Unraveling Key Players of Humoral Immunity: Advanced and Optimized Lymphocyte Isolation Protocol from Murine Peyer's Patches

Summary

In this study, we present a novel and effective protocol for the isolation of lymphocytes from Peyer's Patches (PPs), which can be subsequently used for in vivo and in vitro functional assays as well as flow cytometric studies of follicular T helper and germinal center B cells.

Abstract

In the gut mucosa, immune cells constitute a unique immunological entity, which promotes immune tolerance while concurrently conferring immune defense against pathogens. It is well established that Peyer's patches (PPs) have an essential role in the mucosal immune network by hosting several effector T and B cell subsets. A certain fraction of these effector cells, follicular T helper (TFH) and germinal center (GC) B cells are professionalized in the regulation of humoral immunity. Hence, the characterization of these cell subsets within PPs in terms of their differentiation program and functional properties can provide important information about mucosal immunity. To this end, an easily applicable, efficient and reproducible method of lymphocyte isolation from PPs would be valuable to researchers. In this study, we aimed to generate an effective method to isolate lymphocytes from mouse PPs with high cell yield. Our approach revealed that initial tissue processing such as the use of digestive reagents and tissue agitation, as well as cell staining conditions and selection of antibody panels, have great influence on the quality and identity of the isolated lymphocytes and on experimental outcomes.

Here, we describe a protocol enabling researchers to efficiently isolate lymphocyte populations from PPs allowing reproducible flow cytometry-based assessment of T and B cell subsets primarily focusing on TFH and GC B cell subsets.

Introduction

The entire gastrointestinal tract from the beginning to the end is decked with an extensive lymphoid network that contains immune cells more than any other organ in human and mouse¹. Peyer's patches (PPs) constitute a major component of the intestinal branch of this cellular immune organization, so-called gut-associated lymphoid tissue (GALT)^2,3. Within PPs, thousands of millions of antigens derived from dietary materials, commensal microbiota and pathogens are being sampled continuously, and when necessary appropriate immune responses toward them are mounted thus maintaining intestinal immune homeostasis. In that sense, PPs could be named as "tonsils of the small intestine". PPs consist of major sub-compartments: subepithelial dome (SED), large B-cell follicle zones; the overlying follicle-associated epithelium (FAE) and interfollicular region (IFR) where T cells are located⁴. This unique compartmentalization of PPs enables different effector cell subsets to cooperate, thus, confers immunocompetence in the gut.

PPs lack afferent lymphatics, and due to this reason, antigens transported to PPs from small intestine are not being conducted through lymphatic vessels in contrast to most of the other lymphoid organs. Instead, specialized epithelial cells located in FAE, so-called M cells, are responsible for the transfer of luminal antigens into the PPs⁵. Subsequently, the transported antigens are picked up by the dendritic cells (DCs) and phagocytes that are located in the subepithelial dome (SED) region beneath the FAE^6,7. This antigen sorting process by DCs in the PP is crucial to initiate adaptive immune response⁸ and subsequent generation of IgA secreting cells⁹.

Due to the heavy antigenic burden from commensal flora and dietary material, PPs host endogenously activated effector T and B cell subsets in great abundances such as TFH and IgA⁺ GC B cells¹⁰, suggesting that PPs represent a site of active immune response¹¹. Detection of up to 20–25% TFH cells within total CD4⁺ T cell compartment and up to 10–15% GC B cells within total B cells is possible in PPs collected from unimmunized young C57BL/6 mice¹². In contrast to other T helper cell types (i.e., Th1, Th2, Th17 cells), TFH cells show unique tropism into B cell follicles primarily owing to CXCR5 expression, which promotes TFH cell homing along CXCL13 gradient¹³. In the B cell follicle zones of PPs, TFH cells induce IgA class switch recombination and somatic hypermutation in activated B cells from which high-affinity IgA producing cells differentiate¹⁴. Subsequently, these antibody-secreting plasma cells migrate to the lamina propria (LP) and regulate immune homeostasis in the gut¹⁰.

