Isolating And Immunostaining Lymphocytes and Dendritic Cells from Murine Peyer's Patches

Podsumowanie

There is an increasing interest in understanding the immunological functions of specific subpopulations of cells in Peyer's patches (PPs), the primary inductive sites of gut-associated lymphoid tissues. Here we outline parallel protocols for preparing PP single cell preparations for flow cytometric analysis and PP cryosections for immunostaining.

Streszczenie

Peyer's patches (PPs) are integral components of the gut-associated lymphoid tissues (GALT) and play a central role in intestinal immunosurveillance and homeostasis. Particulate antigens and microbes in the intestinal lumen are continuously sampled by PP M cells in the follicle-associated epithelium (FAE) and transported to an underlying network of dendritic cells (DCs), macrophages, and lymphocytes. In this article, we describe protocols in which murine PPs are (i) dissociated into single cell suspensions and subjected to flow cytometry and (ii) prepared for cryosectioning and immunostaining. For flow cytometry, PPs are mechanically dissociated and then filtered through 70 μm membranes to generate single cell suspensions free of epithelial cells and large debris. Starting with 20-25 PPs (from four mice), this quick and reproducible method yields a population of >2.5 x 10⁶ cells with >90% cell viability. For cryosectioning, freshly isolated PPs are immersed in Optimal Cutting Temperature (OCT) medium, snap-frozen in liquid nitrogen, and then sectioned using a cryomicrotome. Tissue sections (5-12 μm) are air-dried, fixed with acetone or methanol, and then subjected to immunolabeling.

Wprowadzenie

Peyer's patches (PPs) are macroscopic aggregates of organized lymphoid follicles present throughout the small intestine of humans and mice (Figure 1) and constitute the primary sites at which mucosal immune responses are initiated against dietary antigens, commensal bacteria, microbial pathogens, and oral vaccines ^1-4. Unlike other peripheral lymphoid tissues such as the mesenteric lymph nodes, PPs lack afferent lymphatics. As such, adaptive immune responses in PPs are driven in response to antigens derived from the intestinal lumen. The sampling of luminal antigens is accomplished the by the follicle-associated epithelium (FAE), which consists of both enterocytes and antigen-sampling cells known as M cells. Beneath the FAE, in the sub-epithelial dome (SED) region, lies a network of dendritic cells (DCs) intermingled with macrophages, B cells, and CD4⁺ T cells ^5-9. At the core of each PP lymphoid follicle are follicular dendritic cells (FDCs) and a B cell-rich central germinal center, flanked by T cell-rich interfollicular zones. Antigen sampling by PPs results in the development of IgA+ B cell plasmablasts and CD4⁺ effector and memory cells that seed the surrounding lamina propria and provide immunity to a wide range to mucosal invaders.

Dissecting the complex immunologic events associated with antigen sampling, processing, and presentation in PPs is a daunting task, considering that PP cells constitute only a tiny fraction of the total lymphoid cells in intestinal mucosa. To aid in the in vitro characterization of cells in this environment, we provide a protocol for preparing total mouse PP cells for flow cytometric and functional analysis, as well as a protocol for preparing PP cryosections for immunofluorescence microscopy and immunohistology. Our protocol for the isolation, characterization, and immunostaining of mouse PP cells is not novel per se, as evidenced by the fact that there are numerous references dating back more than 25 years that cite these techniques ^5,6,9-11. Rather, our protocol provides a streamlined (and visual) method for investigators collecting PPs for the first time. The techniques we describe are easily mastered and readily yield large numbers of cells with >90% cell viability. The cryosectioning protocol yields highly reproducible serial sections ideally suited for immunofluorescence staining and confocal imaging. Furthermore, our protocol complements two other recent JoVE articles. The first, by Fukuda and colleagues, describes the use of ligated ileal loop assays to assess the uptake of pathogenic bacteria by PP M cells ¹². The other, by Geem and colleagues, describes the isolation and characterization of DCs and macrophages from the mouse intestinal mucosa, but explicitly excludes PPs from their analysis ¹³.

