylcellulose as has been reported in multiple studies. We prefer to prepare cytokine or chemokines in CMC as this gives a more robust response.
  2. 15. Prepare 1 ml syringes with 1 ml solvent alone (PBS or CMC, dependent on what test solutions are prepared in) for sham-treated mice and cap with 26-gauge needle.
  3. 16. Prepare 1 ml syringes with 1 ml of exposure stimuli of interest and cap with 26-gauge needle.
     We routinely test the inflammatory response to live bacteria, and thus actively growing bacteria are cultured, washed and resuspended in PBS at the appropriate cell number in 1 ml volume for inoculation to the air pouch. If not using live bacteria and your compound of interest is stable, syringes can be prepared the day before. Syringes of each condition should be color-coded using tape or sharpie to remain organized and reduce delay.
  4. 17. Anesthetize mice as in Step 4.
  5. 18. Inject 1 ml of solvent or stimulus into air pouch.
     You will likely have to refresh the tail markings.
  6. 19. Complete all required monitoring paperwork during the experimental exposure period.
     Mice may have a small increase in weight and should not display any signs of stress or pain.
     Ensure all tubes are labeled and all materials and supplies are ready (Step 14) prior to beginning sample harvest and collection.
     Depending on the cell type, stimulus used and biological response of interest, time points will vary.
     For example, we routinely assess neutrophil recruitment and initial inflammatory response after 2 or 6 hours of exposure, while monocyte/ macrophage recruitment and later inflammatory responses are also assessed after 24 or 48 hours.

Sample collection and tissue harvest (Figure 2D)

• 20. Sacrifice mice using a CO₂ chamber at the relevant time point(s).

• 21. Inject 2 ml PBS (3 ml syringe, 18-gauge needle) into air pouch while holding mouse vertical, tail up, inject so the puncture is at the highest elevation.

• 22. Cover hole with one finger and massage the air pouch gently to loosen cells.

• 23. Cut slit just above air pouch using scissors, puncture air pouch using transfer pipet and pipet up and down to mix, being careful not to allow any leaks.

• 24. Use transfer pipet to remove exudate from air pouch and transfer into a 2 ml centrifuge tube on ice.

  Record the amount of exudate collected from each mouse.

• 25. If isolating the air pouch tissue for analysis, cut around the air pouch between skin and muscle using sterile scissors and forceps.

  1. Intact air pouch membrane and associated skin can be collected together OR the thin air pouch membrane can be separated and collected.
  2. Analysis:

     1. For histological analysis, place tissue in 10% formalin for 24 hours for fixation before proceeding with tissue processing.
     2. For molecular analysis, place tissue in conical tube, snap freeze in liquid nitrogen and store at −80°C until analysis.
        Air pouch tissue can be collected intact or divided into multiple pieces dependent on the analysis to be performed. For example, one air pouch can be divided into multiple pieces to allow histological analysis and molecular analysis such as protein or RNA isolation. The amount of tissue required for each type of downstream analysis may require optimization.

Collecting and analyzing cellular exudate.

• 26. Using collected lavage fluid from Step 24, remove 50 μl to a clean microcentrifuge tube for total cell counting.

  Centrifuge remaining volume for 10 min at 1000 × g, 4°C.

  Cells can be counted using a hemocytometer, automatic cell counter or cellular stain.

• 27. Divide the supernatant into single use aliquots and snap freeze in liquid nitrogen and store at −80°C if further analysis will not be performed immediately.

• 28. The remaining collected cells can now be snap frozen in liquid nitrogen and stored at −80°C OR used immediately for downstream analysis of cellular composition or biological assays.

  The endpoints to be measured will depend on the study goals, and several parameters can be assessed to evaluate inflammation or biological and molecular functions. Examples include measurement of exudate volume, quantification of secreted inflammatory mediators such as cytokines by ELISA or biological function assays. The number of cells recruited to the air pouch can be quantified by using an automatic cell counter or a hemocytometer, and specific cell populations can be identified by flow cytometry (for example: Ly6G for neutrophils, F4/80 for monocytes/macrophages) or quantitated by differential cell staining of cytospin preparations. Biological in vivo functions (for example: bacterial killing, phagocytosis, cellular signaling activation) or molecular analysis (for example: protein or gene expression) can be assessed using collected cell pellets. Tissue samples can be used to quantify molecular changes (protein or gene expression) or histological features by general histological stains or immunohistochemistry using specific antibodies of interest.

Please see representative data in the Understanding Results section (below).

REAGENTS AND SOLUTIONS:

0.5% Carboxymethylcellulose (CMC)

• 250 mg of CMC.