Identification and characterization of TFH and GC B cell populations within PPs might enable researchers to investigate the dynamics of humoral immune responses under steady-state conditions without the need of time-consuming immunization models traditionally used in TFH-GC B cell studies^15,16,17,18. Analyzing TFH cells within PPs is not as straightforward as other cell subsets. Technical challenges include identifying ideal tissue preparation conditions, surface antibody-marker combination, as well as selecting appropriate positive and negative controls. Both TFH and PP research fields exhibit great variability in terms of experimental procedures and are far from conferring a consensus to establish standardized protocols due to several reasons. First, each cell subset within PPs tends to be affected differentially by tissue preparation conditions requiring further modifications in a cell subset-specific manner. Second, there is a significant discrepancy among the reported methods regarding the details of cell preparation from PPs. Third, the number of protocol-based comparative studies investigating ideal tissue preparation techniques and experimental conditions for PP and TFH research is rather limited.

Current protocol-based studies suggested for PP cell preparation^19,20,21,22 were not TFH- or GC B cell-oriented. Moreover, some tissue preparation conditions recommended for PPs^19,20 such as collagenase-based digestion were found to affect the outcome of TFH identification by flow cytometry negatively¹⁸. On this basis, we reasoned that an optimized, standardized and reproducible protocol that can be used to study TFH and GC B cell dynamics within PPs would be valuable to investigators working on this topic. This need gave us the impetus to generate an improved and up-to-date protocol for the isolation and characterization of PP lymphocytes that is finely optimized for cellular recovery, viability, and efficiency for flow cytometric characterization of several T and B cell subsets. We also aimed to exclude several laborious preparation steps suggested in previous protocols, thereby, reducing the required manipulations and time for tissue and cell preparation from PPs.

Protocol

All studies and experiments described in this protocol were conducted under guidelines according to Institutional Animal Care and Use Committee (IACUC) of Beth Israel Deaconess Medical Center.

1. Designing Experimental Set-up and Mouse Groups

1. (Optional) Co-house the experimental mice to facilitate horizontal transmission of gut microbiota between experimental mice and to reduce non-specific variability within PP lymphocytes. Additionally, use littermate controls of the same gender to minimize variability.