Protokół

Animals were housed under conventional, specific pathogen-free conditions and were treated in full compliance with the Wadsworth Center's Institutional Animal Care and Use Committee (IACUC) guidelines.

1. Oral Gavage

1. (Optional) Gavage mouse strain of choice with antigen or microbes of interest using a 22 G x1.5-in. blunt-end feeding needle (Popper Scientific, New Hyde Park, NY). Delivery volumes should not exceed 400 μl per mouse.

2. Isolation of PP Cells for Flow Cytometry

1. Euthanize mice by CO₂ asphyxiation, according to institutional animal care and use committee (IACUC) guidelines.
2. Cleanse the abdomen with 70% ethanol or betadine prior to surgery. Perform a standard laparotomy that involves a single (1 cm) incision using surgical grade scissors along the midline beginning about 1.5 cm from the base of the rib cage. Expose the peritoneal cavity and identify the cecum. Snip the terminal small intestine at the ileal-cecal junction and gently remove the intestine in its entirety. Take care not to hyperextend the intestinal tissue as it is being removed from the peritoneal cavity.
3. Lay the small intestine on a bed of moist paper towels or Kimwipes. Wet the small intestine gently with saline to prevent tissue dehydration. Visually identify individual PPs located on the anti-mesenteric side of the intestine (Figure 1). Typically, a single mouse has 5 to 10 visible PPs that are evenly distributed from the duodenum (proximal) to ileum (distal). On average, four mice will yield 23 PPs.
4. Using curved surgical scissors, gently excise individual PPs and place them in cold Hank's Balanced Salt Solution (HBSS). Note, the scissors should be placed curve-side up just above the PP and then gently applied to the tissue. Excise only the PP (not surrounding tissue) and transfer to cold HBSS.
   Note: All incubations and centrifugation steps from this point forward in Section 2 should be done at 4 °C to preserve cell viability.
5. Transfer PPs into 5 ml Spleen Dissociation Medium and incubate for 15-20 min at 37 °C while vigorously shaking at 250 rpm. Longer incubation time will negatively affect cell viability.
6. To generate a single cell suspension, place PPs on a sterile (autoclaved) 70 μm nylon mesh cell strainer and forcibly grind the tissue into the mesh using the base of a plunger from a 1 cc syringe. Alternatively, sandwich PPs between two frosted sterile glass microscope slides (autoclaved in envelopes) and gently grind tissues with a back and forth motion. Use a sterile transfer pipette to transfer the cell suspension into a 5 ml Falcon tube.
7. Add EDTA to a final concentration of 1 mM to the cell suspension. Incubate on a rocker for 5 min at room temperature.
8. Decant cell suspension through a second 70 μm cell strainer to remove any remaining cellular aggregates or tissue debris. Collect cells in a 15 or 50 ml conical tube.
9. Subject cells to gentle centrifugation (5 min at 500 x g). Decant supernatant and resuspend cells in flow buffer, which is phosphate buffered saline (PBS) containing 1% fetal calf serum (FCS).
10. To determine cell number and viability, dilute cells 1:10 into trypan blue and visually inspect cells using a light microscope and a hemocytometer. Alternatively, a Countess cell counter (Invitrogen) or similar instrument can be used to automatically enumerate cell numbers and viability.
    Starting with 20-25 PPs, this protocol should yield >2.5 x 10⁶ total cells. This equates to ~ 0.8-1.2 x 10⁶ cells per mouse.

