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. Author manuscript; available in PMC: 2023 Jul 12.
Published in final edited form as: Mucosal Immunol. 2023 Jan 16;16(1):17–26. doi: 10.1016/j.mucimm.2022.11.004

Tissue-resident memory T cells trigger rapid exudation and local antibody accumulation

Pamela C Rosato 1,2, Sahar Lotfi-Emran 1, Vineet Joag 1, Sathi Wijeyesinghe 1, Clare F Quarnstrom 1, Hanna N Degefu 2, Rebecca Nedellec 3, Jason M Schenkel 1, Lalit K Beura 1,4, Lars Hangartner 3, Dennis R Burton 3,5, David Masopust 1,✉
PMCID: PMC10338064  NIHMSID: NIHMS1909456  PMID: 36657662

Abstract

Adaptive immunity is didactically partitioned into humoral and cell-mediated effector mechanisms, which may imply that each arm is separate and does not function together. Here, we report that the activation of CD8+ resident memory T cells (TRM) in nonlymphoid tissues triggers vascular permeability, which facilitates rapid distribution of serum antibodies into local tissues. TRM reactivation was associated with transcriptional upregulation of antiviral signaling pathways as well as Fc receptors and components of the complement cascade. Effects were local, but evidence is presented that TRM in brain and reproductive mucosa are both competent to induce rapid antibody exudation. TRM reactivation in the mouse female genital tract increased local concentrations of virus-specific neutralizing antibodies, including anti-vesicular stomatitis virus, and passively transferred anti-HIV antibodies. We showed that this response was sufficient to increase the efficacy of ex vivo vesicular stomatitis virus neutralization. These results indicate that CD8+ TRM antigen recognition can enhance local humoral immunity.

INTRODUCTION

Cell-mediated immunity, comprised principally of CD8+ and CD4+ T cells, controls intracellular infections and tumor cells via mechanisms requiring cell-to-cell contact. Humoral immunity, comprised of B cells and antibody, controls infectious agents within the extracellular environment. Cell-mediated and humoral immunity are generally conceptualized as two separate arms of the immune system; however, there are examples of crosstalk. For example, some evidence indicates that CD8+ T cells may be able to execute antibody-dependent cell-mediated cytotoxicity1. Moreover, recent vaccine studies indicate that cell-mediated immunity may enhance protective immunity against vaginal simian immunodeficiency virus (SIV) or simian-HIV (SHIV) challenge in the presence of systemic humoral immunity2,3. Whether enhanced protection resulted from two completely independent mechanisms of protection or synergy between antibody and T cells was not investigated.

Tissue-resident memory T cells (TRM) stably patrol regional compartments without recirculating through blood or other organs and are abundant in mice and humans4–9. When established CD8+ TRM re-encounter cognate antigen, they not only kill target cells but also communicate antigen recognition to other local parenchymal and hematopoietic cells, inducing a local anti-pathogen and immunostimulatory state10–13. This response can protect against pathogen re-encounter, which has been studied extensively at barrier sites such as the skin13–16, and mucosal tissues including the lung17–25, and female reproductive tract10,26–29. The ability of TRM to orchestrate immune responses that span innate and adaptive immunity is one of many examples by which the immune system shares information between cell types. Antibody, complement proteins, and other immune effectors patrol vascular compartments and are largely dependent on transudation from serum to accumulate within tissues30,31. Neutralization of many pathogens, SIV or HIV, for example, at mucosal barrier sites, requires high local antibody concentrations that are difficult to durably establish through vaccination2,32–35. Whether tissue-resident T cells can trigger the local exudation of serum immune effector molecules is unknown.

Here, we describe a new function for CD8+ TRM in mediating local increases in vascular permeability within hours of reactivation, resulting in extravasation of circulating antibody. This increase in antibody afforded heightened neutralization of virus ex vivo. These data indicate a form of synergy between CD8+ T cells and humoral immunity and provide one rationale for including both arms of the immune system in vaccine design against certain intracellular pathogens in cases where antibody alone may not be locally available and durably protective.