2. Surgical Excision and Tissue Preparation Steps:

1. Surgical removal of the Small Intestine (SI)
   1. Euthanize mice using CO₂ asphyxiation or any equivalent method approved by the institutional animal ethics committee.
   2. Transfer the mouse to a dedicated area for surgical excision. Place backside down and sanitize the abdomen with 70% ethanol. Perform a laparotomy by cutting the abdominal skin and peritoneum along the midline from pubis to the rib cage thus opening the peritoneal cavity.
      NOTE: Perform the first incision on a relatively small area of the skin to avoid penetrating the peritoneal cavity and damaging intestinal tissue. Continue the excision until desired anatomical border.
   3. Identify the caecum, which is an ideal landmark for the detection of the terminal ileum, which constitutes the distal segment of the small intestine.
      NOTE: Caecum is usually located on the lower left side of the mouse abdomen.
   4. Pinpoint ileocaecal junction and make a cut at this level as distal as possible to separate the small intestine from the caecum. Throughout the next steps, avoid excessive physical contact with the intestinal wall because fragile PPs collapse easily upon touch.
   5. Gently remove the entire small intestine until the pyloric sphincter by cutting the mesentery using scissors. Identify the junction between pylorus and duodenum, and snip the duodenum at this level, which will lead to complete detachment of small intestine from the abdominal cavity.
      NOTE: (i) Avoid hyperextension as this might cause the rupture of the intestinal wall. (ii) If LP lymphocyte isolation is desired in addition to PP lymphocytes, complete removal of the mesenteric fat is necessary. However, for PP isolation only, remaining mesenteric fat could provide some benefits during the further isolation steps; therefore, it should be preserved.
   6. Place detached small intestines in a 6-well plate filled with cold RPMI + 10% fetal bovine serum (FBS) and gently agitate the tissues manually until all intestinal segments are submerged in the cold media. Maintain the tissues on ice throughout the next steps.
   7. After dissecting the desired number of mouse small intestines, proceed to PP excision from collected small intestines.
2. Surgical Excision of PPs and Preparation of Single Cell Suspension:
   1. Gently transfer the small intestine on a paper towel by gripping the mesenteric fat using forceps and place the mesenteric side facing the paper towel. Moisten the entire intestinal segment with cold RPMI + 10% FBS to avoid tissue dehydration and stickiness.
      NOTE: Remaining mesenteric fat can be helpful at this stage because fat tissue segments on the mesenteric site of SI will stick to the paper towel keeping the anti-mesenteric site facing up.
   2. Identify the PPs, which appear as white multi-lobulated aggregates in a “cauliflower-like” shape on the anti-mesenteric side of the intestinal wall.
      NOTE: Flushing out the luminal content is not recommended until all PPs are excised. Emptying the luminal content might cause the collapse of the PPs and will prevent the color contrast between PPs and the intestinal wall, which is very helpful for visual identification of PPs.
   3. After identifying PPs on the anti-mesenteric side, place the surgical curved-end scissor on PPs (curve should face up) restraining the PP from its distal and proximal border.
      Optional: Push the PP gently toward the blades of scissor using a fingertip. This maneuver will lead to the better exclusion of surrounding non-PP tissue. Excise the PPs gently, excluding the surrounding intestinal tissue.
      NOTE: (i) This step is crucial to obtain maximal PP cell yield while minimizing the cell contamination from neighboring intestinal compartments such as LP and intestinal epithelium, which are also rich in T cells. (ii) From one SI excised from C57BL/6 mouse, 5-10 PPs (average size, multi-lobulated) can be collected. By aiming even smaller PPs (not multi-lobulated), collection of up to 12-13 PPs per mouse (C57BL/6) is possible using this protocol.
   4. Transfer the excised PPs to a 12-well tissue culture plate filled with ice-cold RPMI + 10% FBS and maintained on ice using forceps or curved surgical scissors.
      NOTE: (i) Immediately after excision of PPs, mucus and intestinal content on the PP surface can be cleaned by rubbing the tissue gently on a paper towel. This step will help improve the viability of PP lymphocytes. (ii) Instead of transferring the PPs to a plate, transferring to separated tubes can also be considered depending on total sample number.
   5. Optional: When PP excision and subsequent placement into the well plate are completed, the number and size of PPs collected from different experimental/mouse groups can be documented by taking a picture of the tissue culture plate containing PPs.
   6. Prepare a set of 50 mL conical tubes filled with 25 mL of RPMI + 10% FBS (pre-warmed at 37 °C). Using a pair of scissors, cut the edge of a 1,000 µL tip from the distance that will allow the aspiration of the PPs with 1 mL pipette. Aspirate the PPs with 1 mL pipette and transfer them from the 12-well tissue culture plate to the prepared 50 mL conical tubes.
      NOTE: Use a new tip for each mouse sample to avoid cross-contamination among the samples.
   7. Secure the lid and place the tubes vertically in an orbital shaker at 37 °C, with continuous agitation at 125–150 rpm for 10 min. Meanwhile, prepare a new set of 50 mL tubes and place a 40 µm cell strainer on the top of each tube, through which single cell suspension will be prepared.
      NOTE: (i) The agitation step will remove the remaining intestinal content, mucus and cell debris, which decrease the cell viability and recovery of PP lymphocytes if not removed. (ii) Do not apply any kind of digestive enzymes on PP tissue because the digestion process causes a dramatic loss of CXCR5 expression from the cell surface.
   8. After the agitation, transfer the PPs to the 40 µm cell strainer placed on the top of the newly prepared conical 50 mL tubes. Using the rounded side of a 10 mL syringe plunger, gently disrupt the PPs through the cell strainer to generate single cell suspension. Rinse the strainer with 15–20 mL of cold RPMI + 10 % FBS.
      NOTE: (i) Before filtering, shake the tubes containing the PPs horizontally. This short shake will facilitate the transfer of PPs into cell strainers. (ii) Using a 70 µm cell strainer for the isolation of non-lymphoid immune cells (e.g., monocytes, macrophages, DCs) is recommended.
   9. Centrifuge the single cell suspensions at 350–400 x g for 10 min at 4 °C.
   10. Carefully discard the supernatant and resuspend the cells at a concentration of 10 x 10⁶ cells/mL. Count the cells using a hemocytometer.
       NOTE: (i) Prior to cell counting, the total cell number for each mouse PP group can be approximately estimated using the following formula: “0.5-1 x 10⁶ cells x (number of PPs) = total cell count”. Further dilutions with trypan blue for cell counting might be necessary. (ii) As an alternative to manual counting, automated cell counters can be used.
   11. Transfer 2-2.5 x 10⁶ cells in appropriate volume (e.g., 200 µL) into a 96-well round-bottom plate.
       NOTE: For single color and negative control samples, 0.5-1 x 10⁶ cells might be sufficient.
   12. Centrifuge the plate at 350 x g for 5 min at 4 °C. Flick the plate.
   13. Wash the cells in 200 µL of Staining Buffer.