• 50 ml of sterile PBS.

• Sprinkle CMC powder onto PBS solution under continuous stirring at room temperature until fully dissolved (may take up to 30 minutes).

• Centrifuge solution at 400 x g for 10 minutes.

• Store solution on ice until ready to use. Make fresh on the day of use only.

COMMENTARY:

Background Information:

Neutrophil Collection from Blood.

Circulating neutrophils are a central component of the innate immune system and serve as the first line of host defense against invading microbes (Oh et al., 2008). While traditionally thought of as static cells, new knowledge of their ability to actively respond to their environment and modulate inflammation has renewed the focus of scientific investigation. To study neutrophil responses effectively, it is imperative to have a highly pure homogenous cell population (Lakschevitz & Glogauer, 2014; Pelletier et al., 2010) as contamination with monocytes or eosinophils can lead to erroneous interpretations and results (Davey et al., 2011). Several methods for isolating neutrophils from human blood have been developed. These methods include density and gradient centrifugation methods using media such as Ficoll-Hypaque (Ferrante & Thong, 1978, 1980), Percoll (Kuhns et al., 2015) or Ficoll-Hypaque coupled with dextran sedimentation to remove red blood cells (Quach & Ferrante, 2017), as well as antibody-based methods including flow cytometry cell sorting (Dorward et al., 2013) and immunomagnetic bead positive or negative selection (Zahler et al., 1997). Use of the 1-step Polymorphprep density media is advantageous as it is ready to use, easily scalable to accommodate various blood volumes and does not require gradient formation or secondary sedimentation to remove the majority of red blood cells. This method is also more cost and time-effective than flow sorting and antibody-based separation methods. Neutrophil isolation using Polymorphprep typically results in purity greater than 90% with viability exceeding 98% (Ferrante & Thong, 1978; Zhou et al., 2012), which is generally sufficient for most functional assays. However, for analysis requiring extremely pure neutrophil populations, such as transcriptomic analysis, negative-selection antibody based-magnetic separation is recommended.

Oral Collection of Neutrophils.

Neutrophils form a heterogenous population, with various subpopulations having distinct roles at different tissue sites and in health and disease (Beyrau et al., 2012; Hong, 2017; Silvestre-Roig et al., 2016). In the oral cavity, neutrophils are prominent at mucosal surfaces and in saliva, having migrated from the peripheral blood circulation through the gingival microvasculature and gingival sulcus epithelium into the gingival crevicular fluid (GCF). Even in a healthly individual, oral neutrophils are prominent and comprise greater than 90% of the immune cells in GCF where they function to maintain homeostasis (Delima & Van Dyke, 2003; Metcalfe et al., 2021). The oral cavity represents the interface to the external environment and is continuously exposed to dietary, biological and environmental influences and provides an easily accessible and non-invasive source of biological samples for analysis. During oral inflammatory states such as periodontitis, the oral neutrophil load increases up to 10-fold (Bender et al., 2006; Jones et al., 2020; Landzberg et al., 2015) along with altered cell-surface receptor profiles (Chadwick et al., 2021; Fine et al., 2016; Rijkschroeff et al., 2016), transcriptional changes, phenotypic and functional output changes (Jones et al., 2020; Lakschevitz et al., 2013a, 2013b; Moonen et al., 2019) as compared to those observed during oral health. Oral neutrophils also have significant value for characterizing non-dental conditions as well as diagnostic or prognostic potential. For example, oral neutrophil loads are elevated during cardiopulmonary bypass (Wilcox et al., 2014), and neutrophil plasticity and impaired functionality has been characterized during oral cancer (Ueta et al., 1993; Ueta et al., 1994), and functional changes characterized in oral neutrophils of individuals that smoke (Archana et al., 2015).

Murine Air Pouch.