RESULTS

To study the impact of TRM reactivation in the female reproductive tract (FRT), which, in this study, includes both the uterus and vagina unless noted, and gain insights into potential mechanisms of humoral crosstalk, we established a trackable population of virus-specific memory T cells, taking advantage of transgenic P14 T cells specific for the gp33 epitope of lymphocytic choriomeningitis virus (LCMV). We transferred a small number of naïve Thy1.1 or CD45.1 P14 cells into CD45.2 congenically distinct wild-type C57BL/6 host mice. One day after T cell transfer, we infected mice with LCMV via the intraperitoneal route, which results in the durable establishment of broadly distributed memory T cells, including TRM in the FRT (Fig. 1A)10,11,36. These mice will be referred to as P14 immune chimeras. To first understand the transcriptional changes that occur locally in the FRT when TRM re-encounter antigen, we took a reductionist approach to trigger TRM reactivation in the absence of viral reinfection by delivering gp33 peptide or an irrelevant control peptide (SIINFEKL) transcervically. This was performed 120 days after infection. To gain an understanding of the early transcriptional changes that occur in the FRT following TRM reactivation, we performed a microarray on whole FRT tissue 9 hours following gp33 peptide or control SIINFEKL peptide delivery. We observed significant upregulation of pro-inflammatory transcripts, including cytokines, genes important for T cell chemotaxis and tissue migration, and antiviral responses, including interferon (IFN)-stimulated genes (Fig. 1B and 1C). This is consistent with what has been reported for TRM responses in the skin12,13 and demonstrates durability of TRM sensing and alarm functions. These data also validated our previously published studies in the FRT, which found an upregulation of VCAM-1 and CXCL9 on endothelial cells, CCL2 and CXCL9 on DCs, and CCL3/CCL4 and IFN-γ on T cells following TRM reactivation10,11. In probing this data set, we noted upregulation of genes associated with antibody effector functions, including components of the complement cascade, Fc gamma receptors, as well as the polymeric immunoglobulin (Ig) receptor (PIGR) (Fig. 1D). Levels of PIGR have been reported to change with estrus cycle37. As we did not note the stage of estrus cycle for microarray experiments, we wanted to confirm the upregulation of PIGR was independent of cycle and thus treated mice with medroxyprogesterone acetate (Depo-Provera) to synchronize mice in diestrus. Six hours following TRM reactivation with transcervically delivered peptide, we found an increase in PIGR expression in the uterus by immunofluorescence, consistent with the microarray results (Fig. 1E and F).

Fig. 1.

Fig. 1

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TRM reactivation induces immune-activating transcriptional changes. (A) Schematic of experimental set-up. P14 immune chimeras 120 days post-infection were challenged transcervically with gp33 or control SIINFEKL peptide and microarray was performed 9 hours later; (B) Volcano plot of differentially expressed genes in FRT where TRM were recalled versus control; (C) Top 100 differentially expressed genes categorized by function; (D) Fc receptor and complement gene expression in FRT tissue where TRM were recalled versus control. Data from n = 3 mice in each group; experiment performed once; (E) Representative PIGR staining in the uterine horns of Depo-Provera treated P14 immune chimera mice 35 days post-infection transcervically challenged with gp33 or control SIINFEKL peptide. Tissue taken 6 hours after peptide challenge. PIGR in magenta, DAPI in blue. < scale bar = 500 μm>; (F) quantification of E (n = 5 mice for TRM recall and n = 6 for control). Data are pooled from two independent experiments. Statistical significance was determined by unpaired two-tailed Mann–Whitney test where *p < 0.05. Error bars indicate standard error of mean. DAPI = 4′,6-diamidino-2-phenylindole dihydrochloride; LCMV = lymphocytic choriomeningitis virus; PIGR = polymeric immunoglobulin receptor; TRM = Tissue-resident memory T cells; WT = wild-type.

