Reduced T Cell Priming in Microbially Experienced “Dirty” Mice Results from Limited IL-27 Production by XCR1⁺ Dendritic Cells

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Multiple studies over the past quarter century have rigorously defined the T cell response to vaccination, providing exquisite detail to the processes essential for the development and maintenance of memory T cells that provide protection to subsequent challenges (1–3). Although most vaccination research falls under the “infectious disease umbrella,” there has been interest in recent years in using vaccination to stimulate antitumor immunity (4, 5), especially with the identification of tumor-specific immunogenic neoantigens. Current vaccines contain attenuated microorganisms (i.e., bacteria or viruses), purified proteins or toxins, synthetic peptides, or mRNAs that encode for a defined protein to induce protective immunity (6). Vaccines also frequently include an adjuvant, which is designed to increase the activity of APCs and potency of the immunogen (7).

A variety of immunological techniques and reagents, including the extensive array of genetically altered mice, have given researchers the means to interrogate how stimulation through the TCR shapes an Ag-specific T cell response (i.e., T cell expansion, contraction, differentiation, and function) after vaccination or infection at the cellular and molecular levels. Recently, there has been growing interest in understanding how environmental microbial exposure impacts the ability of the immune system to respond to new challenges (8–10). Justification for these studies stems from the fact that the human immune system is shaped and trained by the vaccinations received and pathogenic and commensal microbes encountered daily from birth. In contrast, nearly all laboratory studies use mice housed in specific pathogen-free (SPF) conditions, which keeps their immune systems in a more naive state (8, 11). The landmark study by Beura et al. (12) using SPF and microbially experienced “dirty” mice revealed key immunological parallels between SPF and dirty mice to neonatal and adult humans, respectively. Subsequent studies from our laboratory and others have shown the power of using dirty mice to complement experiments done in SPF mice, revealing how prior microbial exposure can better replicate clinical observations (13–16).

The objective of this study was to define how previous microbial exposure impacts naive T cell expansion, differentiation into memory populations, and function after immunization with a defined peptide subunit vaccine. To investigate this question, we used a vaccine platform consisting of synthetic CD8 and/or CD4 T cell epitope peptides, the TLR3 agonist polyinosinic-polycytidylic acid [poly(I:C)], and agonistic anti-CD40 mAb (17), to stimulate a primary Ag-specific CD8 or CD4 T cell response. Our data show this immunization protocol induced dramatically muted T cell immunity in microbially experienced dirty mice compared with SPF mice. The observed differences in T cell expansion, differentiation, and function did not appear to be intrinsic to the Ag-specific T cells. Instead, the attenuated T cell response in dirty mice correlated with reduced IL-27 production from a XCR1⁺ dendritic cell (DC), which is a known predictor of the magnitude of the T cell response after subunit vaccination (18). Overall, the data presented in this article further highlight the impact of microbial exposure on T cell immune responses to new Ag encounters.

Female CD45.2⁺ C57BL/6N (B6; 8–10 wk of age) and CD45.1⁺ B6.SJL mice were purchased from Charles River (Wilmington, MA) and Jackson Laboratories (Bar Harbor, ME), respectively. Female pet store mice were purchased from local pet stores in the Minneapolis-St. Paul, Minnesota metropolitan area. All mice were housed in American Association of Laboratory Animal Care–approved animal facilities at the University of Minnesota (BSL-1 for SPF B6 mice and BSL-3 for cohoused [CoH] B6 and pet store mice). SPF B6 or B6.SJL and pet store mice were CoH at a ratio of 8:1 in large rat cages for 30–60 d to facilitate microbe transfer (19). Mice within each cohousing cohort were randomly assigned to experimental groups. Experimental procedures were approved by the University of Minneapolis Animal Care and Use Committee and performed following the Office of Laboratory Animal Welfare guidelines and Public Health Service Policy on Human Cancer and Use of Laboratory Animals.

Mice were vaccinated i.v. with a mixture of synthetic peptide (50 µg; Vivitide), poly(I:C) (50 µg; Invivogen), and agonistic anti-CD40 mAb (100 µg; BioXcel). We refer to this mixture as “TriVax,” with the specific peptide(s) as a prefix. Alternatively, mice were immunized s.c. with peptide emulsified in CFA (Sigma) or Alhydrogel (Invivogen).