Antibody Labeling of Cells for Flow Cytometry

11. Dispense cells (10⁵ per well) into wells of a 96-well round bottom plate.
12. Subject plate to gentle centrifugation (5 min at 1,000 x g), and decant supernatant by gently inverting the plate.
13. Resuspend cells in Fc block buffer and incubate on ice for 15 min. While there are a number of commercially available Fc block buffers (e.g. rat anti-mouse CD16/CD32, BD Biosciences), we simply use spent medium from a rat B cell hybridoma (ATCC 2.4.G2) that secretes a monoclonal IgG₁ against murine Fcγ receptors for this step.
14. Subject plate to gentle centrifugation (5 min at 1,000 x g) as above, and decant supernatant by gently inverting the plate.
15. Add fluorophore-conjugated antibodies directly to cell suspensions at desired dilution, and incubate on ice for 30 min with continuous rocking.
16. Subject plate to gentle centrifugation (5 min at 1,000 x g), and decant supernatant by gently inverting the plate.
17. Wash cells with 100 μl of flow buffer.
18. Centrifuge plate at 1,000 x g for 5 min, and decant supernatant by gently inverting the plate.
19. Resuspend cells 400 μl fixation buffer, which consists of 300 μl of flow buffer and 100 μl of 1% paraformaldehyde in PHEM buffer (60 mM PIPES, 25 mM HEPES, 10 mM EGTA, 4 mM MgCl₂ at pH 6).
20. Subject cells to flow cytometry using a FACSCalibur or equivalent. Analyze results using Cell Quest Pro software version 5.2.

3. Preparation of PP Cryosections

1. In advance of tissue collection, add Optimal Cutting Temperature (OCT) compound to 7x7x5 mm plastic base molds. Also prepare a pool of liquid nitrogen (>750 ml) in a dewar or ice bucket.
2. Euthanize mouse and perform a laparotamy, as described above. Remove intestine and place on wet paper towels.
3. To isolate PPs for cyrosectioning, cut 0.5 cm transverse segments of small intestine containing a single PP and transfer tissue to a Petri dish containing PBS. Gently rinse the lumen of these segments with PBS to remove unwanted fecal debris.
4. Place intestinal tissue segments vertically inside base molds containing OCT. Immerse the molds in liquid nitrogen and allow tissue to freeze completely (1-2 min). The color of the OCT will change from clear to white when the tissue is fully frozen. For a more gradual cooling (e.g. to reduce cracking of the OCT), immerse mold in a bath of isopentane cooled with liquid nitrogen vapors. Note: Wear appropriate safety goggles when working with liquid nitrogen.
5. Remove frozen base molds from the liquid nitrogen using forceps and allow the mold to warm at room temperature just long enough (~10-15 sec) to allow the edges of the OCT block to soften. To then remove the frozen block of OCT from the mold, invert the mold and press gently on the back to eject the block onto a clean 4 x 4 cm piece of aluminum foil. Immediately wrap the tissue in the foil and place in a labeled specimen bag stored in liquid nitrogen or on dry ice. Alternatively, transfer tissue in a freezer (<-20 °C). Use the tissue within a week, otherwise the blocks will become brittle with age.

Cryosectioning

6. Cryosection the tissue using a Leica CM3050S cryomicrotome or equivalent. The chamber temperature on the cryostat should be set to -21 °C.
7. Strive for sections that are 12-14 μm in thickness.
8. Collect sections onto Fisher SuperFrost Plus (or equivalent) glass microscope slides and inspect visually using a standard low power light microscope. If multiple sections are being collected on a slide, ensure that they are clustered together for downstream staining applications.
9. Store the slides at room temperature in a slide box and, ideally, use them within a few days.