Given this increase in antibody effector and transcytosis pathways, we next tested if TRM can trigger an increase in antibody into tissue. To do this, we repurposed a well-established technique to distinguish between T cells in the vasculature and tissue parenchyma by using an intravenous (IV)-injected anti-CD8α antibody prior to animal euthanasia (Fig. 2A). This method is routinely used to selectively stain T cells in the vasculature while sparing cells in the tissue due to the short time (antibody is injected only a few minutes before cell isolation) and restrictive nature of vascular barriers38–40. We hypothesized that TRM reactivation would promote access of IV-injected antibody into tissue, which would manifest as intermediate staining of tissue-restricted CD8+ T cells. To assess this, we quantified the mean fluorescent intensity (MFI) of IV-injected, labeled anti-CD8α antibody on CD8β+ T cells from the FRT following TRM reactivation. Consistent with our hypothesis, we found that there was a significantly higher MFI of a CD8α IV-negative fraction in tissues when TRM were reactivated (Fig. 2A). This suggested that TRM reactivation promoted infiltration of IV-transferred antibody.

Fig. 2.

Fig. 2

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TRM reactivation induces changes in IV antibody staining in non-lymphoid tissue. (A) P14 immune chimeras were challenged transcervically with gp33 peptide (n = 6 mice) or control peptide (SIINFEKL; n = 6 mice) then 6 hours later biotinylated anti-CD8α antibody was transferred intravenously and mice were euthanized after 3 minutes. Representative flow cytometry plots from FRT tissue. Cells gated on CD8β +. Quantification of mean fluorescent intensity of IV-negative stained cells (gate indicated in histogram); (B) Levels of IgG2c, IgA, and IgM in the vaginal lumen of P14 immune chimera mice, stratified by estrus cycle (Di; n = 34 mice), (Pro; n = 13 mice), (Est; n = 16 mice), (Met; n = 6 mice); (C) Experiment was designed as in (A) except CD8α antibody was injected intravenously 12 hours after intracranial delivered peptides. Representative flow cytometry plots from brain tissue. Cells gated on CD8β +. Quantification of mean fluorescent intensity of IV-negative stained cells (gate indicated in histogram) with n = 4 for control and n = 4 for TRM recall. All data are pooled from at least two independent experiments. Statistical significance was determined by unpaired two-tailed t test in all instances except the IgM panel in (B) where an unpaired two-tailed Mann–Whitney was performed (due to non-normal distribution of the data) where * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001. Error bars indicate standard error of mean. Di = diestrus; Est = estrus; FRT = female reproductive tract; LCMV = lymphocytic choriomeningitis virus; Ig = immunoglobulin; i.p. = intraperitoneal; IV = intravenous; Met = metestrus; Pro = proestrus; TRM = Tissue-resident memory T cells; WT = wild-type.

To test if endogenous antibody levels were increased upon TRM reactivation in the FRT, we first performed enzyme-linked immunosorbent assays (ELISAs) for antibody on washes from the vaginal lumen of P14 immune chimeras. Steady-state, endogenous antibody levels in the FRT were widely variable and closely associated with estrus cycle (Fig. 2B), consistent with other reports41. Because of this, we wanted to confirm in an alternative tissue site that TRM reactivation leads to increased IV antibody access. We chose to assess this question in the brain because it is surrounded by a highly restrictive blood–brain barrier. We established a population of brain TRM by transferring P14 cells and infecting mice with LCMV intranasally instead of intraperitoneally because the intranasal infection route establishes a higher magnitude of brain TRM (data not shown). At a memory timepoint, we delivered gp33 or control peptide intracranially and 12 hours later, injected IV anti-CD8α antibody, and euthanized mice 3 minutes later, as before. Like the FRT, we found that the MFI of tissue-restricted IV-negative cells was significantly higher, confirming that antibody gains access to tissue when TRM are reactivated (Fig. 2C).