Mice were immunized with OVA₂₅₇/OVA₃₂₃-TriVax 30 d before being challenged with B16-OVA tumor cells (2.5 × 10⁵ cells in 0.1 ml PBS s.c.). Tumor size (mm²) was measured over time with calipers.

K^b-specific tetramers containing B8R_20–27 (TSYKFESV), OVA_257–264 (SIINFEKL), HSV peptide gp B_498–505 (SSIEFARL), Plasmodium ovale glideosome-associated protein 50_40–48 (GAP-50; SQLLNAKYL), and lymphocytic choriomeningitis virus (LCMV) gp_33–41 (KAVYNFATM) were used to identify and quantify Ag-specific CD8 T cells by flow cytometry. Biotinylated peptide:K^b monomers were obtained from the National Institutes of Health Tetramer Core. I-A^b–specific tetramers containing 2W1S (EAWGALANWAVDSA), Ag28m (VEIHRPVPGTA), LCMV gp_66–77 (DIYKGVYQFKSV), or L. monocytogenes LLO_190–201 (NEKYAQAYPNVS) peptides were used to identify and quantify Ag-specific CD4 T cells by flow cytometry. Biotinylated peptide:I-A^b monomers were made into tetramers with streptavidin-PE or streptavidin-allophycocyanin (Prozyme) as previously described (20, 21).

To quantify the number of Ag-specific CD8 or CD4 T cells within the spleens of SPF or CoH mice, we used a tetramer-based enrichment protocol (20) using p:K^b or p:I-A^b tetramers. In brief, spleens were harvested for each mouse analyzed, a single-cell suspension was prepared, and allophycocyanin- and PE-conjugated tetramers were added at a 1:400 dilution in tetramer staining buffer (PBS containing 5% BCS, 2 mM EDTA, 1:50 normal mouse serum, and 1:100 anti-CD16/32 mAb). The cells were incubated in the dark on ice (for p:K^b tetramers) or at room temperature (for p:I-A^b tetramers) for 1 h, followed by a wash in 10 ml cold FACS buffer. The tetramer-stained cells were then resuspended in 0.2 ml FACS buffer, mixed with 0.05 ml of both anti-allophycocyanin and anti-PE mAb-conjugated magnetic microbeads (STEMCELL Technologies), and incubated in the dark on ice for 30 min. The cells were washed and resuspended in 3 ml cold FACS buffer and passed over a magnet (STEMCELL Technologies) to enrich for tetramer-specific cells. The resulting enriched fractions were then stained with a mixture of fluorochrome-labeled mAb (see later). Cell numbers for each sample were determined using AccuCheck Counting Beads (Invitrogen). Samples were then analyzed using an LSR II flow cytometer (BD) and FlowJo software (Tree Star). The percentage of tetramer-positive events was multiplied by the total number of cells in the enriched fraction to calculate the total number of Ag-specific CD8 or CD4 T cells in the spleen.

Peripheral blood leukocytes or splenocytes were evaluated by flow cytometry. RBCs were removed from heparinized whole blood or single-cell splenocyte suspensions using ACK lysis buffer. To assess the expression of cell-surface proteins, we incubated cells with fluorochrome-conjugated mAb at 4°C for 30 min. The cells were then washed with FACS buffer (PBS containing 2% BCS and 0.2% NaN₃). For some experiments, the cells were then fixed with PBS containing 2% paraformaldehyde. In procedures requiring intracellular staining (i.e., to detect transcription factor or intracellular cytokine presence), cells were permeabilized after surface staining using the Foxp3 transcription factor staining kit (Tonbo) for transcription factors or the BD Cytofix/Cytoperm kit (BD Biosciences), stained for 1 h at 4°C with a second set of fluorochrome-conjugated mAbs, and suspended in FACS buffer for acquisition. The fluorochrome-conjugated mAbs used in both surface and intracellular stainings were Horizon V500 CD90.2 (clone 53-2.1; BD Biosciences), Brilliant Violet (BV) 510 and FITC CD3 (clone 17A2; BioLegend), BV421 and BV605 CD4 (clone GK1.5; BioLegend), BV650 CD8 (clone 53-6.7; BioLegend), PE-CD45.1 (clone A20; eBioscience), FITC-CD45.2 (clone 104; eBioscience), AlexaFluor700 CD44 (clone IM7; BioLegend), PE-Cy7 KLRG1 (clone 2F1; eBioscience), PerCP-Cy5.5 CD62L (clone MEL-14; BD Horizon), BV510 CD127 (clone A7R34; BioLegend), allophycocyanin and BV650 IFN-γ (clone XMG1.2; BioLegend), PE-Cy7 IL-2 (clone JES6-5H4; BioLegend), PerCP-Cy5.5 B220 (clone RA3-6B2; eBioscience), PerCP-Cy5.5 CD11b (clone M1/70; eBioscience), PerCP-Cy5.5 CD11c (clone N418; eBioscience), PerCP-Cy5.5 F4/80 (clone BM8; eBioscience), FITC Foxp3 (clone FJK-15S; eBioscience), FITC TNF-α (clone MP6-XT22; eBioscience), PE IL-12p40 (clone C15.6; BioLegend), and PE IL-27p28 (clone MM27-7B1; BioLegend).