The air pouch model is a useful in vivo model of localized inflammation. The repeated injection of air into the dorsal interscapular region of the mouse separates the dermis and epidermis from the subcutaneous tissues, forming an enclosed cavity. The air-pouch model was initially developed as an in vivo model to represent inflamed synovium tissue as a thin lining membrane tissue of 2- 3 cells in thickness forms, which is composed primarily fibroblast-like cells and macrophages with surrounding microvasculature (Edwards et al., 1981). This protocol describes use of a 6-day air pouch with short biological exposure times to study acute inflammatory responses. However, adjusting the duration of air-pouch formation (with repeated air injection to maintain inflation) to allow further maturation of the formed lining tissue provides flexibility in parameters and microenvironment of study. With prolonged duration of up to 14 or even 30 days a chronic system may be modeled, as the air pouch lining tissue becomes thicker and more fibrous (Edwards et al., 1981). Within the 6-day period described in this protocol, development of the lining including increased vasculature and cellular organization of the tissue, provides an effective barrier which also contributes to increased reactivity of the air pouch (Sedgwick et al., 1983). Injection of irritants (such as croton oil or carragean) or chemokines/cytokines into this subcutaneous cavity provides a model to assess general inflammatory response or specific immune activation pathways. This model system has been adapted for many biological stimuli or exposures; for example, it has been used to study granulomatous inflammation (Vane et al., 1994), angiogenesis (Eteraf-Oskouei et al., 2014), tumor development (Mishalian et al., 2014), biomaterial compatibility (Vandooren et al., 2013), and oxidative stress (Li & Frei, 2006). This model can also be used to assess the role of live bacteria and bacterial products in the promotion or inhibition of inflammatory and biological responses (Jones et al., 2020; Koh et al., 2015), as well as screening of potential anti-inflammatory compounds (Li et al., 2021) or therapeutic molecules. Efficacy of therapeutic agents, antagonists or agonists can be assessed by either by local injection in the air pouch to examine direct interactions (Herrera et al., 2015; Koh et al., 2015; Perretti & Flower, 1993b) or inoculation intravenously or interperitoneally (Perretti & Flower, 1993a) to examine cell recruitment and activity to the air pouch, prior to stimulus of interest. The use of mice with genetic mutant backgrounds also allows investigation of the role of specific host cell pathways in inflammation and immune cell function. The air pouch model is advantageous compared to other in vivo models of immune cell recruitment and responses, such as the peritonitis model, as there is less risk of blood or other biological contamination of samples due to the high vascularization and presence of nearby organs in the peritoneal cavity. Most importantly, the air pouch model allows formation of an enclosed chamber allowing reproducibility, detailed monitoring of biological responses and immune cell recruitment together with collection of a sufficient amount of sample to allow analysis of multiple parameters from the same animal.

Critical Parameters:

Neutrophil Collection from Blood.

When isolating neutrophils from peripheral blood, it is imperative to store blood in appropriate vacutainers with a reputable anticoagulant. Numerous compounds have been tested as anticoagulants, including sodium citrate (Ernst, 2007), lithium heparin (Çuhadar, 2013), and EDTA (Ernst, 2007). Of these, EDTA is our preferred choice of anticoagulant with respect to maintaining neutrophil responses (Banfi et al., 2007; Lam et al., 2004; Shabnam et al., 2014). During collection, invert each tube three to four times as soon as it is filled to ensure even incorporation of the anticoagulant. After blood collection, the anticoagulated blood should be kept at room temperature and used within two hours. During separation, the blood, reagents, and centrifuge should not be below 17C as Polymorphprep uses its high osmolarity to cause water loss from red blood cells, causing them to sediment out of the blood and this process is most effective at room temperature. Neutrophils are sensitive to vibration and physical stress and are prone to activation, so it is crucial to ensure centrifugation steps are performed with moderate acceleration and minimal brake and to handle them gently. To reduce cell death and/or spontaneous activation, the isolated cells should be utilized with 2-3 hours. If cells will be incubated for extended periods for experimental analysis (1 hour or more), we recommend storage and use in a cell culture media such as RPMI 1640 while short term incubations can be performed in either a buffer solution (such as HBSS) containing magnesium and calcium, or a cell culture medium.

Oral Collection of Neutrophils.

Many of the critical parameters for blood neutrophil collection are also of concern for oral neutrophil isolation, in addition to these noted below. When collecting oral salivary rinses, it is important to keep samples on ice due to the high lytic nature of oral neutrophils. Appropriate timing of 3 minutes between rinses is also required to allow trafficking and repopulation of neutrophils to the oral cavity. During neutrophil enrichment, it is critical to filter collected saliva through sequential filtration by gravity flow rather than vacuum or pressure, as the latter two can increase cell activation and decrease purity of neutrophil samples due to contamination with epithelial cells, monocytes or other debris.

Murine Air Pouch.

Mice must be acclimated to their environment for a minimum of 1 week prior to experiments to prevent undue stress from interfering with results. Lavage fluid volume collected can vary between mice, but large deviation from the expected volume (approx. 2 ml) indicates an issue such as a leakage of the pouch. It is important to immediately separate cells from the exudate fluid and process the cells in order to minimize cell death. Failure to separate cells and exudate can compromise results as the neutrophils will continue to produce inflammatory mediators or consume those that are already present. Finally, following the removal of cells, the exudate must be aliquoted and snap frozen in liquid nitrogen (or ethanol/dry ice) as quickly as possible to minimize degradation of inflammatory mediators. For the same reason, avoid repeated freeze-thaw cycles of the aliquoted exudate.