Given the variability in antibody levels with estrus cycle, we synchronized the cycles by treating mice with Depo-Provera to test if endogenous antibody increased upon TRM reactivation in the FRT. Six to 12 days later, when mice were in diestrus, we reactivated P14 TRM with gp33 or control peptide delivered transcervically. Six hours after TRM recall, we performed ELISAs on vaginal washes (Fig. 3B) and whole FRT tissue lysate (Fig. 3C) for different Ig isotypes. While the amount of antibody varied, we found that TRM reactivation led to a significant increase in IgA in the vaginal lumen and IgG1, IgG2b, and IgG2c in FRT tissue. Moreover, although differences were undetectable by ELISA, we found a significant increase in IgM staining in uterine tissue by immunofluorescence (Fig. 3D and 3E). These data also suggest that TRM reactivation in the uterus can have distal effects on IgA in the vagina.

Fig. 3.

Fig. 3

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TRM reactivation triggers rapid increase in antibody in the FRT. (A) Depo-Provera treated P14 chimeras were challenged with gp33 or control peptide transcervically and then 6 hours later, vaginal washes and FRT tissues were assayed for antibody by enzyme-linked immunosorbent assay; (B) Levels of antibody in vaginal washes 6 hours following TRM reactivation; n = 12 mice for control and n = 14 mice for TRM recall; (C) levels of antibody in FRT tissue 6 hours following TRM reactivation; n = 10 mice for control and n = 12 mice for TRM recall; (D) Representative IgM staining of FRT tissue where green = IgM and blue = DAPI stained nuclei. Scale bar = 500 μm; E: Quantification of D; n = 5 mice. All data are pooled from at least two independent experiments. Statistical significance was determined by unpaired two-tailed Mann–Whitney in (B) and (C) panel IgG2c, or by unpaired two-tailed t test in (E) and (C) panel IgG1 and IgG2b, where * p < 0.05, ** p < 0.01. Error bars indicate standard error of mean. DAPI = 4′,6-diamidino-2-phenylindole dihydrochloride; FRT = female reproductive tract; LCMV = lymphocytic choriomeningitis virus; Ig = immunoglobulin; TRM = Tissue-resident memory T cells.

Due to the observed rapid changes in immunoglobulin levels within 6 hours, we hypothesized that the observed increase in antibody was at least partly due to TRM-triggered vascular permeability. To test this, we performed an ELISA for albumin, which is a protein that is normally found in circulation, in mice treated with Depo-Provera to synchronize estrus cycles in diestrus. Consistent with our hypothesis, the amount of albumin was significantly increased, by approximately 40%, in FRT tissue following TRM reactivation (Fig. 4A). This was validated through IV injection of Evans blue dye (EBD), which binds to serum albumin and is normally excluded from tissue. Six hours following TRM reactivation, as with albumin, we found a significant increase in EBD in FRT tissue (Fig. 4B). To test if this increase in vasodilation permitted an increase in antibody redistribution from the vasculature into the tissue, we injected anti-chicken ovalbumin (OVA) antibody IV into P14 immune chimeras. Two hours following antibody transfer, we reactivated TRM and then measured the amount of anti-OVA antibody in the FRT 6 hours later. We found that TRM reactivation triggered anti-OVA antibody infiltration into the tissue, which was three times the amount of control.

Fig. 4.

Fig. 4

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TRM reactivation triggers vascular permeability and neutralizing ab infiltration into the FRT. (A-C) Levels of albumin (n = 10 mice for control and n = 12 for TRM recall) (A), intravenous transferred EBD (n = 8 mice) (B), and IV-transferred anti-OVA ab (n = 8 mice for control and n = 6 for TRM recall) (C) in FRT tissue 6 hours following transcervical delivery of gp33 or control peptide to Depo-Provera treated P14 immune chimera mice. (D) Levels of PGDM1400 HIV neutralizing ab in the FRT after transferring 5 μm (left; n = 8 mice) or 50 μm (right; n = 8 mice) ab intravenous 2.5–3 hours prior to transcervical delivery of gp33 or control peptide; (E) Plaque reduction neutralization assay against VSVOVA using FRT lysate from Depo-Provera treated P14 immune chimeras that were (right) or were not (left) previously infected with VSVOVA. Data are 6 hours following transcervical delivery of gp33 or control peptide. Non-immune; n = 2 mice for control, experiment performed once, and n = 5 for TRM recall. VSV immune; n = 7 mice for both groups. All data are pooled from at least two independent experiments unless noted. Statistical significance was determined by unpaired two-tailed t test in (A–C), unpaired two-tailed Mann–Whitney in (D), and a two-way analysis of variance with Sidak’s multiple comparisons test (E) where * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001. Error bars indicate standard error of mean. ab = antibody; aOVA = anti-chicken ovalbumin; EBD = evans blue dye; FRT = female reproductive tract; TRM = Tissue-resident memory T cells; VSV = virus expressing ova.