Ag-specific CD8 T cell responses were measured after ex vivo peptide stimulation. In brief, spleens were harvested from SPF and CoH mice 30 d after vaccination with B8R-TriVax. Approximately 10⁶ splenocytes were then stimulated with B8R peptide (1 µg/ml) in media containing 3 µg/ml brefeldin A (Tonbo Biosciences). Cells were incubated for 6 h at 37°C. Cells were then stained for surface Abs to identify CD8 T cells and then fixed and permeabilized (BD Biosciences Cytofix/Cytoperm Kit). After permeabilization, cells were stained for intracellular cytokines with anti–IFN-γ and anti–TNF-α mAbs. After staining, cells were resuspended in FACS buffer, and 0.02 ml of counting beads (eBioscience) was added to each sample immediately before acquisition.

In vivo peptide stimulation was used to determine Ag-specific CD4 T cell function by intracellular cytokine production, as previously described (22–24). In brief, SPF and CoH mice were injected i.v. with 100 µg of the 2W1S peptide 28 d after vaccination with 2W1S-TriVax. After 3 h, spleens were harvested in media containing 10 µg/ml brefeldin A. The resulting cell suspensions were first stained with 2W1S:I-A^b tetramer and other mAb to identify Ag-specific CD4 T cells, as described earlier. The cells were then fixed, permeabilized, and stained with anti–IFN-γ, –TNF-α, and –IL-2 mAb. Cells were resuspended in FACS buffer after staining, and 0.02 ml of counting beads (eBioscience) was added to each sample immediately before acquisition.

SPF CD45.1⁺ P14 donor mice were bled, and 5 × 10⁴ P14 T cells were adoptively transferred into SPF or CoH CD45.2⁺ B6 mice by i.v. injection 1 d before gp₃₃-TriVax immunization. Mice were bled on days 3, 7, 14, and 21 after immunization to identify CD45.1⁺ P14 T cells in the circulation by flow cytometry after staining with gp₃₃:K^b tetramers and anti-CD45.1 mAb. Spleens were also collected on day 21 postimmunization to determine the number of CD45.1⁺ P14 T cells and endogenous CD45.2⁺ gp₃₃-specific CD8 T cells using gp₃₃:K^b tetramers, anti-CD45.1, and anti-CD45.2 mAb. Alternatively, 4 × 10⁶ bulk splenocytes from SPF or CoH CD45.1⁺ B6.SJL mice were transferred into SPF or CoH CD45.2⁺ B6 mice by i.v. injection 1 d before B8R-TriVax immunization. Spleens were then collected 7 d later to determine the number of CD45.1⁺ B8R-specific CD8 T cells using B8R:K^b tetramers, anti-CD45.1 mAb, and anti-CD45.2 mAb.

CoH mice received recombinant mouse IL-27p28:EBI3 complex (250 µg/mouse; Sino Biological) i.v. 2 h after B8R/2W1S-TriVax immunization. The frequency and number of B8R-specific CD8 T cells and 2W1S-specific CD4 T cells in the spleen were determined 5 d later by flow cytometry.