Troubleshooting:

Table 1.

Troubleshooting Guide for Neutrophil Isolation from Human Blood

Problem	Possible Cause	Solution
Poor separation of bands	Excessive disturbance of the blood/polymorph boundary	Collect both bands into a fresh tube, fill tube to 14 ml with HBSS −/− and spin 10 min at 400 x g (accel. and decel. 4). Resuspend pellet in 5 ml fresh HBSS −/−, gently layer on top of 5 ml fresh polymorph prep and repeat Step 5.
Excessive number of dead neutrophils in finial isolate	Use of improper solutions; excessive time taken isolating cells	Ensure cells are isolated as quickly as possible. Ensure solutions are being used at the correct dilutions and appropriate buffers and components are used.
Isolated, untreated neutrophils showing high levels of activity in negative control in experiments	Contamination, activation	Ensure no contamination of solutions or container with endotoxin. Be careful to remove all of the mononuclear cell band and take only the lower band when isolating neutrophils. Use sterile disposable polypropylene plasticware when possible. Handle containers with cells gently as excessive disturbance of cells by vortexing or forceful pipetting can cause activation.

Table 2.

Troubleshooting Guide for Neutrophil Isolation from the Oral Cavity

Problem	Possible Cause	Solution
Contamination with epithelial cells and/or monocytes	Improper filtration or high-pressure filtration used	Pooled rinse samples must be filtered sequentially by size to remove debris and larger cells first. Pushing samples through the filter by plunger or other high-pressure methods will increase cell debris and prevent appropriate removal of unwanted cell types.
Clogging of filters or low cell recovery	High mucus content in sample	Very carefully and gently pipette sample up and down to resuspend to filter again and a small volume of extra sterile 0.9% NaCl solution can be added to dilute. Initial collected rinses can be centrifuged at low speed (~120 x g for 5 minutes), resuspended in fresh 0.9% NaCl solution prior to beginning filtration.
Low cell recovery	Cells not allowed to repopulate the oral cavity. Number of cells trafficking to the oral cavity is lower than expected.	At least 3-minute intervals between rinses are required to allow for cells to traffic to the oral cavity. Cell numbers present in the oral cavity vary depending on the overall and local (oral cavity) inflammatory state of the subject. Extending the number of rinses collected will increase the number of neutrophils obtained.

Table 3.

Troubleshooting Guide for Murine Air pouch Model of General Inflammation

Problem	Possible Cause	Solution
Formation of two air pouches.	Instead of reinflating/ maintaining the initial air pouch on Day 3, the needle went to a different layer of skin.	Reinflate the pouch as close to the original location as possible and pinch up the skin and be sure to insert the needle into the cavity.
Recovery of a small volume of exudate.	Leakage of the air pouch caused by excessive roughness in handling or fights between mice.	Carefully handle mice throughout and when collecting extrudate gently check the periphery of the pouch along the sides of the mice.
Large variation in inflammatory markers or other downstream analysis within an experimental group.	Exudate or tissue samples not processed quickly enough. Exudate not kept on ice or snap frozen immediately. Repeated freeze-thaw cycles. Tissue samples not processed immediately (i.e., formalin fixation or snap-frozen).	Quickly process all lavage samples and keep cells and exudate on ice as much as possible. Ensure tissue samples are collected quickly as processed appropriately as soon as possible.
Low cell yield from air pouch.	Cells remain attached to air-pouch membrane. Not using appropriate time point or stimulus exposure.	While washing the air pouch with PBS alone is generally effective, washing with PBS containing 3 mM EDTA may improve cell recovery. Consider modifying the length or exposure and dose and stimulus type depending on cell type of interest. Generally, neutrophils are recruited to the air pouch within 1 to 6 hours.

Understanding Results:

Neutrophil Collection from Blood.

Collecting neutrophils from peripheral blood utilizing the “1 step” method with 1 step polymorph media is a simple, yet delicate process that yields purified neutrophils for in vitro experiments (Figure 1). On average, we isolate 3-4 x 10⁷ neutrophils from 36 ml of blood collected from healthy individuals, with the viability of our cells ranging from 90-95%. Neutrophils can be used to investigate a multitude of neutrophil phenotype-based parameters including, activation, granule acquisition, cytokine release, killing and phagosome maturation. As an example, we have used immunofluorescence to look at early endosome antigen 1 (EEA1) as an indicator of early phagosomes during a time course following phagosome maturation within neutrophils (Figure 3).