Antibody Labeling of Cryosections and Confocal Microscopy Analysis

10. Immerse slides in acetone (or methanol) in tissue staining jar or racks for 2 min. Prolonged incubation may cause tissues to detach from the slide.
11. Transfer slides to new staining jar and wash 3 times with PBS-T (1X PBS with 0.05% Tween-20) for 3 min each.
12. Encircle the sections with an ImmEdge hydrophobic pen. This will ensure that reagents and antibodies will stay confined to that area during incubations.
13. Incubate slides in block buffer (2% goat serum in PBS) for 30 min at 37 °C in a humidified chamber, protected from light.
14. Incubate slides with Fc block buffer (see above) for 10 min at 37 °C.
15. Dip slides in PBS-T and dry area around sections using Kimwipes or other absorbent tissue. Take care not to touch the actual tissue sections.
16. Overlay tissue sections with primary antibody solution and incubate for 30 min at 37 °C in a humidified chamber. Primary antibody is typically diluted in block buffer to a final concentration of 20 μg/ml. The use of directly-labeled primary antibodies is desirable, because as many as three antibodies can be combined directly at this step. Directly-labeled antibodies also eliminate the need for additional antibody incubation steps.
17. Wash slides 3 times (5 min each) by immersion in PBS.
18. If necessary, incubate slides with relevant secondary antibodies for 30 min at 37 °C in a moisture chamber.
19. Wash slides 3 times (5 min each) by immersion in PBS.
20. Incubate slides for 4 min in 1% paraformaldehyde in PHEM buffer, as described above.
21. Rinse slides with PBS for 1 min.
22. Dry area around sections using Kimwipes or other absorbent tissue. Take care not to touch the actual tissue sections. Using a pipette or dropper, gently place a drop of Prolong Gold mounting medium directly onto the section. Apply a glass coverslip onto the mounting medium taking care to avoid bubbles. Use blotting paper to absorb any excess mounting medium.
23. Seal coverslips to microscope slides with commercial nail polish and allow to air dry.
24. View slides with a Leica TCS SP5 or equivalent confocal microscope.

Wyniki

Flow cytometric analysis of monodisperse suspensions of total PP cells reveals a clear distinction between good and poor cell preparations. In good cell preparations with over 80% viability, the vast majority of cells demonstrate high forward scatter (FSC), an indicator of high cell volume, and low side scatter (SSC), an indicator of low cell granularity (Figure 2A). In this experiment, we also intentionally prepared a "poor cell preparation" by incubating PP cells during the isolation steps at ro...

Dyskusje

In this article, we have provided parallel protocols for preparing PP single cell preparations for flow cytometric and functional analysis and cryosections for immunostaining. Both methods are highly reproducible and readily accessible, provided a flow cytometer and cryostat are available. For first time investigators it should be pointed out that when compared to the spleen, total cell yields from PPs are relatively meager. Nonetheless, the protocol we outline generally yields between 0.8-1.2 x 10⁶ total PP c...

Materiały

Name	Company	Catalog Number	Comments
Item	Company	Cat. #	Comments (optional)
OCT Compound	Tissue-Tek	4583
7x7x5 mm Base Molds	Fisherbrand	22-363-552
ImmEdge Pen	Vector Labs	H-4000
Superfrost Plus Slides	Thermo Scientific	4951
Edge Rite Blade	Thermo Scientific	4280L
Anti-Mouse CD11c-PE	eBioscience	17-0114-82
Anti-Mouse CD45R/B220-APC	BD Pharmigen	553092
Anti-Mouse CD3-FITC	BD Pharmigen	561798
Anti-Mouse CD4-PE	BD Pharmigen	553652
Anti- Mouse CD8-PE	BD Pharmigen	553032
Anti-Mouse CD19-PercP	BioLegend	115531
Hank's balanced salt solution (HBSS) without phenol red	Fisher Scientific	14175-079
70 μm cell strainer	BD Falcon	352350
Spleen Dissociation Medium	Stem Cell Technologies	7915
Goat serum	Invitrogen	16210-072
Fc Block	ATCC 2.4.G2	HB-197	Supes obtained from cell line 2.4.G2
Curved Scissor	F.S.T	14061-09
Cryostat	Leica	3050S
FACS Calibur	BD
Countess Cell Counter	Invitrogen
Hematoxylin	Richard Allan	7211
Eosin	Richard Allan	71304
Formalin	Starplex Scientific	3661
			Table 1. Reagents and equipment used in this study.