Data shown are presented as mean values ± SEM. GraphPad Prism 9 was used for statistical analysis, where statistical significance was determined using two-tailed unpaired, nonparametric Mann–Whitney U test (for two individual groups) or group-wise, one-way ANOVA analyses followed by multiple-testing correction using the Holm–Sidak method, with α = 0.05 and *p < 0.05, **p < 0.01, ***p < 0.005, and ****p < 0.001.

Immunization with peptide epitopes can elicit robust priming of Ag-specific T cell populations that mediate effective protection against pathogen challenge or tumor growth. Moreover, inclusion of TLR and costimulatory receptor agonists, which are thought to mimic the multiple immunostimulatory pathways activated during true pathogen infection, can maximize the T cell response to the peptides. For example, the combination of defined antigenic peptides mixed with the TLR3 agonist poly(I:C) and agonistic anti-CD40 mAb (17), hereafter referred to as “TriVax,” elicits strong, stable T cell responses in SPF mice capable of mediating protection to viral infection or tumor rejection (25, 26). Because of the documented impact microbial exposure has on shaping and training the composition and function of the immune system to new challenges, we used the TriVax vaccination platform to examine the generation of antitumor immunity in SPF and microbially experienced B6 mice that had been CoH with pet store mice. SPF and CoH mice were immunized with OVA₂₅₇- and OVA₃₂₃-TriVax to activate both OVA-specific CD8 and CD4 T cells. Mice were then challenged with B16-OVA tumor cells 28 d later, and tumor growth was monitored over time (Fig. 1A). Consistent with previous data (25), B16-OVA tumors were reduced in size in the TriVax-immunized SPF mice compared with unimmunized SPF mice (Fig. 1B). To our surprise, we saw no difference in the size of the B16-OVA tumors in untreated and TriVax-immunized CoH mice. Multiple reports have demonstrated TriVax immunization elicits protective immunity (25, 27, 28), so we set out to further examine why this vaccination platform failed to suppress tumor growth in CoH mice.

Peptide immunization, especially when combined with poly(I:C) and anti-CD40 mAb, can dramatically increase the number of Ag-specific T cells (25, 27, 28). To examine the impact of microbial experience on naive T cell priming and expansion, we monitored the frequency and number of B8R-specific CD8 T cells or 2W1S-specific CD4 T cells in the blood and spleens of SPF and CoH mice after vaccination with either B8R- or 2W1S-TriVax, respectively (Fig. 2A). Robust expansion of both B8R-specific CD8 T cells and 2W1S-specific CD4 T cells was noted in the blood of SPF mice, peaking on day 8 and followed by a contraction in population size by days 14 and 28 (Fig. 2B, 2C). We were able to detect expansion of these Ag-specific T cell populations in the blood of CoH mice, but not to the extent seen in SPF mice. Splenic analysis of B8R-specific CD8 T cells or 2W1S-specific CD4 T cells at day 28 revealed similar significant reductions in frequency and number in the CoH mice compared with SPF mice (Fig. 2D, 2E). The reduced T cell expansion in CoH mice was not limited to TriVax immunization, because we also noted reduced numbers of Ag-specific CD8 and CD4 T cells in CoH mice immunized with peptide mixed with either CFA or the alum vaccine adjuvant Alhydrogel (Fig. 2F, 2G). Collectively, these data demonstrate that Ag-specific CD8 and CD4 T cell expansion and the number of cells at a memory time point (after contraction) are diminished in CoH mice (compared with SPF mice) after peptide immunization.