3. Surface Antibody Staining

1. Viability Staining:
   1. After the final wash, resuspend the cells in 100 μL of fixable viability dye diluted in PBS (1:1,000). Incubate for 30 min on ice or at 4 °C in the dark.
      NOTE: (i) Intracellular staining for the detection of key transcription factors such as Foxp3 or Bcl-6 requires fixation of the cells. In that case, non-fixable viability dyes (e.g., 7-AAD, DAPI) cannot be used. (ii) To dilute fixable viability dye, do not use any staining buffer that contains protein. The media used in this step must be protein-free. (iii) Exclusion of dead cells by using viability staining is crucial because dead cells can cause serious technical difficulties in flow cytometry analysis by emitting autofluorescence and by binding surface antibodies nonspecifically, which might lead to false positive results.
   2. Wash the cells twice with staining buffer. Centrifuge at 350 x g for 5 min at 4 °C. Flick the plate.
2. Fc Block and Surface -Layer I:
   1. Prepare the Fc-block solution by diluting anti-CD16/32 antibody (1:200) in staining buffer.
   2. Resuspend the cells in 20 μL of prepared Fc-block solution. Incubate for 15 min on ice.
   3. Without washing, add 80 μL of surface antibody cocktail (see Table 1 for the antibody summary) prepared at appropriate dilutions. Incubate on ice for at least 30 min.
   4. Wash twice by adding excessive staining buffer. Centrifuge at 350 x g for 5 min at 4 °C. Flick the plate.
3. Surface -Layer II:
   1. Prepare Streptavidin staining solution by diluting fluorochrome-conjugated Streptavidin in staining buffer.
   2. After the final wash, resuspend the cells with 100 μL of pre-diluted Streptavidin (1:100) staining solution. Incubate for at least 15 min on ice in the dark.
   3. Wash twice with staining buffer. Centrifuge at 350 x g for 5 min at 4°C. Flick the plate.
      Optional: If intracellular staining for follicular regulatory T cell detection is not desired, after the last wash, resuspend the cells in 200 μL of staining buffer and transfer into appropriate tubes to acquire data in flow cytometer. When the samples are not fixed, acquisition of data by flow cytometry should be performed within 3–4 h to obtain accurate results.

4. Cell Fixation

1. Prepare fixation/permeabilization (Fix/Perm) working solution using the reagents from Foxp3/Transcription Factor Staining Buffer Set. Mix one-part of fixation/permeabilization concentrate with three parts of fixation/permeabilization diluent to the desired final volume.
2. After the final wash, resuspend the cells in 200 μL of Fix/Perm working solution.
3. Incubate the plate on ice or at 4 °C for 20 min. Do not exceed 20 min for this step. Longer incubation time might severely decrease the cell recovery.
4. Centrifuge at 350 x g for 5 min at 4 °C. Flick the plate. (Optional) After this step, fixed cells can be stored for several days at 4 °C in staining buffer containing bovine serum albumin (BSA) or FBS until subsequent intracellular staining or flow cytometry acquisition.
5. Resuspend the cells in 200 μL of permeabilization Buffer (x1) freshly pre-diluted in purified deionized water.
6. Centrifuge at 350 x g for 5 min at room temperature (RT).
7. Wash once with 200 μL of permeabilization buffer and centrifuge at 350 x g for 5 min at RT.