Figure 3. Immunofluorescence analysis of blood neutrophils.

Example of phagocytic uptake and phagosomal analysis of isolated neutrophils. Neutrophils were incubated with IgG opsonized 3 μm latex beads, fixed with paraformaldehyde, then stained to label external beads (red), early phagosomal marker EEA1 (green) and the nucleus stained with DAPI (blue). Examples of positive (arrow) and negative internal beads are shown.

Oral Collection of Neutrophils.

From oral rinses of healthy individuals, we routinely isolate between 0.2 - 2.0 X 10⁶ neutrophils. During inflammatory states or infection, increased neutrophil load is expected: during periodontitis 0.3 - 9.2 X 10⁶ neutrophils are isolated from the oral cavity in our experience. Enriched oral neutrophil samples can be directly analyzed (functional assays, gene or receptor profiling) to characterize in vivo immune responses. As an example, enriched oral neutrophils from individuals with periodontitis secrete more cytokines compared to oral neutrophils in health. Alternatively, the enriched oral neutrophils can also be cultured in vitro with an experimental stimulus of choice followed by subsequent downstream assays.

Murine Air Pouch.

The exudate volume recovered should be equivalent to the total injected volume of the lavage solution plus the stimulant/test solution, though the volumes recovered will likely vary from animal to animal. The time between formation of the pouch and treatment with stimulant will influence the magnitude of the inflammatory response: a one-day-old pouch will show much less reactivity than a six-day-old pouch (Sedgwick et al., 1983). We routinely harvest approximately 0.4 - 1 × 10⁵ cells in total from a 6-day unstimulated air pouch, while a positive-control stimulus such as LPS for 6 hours routinely recruits 1 – 1.5 X 10⁶ cells. Cell recruitment and cellular composition to the air pouch will vary dependent on stimulus used and length of exposure. The majority of cells harvested from an unstimulated pouch are likely to be monocytes and macrophages, with very low numbers of polymorphonuclear cells and low levels of pro-inflammatory mediators. Air pouch membrane tissue lining from sham-treated mice should show low levels of immune cell infiltrate compared to those from mice exposed to bacteria or other inflammatory agents (Figure 4A). A variety of downstream analyses can be performed directly on collected lavage fluid and recruited cells. For example, secreted inflammatory mediators can be measured in lavage fluid (Figure 4B) and cellular signaling activation can be quantified in cells collected from the air pouch (Figure 4C).

Figure 4. Exposure to Treponema denticola increases neutrophil recruitment and inflammatory signaling in the air pouch model.

A) H&E staining of air pouch lining in sham (top) shows fewer neutrophils (indicated with yellow arrows) than T. denticola exposure (bottom). Tissue was harvested from male C57BL/6J mice six hours after inoculation of 6-day old air pouches. Representative downstream analysis of collected lavage samples. B) Selected inflammatory markers are elevated in lavage fluid as measured by ELISA and C) Rac1 signaling is activated in recruited neutrophils (measured by GLISA) following exposure to T. denticola in the air pouch.

Time Considerations:

Neutrophil Collection from Blood

Isolating neutrophils from peripheral blood using 1 Step Polymorphs takes approximately 1-1.5 hours, with time varying depending on the total volume of blood that is being used for cell isolation.

Oral Collection of Neutrophils

Collection of oral rinses from donors takes approximately 1 –2 hours, depending on how many rinses are needed to collect enough volume for cell isolation. Neutrophil enrichment and isolation from oral rinses takes approximately 1- 2 hours.

Murine Air Pouch

The overall time of this experiment will vary greatly depending on the number of animals, how long the experimenters allow the air pouch to develop prior to treatment for, and the number of experimenters. As an example, with 3 experimenters working on 40 mice, creation of the air pouch will take approximately 1.5 hours if the mice are also shaved, while re-inflation should take around 1 hour. Monitoring alone should take approximately 30 minutes. Experimental exposure should again take approximately 1 hour, while sample collection should take about 2 hours at each time point. When collecting samples on the same day as the experimental exposure, expect to need the full day. If the harvested cells require preparation that same day (ex: for flow cytometry), the experiment will likely take into the late evening provided the initial exposure was in the morning. If the experimental exposure includes live bacteria, they must be prepared shortly before the exposure which can require an additional 1 to 1.5 hours.

DATA AVAILABILITY STATEMENT:

The data, tools, and material (or their source) that support these protocols are available from the corresponding author upon request.