CD8 and CD4 T cell proliferation is accompanied by differentiation into effector cells, which is followed by a contraction phase. The T cells that survive contraction establish the memory pool of cells, displaying phenotypic and functional differences from the naive precursor T cells (29–32). The data presented in (Fig. 2 showed significant reductions in Ag-specific CD8 and CD4 T cells at the peak of expansion and at a memory time point (day ≥28) after contraction in TriVax-immunized CoH mice. Thus, we next evaluated the extent of CD8 T cell differentiation in SPF and CoH mice after TriVax immunization (Fig. 3A). CD8 T cell subsets, such as effector memory (T_EM), central memory (T_CM), and long-lived effector cells (LLECs), can be distinguished based on the expression of CD62L and KLRG1 (33). Early (7 d) after TriVax immunization, there was decreased frequency of B8R-specific CD8 T cells among all CD8 T cells in the spleens of CoH mice compared with what was seen in SPF mice (Fig. 3B), but this did not translate to a significant numerical difference. There were, however, notable reductions in the frequency and number of B8R-specific CD8 T cells in CoH mice 30 d after immunization. Interestingly, CoH mice had an increase in memory precursor cell (MPEC) frequency (KLRG1⁻CD127⁺) and a decrease in short-lived effector cell frequency (KLRG1⁺CD127⁻) 7 d after immunization (Fig. 3C, 3D). Further examination of the B8R-specific CD8 T cells via CD62L confirmed a decrease in KLRG1⁺CD62L⁻ and KLRG1⁻CD62L⁻ cells by percentages and numbers of cells from CoH mice at 7 d postimmunization, with an increase in KLRG1⁻CD62L⁺ cells (Fig. 3E–G). Intriguingly, at 30 d post-TriVax, we saw a reduction by percentage and number within the T_CM (CD62L⁺KLRG1⁻) and T_EM (CD62L⁻KLRG1⁻) subsets, and a reduction in number but increase in frequency of the LLEC (CD62L⁻KLRG1⁺) subset (Fig. 3E–G). Interestingly, we also noted the appearance of a CD62L⁺ population with low expression of KLRG1. Thus, at 7 d post-TriVax, the responding CD8 T cells in CoH mice favor MPEC generation, but by memory time points (i.e., 30 d postimmunization) the B8R-specific CD8 T cells are fewer in number and contain a larger proportion of LLECs. We then examined the ability of B8R-specific CD8 T cells to produce effector cytokines after in vitro peptide restimulation (Fig. 3H). Splenocytes from TriVax-immunized CoH mice had a lower percentage of CD44^hi CD8 T cells producing IFN-γ or IFN-γ and TNF-α after restimulation compared with splenocytes from TriVax-immunized SPF mice (Fig. 3I).

Naive CD4 T cells have the potential to differentiate into multiple “helper” populations depending on the cytokines present at the time of activation (34). These subsets can be defined by lineage-specific transcription factor and/or surface protein expression, such as Tbet⁺ Th1, Bcl6⁺CXCR5⁺ follicular helper T cells, or Foxp3⁺ regulatory T cells (35–37). To evaluate how CD4 T cell differentiation in CoH mice after 2W1S-TriVax immunization compares with differentiation in SPF mice, we determined the frequency and number of 2W1S-specific CD4 T cells that expressed Tbet, Bcl6 and CXCR5, or Foxp3⁺ 7 or 28 d after immunization (Fig. 4A). There was no difference in frequency of Tbet⁺, Bcl6⁺CXCR5⁺, or Foxp3⁺ 2W1S-specific CD4 T cells at the day 7 time point, but the frequency of 2W1S-specific CD4 T cells expressing Tbet in CoH mice 28 d after immunization was reduced compared with similarly immunized SPF mice (Fig. 4B, 4C, Supplemental Fig. 1). Moreover, there was a small (but significant) increase in the frequency of Foxp3⁺ 2W1S-specific regulatory CD4 T cells in the CoH mice on day 28. When applying these frequency data to total number of 2W1S-specific CD4 T cells in SPF and CoH mice after 2W1S-TriVax immunization, the CoH mice contained significantly fewer numbers of Tbet⁺, Bcl6⁺CXCR5⁺, and Foxp3⁺ 2W1S-specific CD4 T cells (Fig. 4D). As with the B8R-specific CD8 T cells, we were also interested in assessing the ability of the TriVax-induced 2W1S-specific CD4 T cells in SPF and CoH mice to produce cytokine after peptide restimulation in vivo (Fig. 4E) (24). Data from this assay showed a dramatic reduction in frequency and number of IFN-γ⁺, IFN-γ⁺TNF-α⁺, and IFN-γ⁺TNF-α⁺IL-2⁺ 2W1S-specific CD4 T cells in CoH mice (Fig. 4F). Together, the results in (Figs. 3 and 4 highlight deficiencies in Ag-specific CD8 and CD4 T cell differentiation and percentage/number of cells capable of making cytokines after restimulation in TriVax-immunized CoH mice, in addition to the reduced proliferative capacity shown in (Fig. 2.