These data raise the possibility that TRM activation may enhance antibody-mediated protection from infection. We recently reported that non-human primates that had low antibody titers following vaccination against SHIV were vulnerable to SHIV challenge compared to animals with high antibody titers2. However, if animals with lower titers had also received a T cell vaccine that established SHIV-specific TRM in the FRT, they were protected against SHIV challenge2. Given these data, we wished to evaluate if TRM could synergize with antibody by triggering rapid re-localization of protective antiviral antibody into the FRT. As a proof-of-concept test, we transferred the HIV broadly neutralizing antibody, PGDM140042, IV into Depo-Provera treated P14 immune chimera mice and delivered gp33 or control peptide transcervically to reactivate TRM. Six hours following TRM reactivation, we tested for levels of PGDM1400 in FRT tissue by ELISA and saw a significant twofold increase in antibody titers (Fig. 4D). To test if increases in local antibody triggered by TRM reactivation could contribute to viral neutralization, we infected P14 immune chimeras with vesicular stomatitis virus expressing ova (VSVOVA) to establish VSVOVA-specific humoral immunity. After viral clearance (30+ days), we reactivated LCMV-specific TRM through transcervical gp33 peptide delivery to Depo-Provera treated mice and 6 hours later, purified protein from the FRT tissue lysate. With this, we performed a plaque reduction neutralization test with VSVOVA and found that tissue extract from FRT where LCMV-specific TRM were reactivated could neutralize VSVOVA to a significantly greater extent (p = 0.014) than tissues that received control peptide. Since VSV is rapidly inhibited by type-I IFN, we performed the same assay with FRTs from P14 immune chimeras that were naïve to VSVOVA to confirm that TRM-triggered IFN responses were not responsible for ex vivo viral neutralization. Consistent with a role for antibody neutralization rather than a generalized antiviral response in vivo, we observed minimal ex vivo neutralization in mice with no VSVOVA-specific immunity regardless of prior TRM reactivation (Fig. 4E).

DISCUSSION

Although vaccine approaches inducing humoral immunity have a history of success against many pathogens, attempts have fallen short for several infections such as malaria, tuberculosis, and those infecting reproductive mucosa such as HIV and herpes simplex virus (HSV), indicating novel approaches are needed43–46. Here, we show that CD8+ resident memory T cells can trigger rapid changes in FRT tissue following reactivation, including pathways that have been observed in other tissues (IFN-stimulated genes, cytokines/chemokines)12,13, and those involving antibody effector functions. Furthermore, we showed that CD8+ TRM-triggered exudation of neutralizing antibody into the tissue through increased vascular permeability. We found this increase in vascular antibody transudation to be independent of B cells, as levels of IV-delivered antibody increased in the FRT following TRM recall. These data have implications for vaccine design and suggest targeting both cellular and humoral immunity may be optimal for protective vaccines. Indeed, studies examining immune correlates of protection against COVID-19 indicate an important role for both T and B cells47–49.