5. Intracellular Staining

1. Prepare the Fc-block solution by diluting anti-CD16/32 antibody (1:200) in permeabilization buffer.
   NOTE: After the fixation step, the cells must be maintained in permeabilization buffer until the end of the intracellular staining process.
2. After the final wash, resuspend the cells in 20 μL of Fc-block solution. Incubate for 10–15 min at RT in the dark.
3. Without washing, add 80 μL of intracellular antibody cocktail (100 μL final volume) pre-diluted in permeabilization buffer. Incubate for 30 min at RT.
4. Add 100 μL of Perm Buffer, and centrifuge at 350 x g for 5 min at RT.
5. Wash once with 200 μL of permeabilization buffer, centrifuge at 350 x g for 5 min at RT.
6. After the final wash, resuspend the cells in 200 μL of staining buffer and transfer the cells into appropriate tubes (final volume of 400 μL in staining buffer) and acquire data by flow cytometry. Samples stained in 96-well plate may also be run without transferring into tubes if a flow cytometer with plate reader option is available. This step minimizes cell loss during cell transferring.
7. Acquire a minimum of 5 x 10⁵ total cells on the flow cytometer to be able to perform reproducible analysis of TFH, TFR as well as GC B cells.

Results

In contrast to a previous protocol²⁰, we have observed that PPs are not evenly distributed throughout the SI but are localized more densely towards the distal and proximal ends of the SI as shown in Figure 1A. Flow cytometric analysis showed that, if followed correctly, our protocol gives a PP lymphocyte population that demonstrates forward-side scatter distribution similar to splenocytes (

Discussion

Here, we describe a protocol optimized for flow cytometric characterization of TFH and GC B cells. One of the major advantages of our protocol is that it enables the isolation of up to 10⁷ (average 4–5 x 10⁶ cells) total PP cells from a single mouse (C57BL/6 strain) without any digestive process. We observed that the total cell yield was positively correlated with the number of PPs and could be estimated from the following simple equation which is helpful for experimental planning: "total ...

Materials

Name	Company	Catalog Number	Comments
anti-mouse CD4 antibody	eBioscience, Biolegend*	17-0041-81 ,10054*	For detailed information see Table 1
anti-mouse CD19 antibody	eBioscience	MA5-16536	For detailed information see Table 1
anti-mouse PD-1 antibody	eBioscience	61-9985-82	For detailed information see Table 1
anti-mouse ICOS antibody	eBioscience	12-9942-82	For detailed information see Table 1
anti-mouse GL7 antibody	Biolegend	144610	For detailed information see Table 1
anti-mouse CXCR5 antibody	Biolegend*, BD Bioscience	145512*, 551960	For detailed information see Table 1
anti-mouse BCL-6 antibody	Biolegend	358512	For detailed information see Table 1
anti-mouse Foxp3 antibody	eBioscience	17-5773-82	For detailed information see Table 1
Streptavidin-BV421	BD Bioscience	563259	For detailed information see Table 1
FixableViability Dye	eBioscience	L34957	For detailed information see Table 1
7AAD	Biolegend	420404	For detailed information see Table 1
FcBlock (CD16/32)	BD Bioscience	553141	For detailed information see Table 1
Collagenase II	Worthington	LS004176
Collagenase IV	Worthington	LS004188
Foxp3/Transcription Factor Staining Buffer Set	eBioscience	00-5523-00
6-well,12-well & 96-well plates	Falcon/Corning	353046,353043/3596
50 ml conical tubes	Falcon	3520
40 µm cell strainer	Falcon	352340
10 ml syringe-plunger	Exel INT	26265
RPMI	Corning	15-040-CV
PBS	Corning	21-040-CM
FBS	Atlanta Biologicals	S11150
Orbital shaker	VWR	Model 200
Curved-end scissor
Fine Serrated Forceps
Small curved scissor