One possible explanation for the reduced T cell expansion in the TriVax-immunized CoH mice we considered was the presence of fewer naive Ag-specific precursors, because the number of precursors can strongly influence the extent of expansion and number of memory T cells after contraction (20, 31, 38–40). Thus, we quantitated the number of T cell precursors for the panel of Ag-specific CD8 and CD4 T cells. For each of the five Ag-specific CD8 T cell and four Ag-specific CD4 T cell populations examined, there was no difference in the number of naive precursors in the spleens of SPF and CoH mice (Fig. 5A, 5B). These data suggest the reduced naive T cell expansion seen in CoH after TriVax immunization was not due to fewer numbers of Ag-specific precursors.

We next examined the impact of the host environment on naive T cell expansion by transferring 50,000 naive P14 TCR-transgenic (Tg) CD8 T cells obtained from CD45.1⁺ SPF donor mice into either SPF or CoH CD45.2⁺ recipient mice 1 d before immunizing with gp₃₃-TriVax (Fig. 6A). The frequency and number of CD45.1⁺ P14 CD8 T cells in the blood were determined 3, 7, 14, and 21 d later, as well as the number of CD45.1⁺ P14 CD8 T cells and endogenous CD45.2⁺ gp₃₃-specific CD8 T cells in the spleens 21 d after immunization. Interestingly, P14 T cell expansion on day 7 postimmunization was considerably blunted in the CoH recipients (Fig. 6B, 6C). Moreover, the reduced number of P14 T cell numbers in the spleens of CoH recipient mice (versus SPF recipient mice) on day 21 after immunization mirrored that for the endogenous gp₃₃-specific CD8 T cell populations present in the same SPF and CoH mice (Fig. 6D–F). Because of the inability to cohouse P14 mice, we were unable to perform the desired reciprocal transfers with the P14 T cells. Instead, we transferred 4 × 10⁶ bulk splenocytes from SPF or CoH CD45.1⁺ mice and transferred them to either SPF or CoH CD45.2⁺ recipients 1 d before B8R-TriVax immunization (Fig. 6G). This approach allowed us to use B8R:K^b tetramers to identify the transferred CD45.1⁺ B8R-specific CD8 T cells that responded to immunization. The results clearly showed equivalent expansion of SPF- and CoH-derived CD45.1⁺ B8R-specific CD8 T cells in SPF recipients, whereas the expansion of either donor CD45.1⁺ B8R-specific CD8 T cell population was significantly reduced in CoH recipients (Fig. 6H, 6I). Thus, data in (Figs. 5 and 6 together suggest the differences in T cell expansion/differentiation seen between SPF and CoH mice after TriVax immunization are not the result of intrinsic T cell differences and are instead governed by T cell–extrinsic factors.

Optimal CD8 and CD4 T cell responses require the right combination of signaling pathways activated via Ag stimulation through the TCR, CD80/86-CD28 costimulation, and cytokines (41, 42). To evaluate the potential impact of changes within the DC compartment in CoH mice on the reduced capacity of TriVax immunization to stimulate a naive T cell response, we first determined the number of XCR1⁺ cDC1, SIRPα⁺ cDC2, and Siglec H⁺ plasmacytoid DCs (pDCs) in the spleens of SPF and CoH mice. Interestingly, each of these DC populations was expanded in CoH mice (Fig. 7A). We also assessed CD40, CD80, and CD86 expression on these DC subsets, finding no differences on XCR1⁺ cDC1, increased CD86 geometric mean fluorescence intensity (gMFI) on CoH SIRPα⁺ cDC2, and increased CD80 and CD86 gMFI on CoH Siglec H⁺ pDCs (Fig. 7B–D). Several recent publications from the Kedl laboratory have described the importance of IL-27p28–expressing XCR1⁺ cDC1 cells in defining the magnitude of T cell expansion after TriVax immunization (18, 43). Thus, we next assessed the extent of IL-27p28 production by XCR1⁺ cDC1 cells from TriVax-immunized SPF and CoH mice. We did not detect any IL-27p28–producing XCR1⁺ cDC1 cells in either SPF or CoH mice at baseline, but there was a substantial increase in IL-27p28 production by these cells in SPF mice 6 h after TriVax immunization (Fig. 7E–H). Strikingly, this IL-27p28 production by XCR1⁺ cDC1 was not seen in TriVax-immunized CoH mice. IL-27p28 is part of the IL-12 cytokine family, so we also measured the expression of IL-12p40 by the XCR1⁺ cDC1 cells. In contrast with the lack of IL-27p28–producing XCR1⁺ cDC1 cells in TriVax-immunized CoH mice, we found SPF and CoH mice had similar frequencies and numbers of IL-12p40–expressing XCR1⁺ cDC1 (Fig. 7I, 7J).