A previous report demonstrated that IFN-γ produced by circulating memory CD4+ T cells triggered increases in vascular permeability in the central nervous system resulting in antibody infiltration 6 days following viral rechallenge50. Here, we show that this vasodilation function extends to resident memory CD8+ T cells, which respond rapidly (within hours rather than days), allowing immediate infiltration of circulating antibody locally into the FRT. This correlated with ex vivo viral neutralization. We explored the requirement for IFN-γ; however, blocking IFN-γ with an antibody had no effect on vascular permeability as measured by EBD (unpublished). We also did not observe an effect when we blocked tumor necrosis factor-α, suggesting there may alternative or synergistic mechanisms of TRM-induced vascular permeability such as IFN-γ and tumor necrosis factor-α combined, or recruitment of histamine or bradykinin-producing basophils or mast cells. This is an area of great interest that warrants future studies. These findings suggest vaccines aimed at establishing both antibody responses and TRM may heighten protective immunity in the FRT.

Although we took a reductionist approach to interrogate TRM functions using viral peptide, it will be important in future studies to understand how this heightened protection is achieved with in vivo viral challenges. By performing passive antibody transfers, we have demonstrated that CD8+ TRM reactivation rapidly increases bloodborne antibody to local tissue. Experimentally, it will be challenging to isolate the in vivo effects of increased antibody transudation from myriad other outcomes of local TRM reactivation that could contribute to protection, including the induction of innate antiviral pathways (Fig. 1)12,13. Moreover, TRM have been shown to recruit B cells, which may eventually contribute to increases in local antibody titers as well10,51. Indeed, this has been demonstrated for CD4 + TRM in the context of HSV infection in the vagina, where antibody increases were seen as early as 24 hours, which were attributed to B cell recruitment into the tissue upon HSV rechallenge of immunized mice51. As we see increases in complement effectors and Fc receptors transcriptionally (Fig. 1) and natural killer cell activation10,11, it will be interesting to see how TRM can affect additional humoral effector mechanisms, including complement activation, antibody-dependent cellular cytotoxicity, and opsonization. Moreover, we noted that levels of IgM were seen to increase by immunofluorescence but not by ELISA. Although we did not observe co-localization of IgM with the B cell marker B220, it may be that increases in IgM+ B cells contribute to this increase or that the ELISA assay was not sensitive enough to detect differences seen by immunofluorescence.

In summary, this study describes a novel function for CD8+ TRM in mediating vascular permeability and antibody extravasation into tissue. These data suggest that antibody-based vaccines may synergize with T-cell vaccines and that targeting both arms of adaptive immunity may provide superior protection against mucosal pathogens.

METHODS

Mice

C57BL/6J (B6) female mice were purchased from The Jackson Laboratory (Bar Harbor, ME) and were maintained in specific-pathogen-free conditions at the University of Minnesota or Dartmouth College. Thy1.1 + P14 and CD45.1 OT-I mice were fully backcrossed to C57BL/6J mice and maintained in our animal colony. Sample size was chosen based on previous experience. No sample exclusion criteria were applied. No method of randomization was used during group allocation, and investigators were not blinded. All mice used in experiments were 6–25 weeks of age. All mice were used in accordance with the Institutional Animal Care and Use Committees guidelines at the University of Minnesota and Dartmouth College.

Adoptive transfers and infections

Immune memory mice were generated by transferring 5 × 104 Thy1.1 + P14 CD8+ T cells from female mice into naïve 6–8 week-old C57BL/6J female mice. One day following transfer, mice were infected with 5 × 105 plaque-forming units LCMV Armstrong in 200μl intraperitoneally. For experiments with VSVOVA, 30 days post LCMV infection, mice received 5 × 104 CD45.1 OT-I IV and then were infected the next day with VSVOVA strain Indiana at 105 plaque-forming units intravenously. For intranasal infections, mice were anesthetized with a ketamine/xylazine cocktail and inoculated with 2 x 105 PFU LCMV Armstrong in 30μl PBS intranasally.