We were intrigued by the limited expression of IL-27p28 by the XCR1⁺ cDC1 cells and wondered whether administration of recombinant IL-27p28:EBI3 complex shortly after TriVax immunization would boost T cell expansion in CoH mice. Using a protocol similar to that described by Pennock et al. (44), we saw a significant increase in the frequency and number of B8R-specific CD8 T cells and 2W1S-specific CD4 T cells in CoH mice that were given a single dose of IL-27p28:EBI3 complex 2 h after immunization compared with CoH mice that did not receive exogenous IL-27 (Fig. 8). Together with the data in (Fig. 7, these data suggest the limited capacity of TriVax immunization to expand Ag-specific CD8 T cells in CoH mice is due (in part) to the reduced frequency/number of IL-27–producing XCR1⁺ cDC1, as well as the reduced amount of IL-27 produced on a per-cell basis (based on gMFI). Moreover, these data further support the idea that the defect restricting vaccine-driven T cell expansion in CoH mice is T cell extrinsic.

Vaccines that elicit robust adaptive immune responses using synthetic peptides combined with adjuvants have generated impressive preclinical data inducing immunity against tumors and pathogens. Prophylactic vaccination for infectious disease typically stimulates the production of neutralizing Abs specific for a conserved region of the microbe. Cancer vaccines can be given prophylactically or therapeutically to induce a robust neutralizing Ab response against immunodominant viral Ags (e.g., L1 proteins from human papillomavirus) or a tumor Ag-specific T cell response, respectively. Regardless of the target of the induced immune response, vaccine efficacy relies on many factors, including the route of delivery, adjuvant, and immunogen used. Interest in peptide subunit vaccines (especially for cancer) has increased in recent years because of detailed knowledge of immunogenic tumor Ag and translation of cancer immunology from the bench to the clinic. Yet, objective response rates have been underwhelming in cancer vaccine clinical trials, and few cancer vaccines have progressed to phase III testing (4, 45–47).

Recent work with peptide subunit immunization has shown differing requirements for supporting factors (such as IL-27, IL-15, and Eomes) when comparing priming of vaccine-stimulated T cells versus infection-stimulated T cells (44). However, it is important to recall that most of the preclinical research on peptide subunit immunization has been conducted in SPF mice, which have limited circulating inflammatory cytokines and are more immunologically naive compared with pet shop or wild mice (12, 13, 15, 16). With that in mind, we were interested to determine to what extent the modified immune environment in laboratory mice that had been CoH with pet store mice to induce immune system maturation still supports robust naive T cell responses after peptide subunit immunization. The data presented in this article provide evidence to suggest reduced ability of peptide subunit immunization to prime a naive CD8 or CD4 T cell response in microbially experienced “dirty” mice (compared with traditional SPF-housed laboratory mice) stemming, in part, from reduced IL-27p28 production from XCR1⁺ cDC1 cells. Consequently, antitumor immunity in peptide subunit–immunized dirty mice was less effective in controlling experimental tumor growth compared with that in SPF mice, which is consistent with the limited success of peptide subunit cancer vaccines in the clinical setting.