Lymphocyte isolation for flow cytometry

We repurposed an intravascular cell labeling technique, as described previously39, to assess availability of antibody to vascular and parenchymal T cells. Briefly, mice were injected intravenously with biotin-conjugated anti-CD8α. Three minutes after the injection, mice were euthanized and tissues were harvested as described10,36. In brief, FRT tissue was removed, digested in Collagenase IV (Sigma) with DNAse for 1 hour, then dissociated via gentleMACS Dissociator (Miltenyi Biotec), and lymphocytes were purified on a 44/67% Percoll (GE Healthcare) gradient. Isolated cells were stained with Steptavidin-BV650 (Biolegend) to label biotin-CD8α bound cells, and antibodies to CD44 (clone IM7, BioLegend), CD45.1 (clone A20, Biolegend), Thy1.1 (Clone OX7, Biolegend), and CD8β (clone Ly-3, Biolegend). All cells were stained at antibody dilutions of 1:100 except Thy1.1, which was 1:1000. Cell viability was determined with Ghost Dye 780 (Tonbo Biosciences). The stained samples were acquired with LSRII or LSR Fortessa flow cytometers (BD Biosciences) and analyzed with FlowJo software (Treestar).

RNA isolation and microarray

FRT tissue was suspended in 1 ml TRIZOL (Invitrogen) and then homogenized with a glass Dounce homogenizer. RNA was then isolated following the TRIZOL recommended protocol. Resulting RNA was then further purified using Qiagen RNA Cleanup Kit. Total RNA samples were processed using the MouseWG-6 v2.0 Expression BeadChip (Illumina). Background correction, normalization, and identification of differentially expressed genes (false discovery rate < 0.05) were performed in R using the limma package.

Immunofluorescence microscopy and quantification

FRT tissue was harvested, then fixed in 1% BD cytofix for 2 hours before being treated with 30% sucrose overnight for cryoprotection. The sucrose-treated tissue was embedded in OCT tissue-freezing medium and frozen in an isopentane liquid bath. Frozen blocks were sectioned into 7 μm sections and stained with antibodies to IgM-FITC (clone II/41, BD Biosciences), and PIGR (clone 7C1, Abcam) conjugated in-house to CF647 (Biotium), and counterstained with 4′,6-diamidino-2-phenylindole dihydrochloride to detect nuclei. Images were captured by confocal microscopy using the Leica Stellaris 8 at 40x final magnification, using a 20x objective (NA = 0.75). Image analysis, cell counting, and calculation of MFI were performed using the spots function in Imaris version 9.8.

Peptide injections

Intracranial:

Intracerebroventricular injections were performed as described previously52. Briefly, 3 μm of control (SIINFEKL), or gp33 peptide (KAVYNFATM) suspended in phosphate-buffered saline (PBS) was injected into the right cerebral ventricle using a Hamilton Syringe, left for 10 seconds, then slowly removed.

Transcervical:

Transcervical injections were done as previously described11. Briefly, 50 μg of gp33 or SIINFEKL control peptide suspended in 30 μl PBS was delivered past the cervix with a gel loading pipette tip and then dispensed. The tip was left for 10 seconds, then slowly removed.

Depo-Provera treatment and estrus cycle determination

Medroxyprogesterone acetate (Depo-Provera) was administered subcutaneously at 100 mg/kg.

Diestrus was confirmed by vaginal cytology 6–12 days later. Estrus cycle staging was also determined by cytology on vaginal washes as described previously53. Briefly, vaginal washes were taken by pipetting 30 μl PBS in and out of the vagina five times. Then, 3 μl of vaginal wash was mounted on a microscope slide with coverslip, and estrus cycle (proestrus, estrus, metestrus, or diestrus) was determined based on the proportion of leukocytes, cornified epithelial, and nucleated epithelial cells.

EBD assay

200 μl of a 1% solution of sterile filtered EBD was injected into the tail vein of mice. Twenty minutes later, mice were thoroughly perfused with 20 ml ice-cold PBS. FRT tissue was removed and incubated in 500 μl formamide overnight at 56 ° C. Tubes were then spun at 2000 × g for 5 minutes, and supernatant was read on a plate reader at 620 nm.

Protein isolation from tissue

FRTs were homogenized in 1.5 ml NP-40 cell lysis buffer (Invitrogen) with protease inhibitor cocktail (Sigma). Detergents were removed via columns (Pierce) then sample was desalted via 7MWCO column (Zeba). The resulting lysate was used for downstream ELISA assays, BCA total protein assay (Pierce), and VSV neutralization assays.