Over their lifetime, humans are exposed to numerous commensal and pathogenic microbes. This microbial exposure, along with vaccination, results in an adaptive immune system primed to respond rapidly and vigorously against subsequent microbial challenge. In contrast, the immune systems of most laboratory mice used in biomedical research are composed primarily of naive cells, largely resulting from the SPF housing conditions used in most animal facilities (11, 48). There is no question that the implementation of SPF housing has had a positive impact on preclinical research (e.g., improving experimental reproducibility), but data published in recent years showing how the maturation of the immune system in laboratory mice with a normalized microbial experience (i.e., exposure to natural pathogenic and commensal microbes present in the environment or transferred from other mice) can significantly affect responses to new challenges demonstrate the benefit of including animals with a matured immune system (like that seen in adult humans) in preclinical experimentation (12–16). With this in mind, we were very intrigued by the lack of antitumor immunity in CoH mice after peptide subunit immunization (Fig. 1). Interestingly, these data are consistent with the subdued Ab responses seen in sequentially infected or CoH mice after yellow fever vaccine or influenza virus vaccine administration, respectively (14, 49). Subsequent investigation found clear reductions in CD8 and CD4 T cell expansion, differentiation into memory populations, and percentage/number of cells capable of producing cytokine after restimulation in TriVax-immunized CoH mice, even though naive precursor T cell numbers were equivalent to that found in SPF mice. We reasoned the reduced T cell priming in CoH mice could be caused by either T cell–intrinsic or –extrinsic factors. Limited expansion of adoptively transferred TCR-Tg or Ag-specific CD8 T cells obtained from SPF donor mice in CoH recipients pointed to T cell–extrinsic differences that throttled the potential for T cell expansion.

Cohousing laboratory mice with pet store mice results in a constitutive increase in many adaptive and innate immune cell populations (12–16), but the data in (Fig. 7 reveal these mice also have increased numbers of multiple DC subsets in the spleen. In many ways, it makes sense for there to be an increased DC load in CoH mice to deal with the increased amount of microbial Ag presentation needed to prime the various pathogen- and commensal-specific T cell populations needed to clear these microbes. Similarly, the modest (but significant) differences in costimulatory molecule expression on the SIRPα⁺ cDC2 and Siglec H⁺ pDCs from CoH mice should be expected because of the elevated steady-state inflammation present, because proinflammatory cytokines (e.g., IFN-γ and TNF-α) can increase CD80 and CD86 expression on DCs (50). Based on these parameters—more XCR1⁺ cDC1, SIRPα⁺ cDC2, and Siglec H⁺ pDCs in the spleens of CoH mice at baseline and higher costimulatory molecule expression—it would have been reasonable to predict CoH mice would exhibit similar (or even more robust) naive T cell expansion/differentiation/function compared with SPF mice. However, the most profound finding from our DC analyses was the inability of XCR1⁺ cDC1 to make IL-27p28 after TriVax immunization. Work from the Kedl laboratory has identified IL-27 as being key for supporting T cell responses after peptide subunit vaccination (18, 43, 44). T cell dependence on IL-27 is thought to regulate the metabolic state of vaccine-elicited T cells (51), and many factors, including several TLR agonists and IFN-γ, induce the production of IL-27 (52). Interestingly, we have previously noted the reduced abundance of Tlr3 transcripts in splenic adherent myeloid cells, which include DCs (53), from CoH mice compared with SPF mice (13). Altered TLR3 expression could be a means to explain the reduced stimulatory capacity of the poly(I:C)-containing TriVax immunization in CoH mice, but it is important to recall that we also saw reduced T cell expansion in CoH mice immunized with peptide combined with CFA or Alhydrogel (Fig. 2F, 2G). These data further demonstrate the need for additional investigation into why XCR1⁺ cDC1 fails to make IL-27 in CoH mice.

Increasing evidence supports the notion that mice exposed to diverse microbes exhibit substantial changes in the immune response to pathogens. Moreover, these changes in the baseline immune cell activation profiles correlate more closely with human immune profiles than that observed in SPF mice, suggesting the potential use of “dirty” mice as an intermediate preclinical step between traditional SPF mice and human testing. These changes can benefit the host in some cases, but they can result in detrimental outcomes in other settings. Indeed, epidemiological data show increased vaccine efficacy in humans living in more developed parts of the world where microbial encounter/infections are less frequent compared with that observed in developing countries (54–56). Our data suggest caution in the interpretation that peptide subunit/adjuvant vaccines will induce robust T cell responses when only tested preclinically using hosts with a limited microbial history (i.e., SPF mice), and they point to the need for additional analysis of vaccine efficacy in dirty mice that includes an assessment of DC populations presenting the Ag of interest.

We thank the members of our laboratories for technical assistance and helpful discussions. We also thank the University of Minnesota Flow Cytometry Resource Facility, CFI Dirty Mouse Colony, and BSL-3 Program for support.

The authors have no financial conflicts of interest.