Enzyme-linked immunosorbent assays

Mouse Ig:

Ready-SET-Go mouse Ig ELISA kits (eBioscience) were used per manufacturer’s instruction for all antibody isotypes in vaginal washes and FRT lysate.

Mouse Albumin:

Mouse Albumin ELISA was performed on FRT lysates per manufacturer’s instruction (Immunology Consultants Laboratory, Inc).

PGDM1400:

ELISA to detect PGDM1400 was performed on FRT lysate as previously described42. Briefly, goat-anti-human IgG Fc (Jackson ImmunoResearch) was used to coat 96-well plates overnight. Following washing and blocking with 3% BSA, sample diluted in 1%BSA/0.02% Tween was added and incubated for 1 hour at room temperature. Following washing, detection antibody goat-anti-human IgG Fab’2-Alkaline phosphatase (Jackson ImmunoResearch) was added and incubated for 1 hour. Alkaline phosphatase stain buffer and substrate (Sigma-Aldrich) were added, and plates were read at 10–20 minutes.

Anti-OVA antibody:

96-well plates were coated with OVA protein at a concentration of 10 μg/ml overnight at 4 °C. Plates were washed with 0.05% Tween and blocked with the blocking solution from Ready-SET-Go ELISA kits (eBiosciences). Following washing, sample was incubated with detection antibody goat-anti-mouse IgG HRP (Invitrogen) for 3 hours at room temperature. Following washes, TMB substrate was added and plates read 15 minutes later at 450 nm on a plate reader.

Plaque reduction neutralization assay

Purified FRT lysate free of detergent was heat inactivated for 30 minutes at 56 °C. Samples were diluted in Dulbecco’s Modified Eagle Medium with 1% FBS and twofold serial dilutions were made. Samples were incubated with VSVOVA for 90 minutes at 37 °C, then 200 ul was plated on Vero cell monolayers in 6-well plates and incubated for 1 hour, shaking every 15 minutes. Sample was then removed, and 2 ml of a 0.3% SeaKem Agarose (Lonza) overlay was added. After 24 hours, agarose was removed, and cells stained with 0.1% Crystal Violet. Plaques were counted, and neutralization was determined as the −log2 of the dilution at which the sample can neutralize 50% of infectious virus.

Statistics

Data were subjected to the Shapiro–Wilk normality test to determine whether they were sampled from a Gaussian distribution. If a Gaussian model of sampling was satisfied, parametric tests (unpaired two-tailed Student’s t test for two groups and one-way analysis of variance or two-way analysis of variance with Sidak’s multiple comparison test for more than two groups) were used. If the samples deviated from a Gaussian distribution, non-parametric tests were used (Mann–Whitney U test for two groups, Kruskal–Wallis with Dunn’s multiple comparison test for more than two groups). All statistical analysis was done in GraphPad Prism (GraphPad Software Inc.). A p-value < 0.05 was considered significant.

Cell definitions

TRM:

In this study, we have defined TRM as P14 T cells in the FRT. This is based on our prior data in the same mouse model demonstrating through parabiosis studies > 95% residency36.

Supplementary Material

1

FUNDING

This work was supported by 1R01AI146032 (DM), 3R01AI084913 (DM), 2P20GM113132-06 (PR), and K22AI148508-02 (PR).

Footnotes

DECLARATION OF COMPETING INTEREST

The authors have no competing interests to declare.

APPENDIX A. SUPPLEMENTARY MATERIAL

Supplementary data to this article can be found online at https://doi.org/10.1016/j.mucimm.2022.11.004.

DATA AVAILABILITY

The datasets generated during and/or analyzed during the current study are available from the corresponding author on request. Microarray data is available through the GEO database under the accession code GSE219002.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

1
NIHMS1909456-supplement-1.pdf (96.3KB, pdf)

Data Availability Statement

The datasets generated during and/or analyzed during the current study are available from the corresponding author on request. Microarray data is available through the GEO database under the accession code GSE219002.

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