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Cell Immunol. Author manuscript; available in PMC Mar 11, 2012.
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PMCID: PMC3078036

Distinct Responses of Splenic Dendritic Cell Subsets to infection with Listeria monocytogenes: maturation phenotype, level of infection, and T cell priming capacity ex vivo


To determine the relative contributions of DC subsets in the development of protective immunity to Listeria monocytogenes we examined the relationship between maturation, bacterial burden, and T cell priming capacity of four well characterized subsets of splenic DC following infection with Lm. CD8α+, CD4+, and CD8αCD4 DC and the B220+ plasmacytoid DC (pDC) were compared for abundance and costimulatory molecule expression at 24, 48, and 72h post i.v. infection. We further determined the bacterial burden associated with each DC subset and their relative capacities to prime CD8+ T cells at 24hpi. The CD8α+ DC displayed the highest level of maturation, association with live bacteria, and T cell activation potential. Second, the CD4+ DC were also mature, yet were associated with fewer bacteria, and stimulated T cell proliferation, but not IFN-γ production. The CD8αCD4 DC showed a modest maturation response and were associated with a high number of bacteria, but failed to induce T cell proliferation ex vivo. pDC displayed a strong maturation response, but were not associated with detectable bacteria and also failed to stimulate T cell activation. Finally, we measured the cytokine responses in these subsets and determined that IL-12 was produced predominantly by the CD8+ DC, correlating with the ability of this subset DC to induce IFN-γ production in T cells. We conclude that Listeria-specific CD8+ T cell activation in the spleen is most effectively achieved by infection-induced maturation of the CD8α+ DC subset.

Keywords: dendritic cells, Listeria, T cell activation, costimulation, innate immunity


Dendritic cells play a major role in the activation of adaptive immunity by capturing, processing, and presenting antigen to naïve T cells in lymphoid organs [1]. DC are important for protective immunity to the Gram-positive, intracellular bacterial pathogen, Listeria monocytogenes (Lm) [2; 3]. Infection with sub-lethal doses of wild type Lm induces the maturation of DC in vivo and primes protective T cell responses [4; 5]. In contrast, infection with even high doses of an avirulent mutant Lm that cannot enter the host cell cytoplasm (referred to herein as vacuolar Lm) is not as effective at inducing DC maturation [4; 5; 6], and is, for the most part, ineffective in conferring protection [7; 8; 9; 10; 11; 12]. In fact, new evidence suggests that recognition of Lm that are not able to escape the phagosome actually suppresses protective T cell responses [11].

A range of migratory and resident dendritic cell subsets have been identified in the spleen [13; 14; 15]. The resident, splenic conventional DC (cDC) include CD8α+, CD4+, and CD8αCD4 DC (DNDC). The interferon-producing, B220+ DC are the plasmacytoid DC (pDC) [13; 16; 17]. CD8α+DC play an important role in the initiation of CD8+ T cell responses and possess potent cross-presentation activity, reviewed in [18]. The expression of 33D1 by CD8α DC is associated with the co-expression of antigen processing machinery for MHC class II presentation and thus the ability to preferentially present antigen to CD4+ T cells [19]. Finally, pDC are best known for their production of type I interferon upon viral infection, but their role in antigen presentation is modest [20; 21; 22].

While CD8α+ DC have been predominantly implicated in the priming of the CD8+T cell responses critical to protective immunity to Lm [23; 24; 25], the role of other DC subsets in the immune response to Lm has not been well defined. In particular, little has been done to examine the number maturation state of individual DC subsets in the first 72 hours following infection. Intriguingly, CD8α+ DC have also been shown to be required for establishing Lm infection in the spleen [25]. Therefore, assessing their role in antigen presentation vs. transport of bacteria (and their antigens) into lymphoid organs has been experimentally difficult using approaches to deplete these cells. Furthermore, the relationship between direct infection and antigen presentation/T cell priming remains to be defined.

To address these questions, we sought to determine how distinct DC subsets responded to either wild type or vacuolar Lm in vivo during the first 72hpi. We examined the response (costimulatory molecule expression, number, and level of infection) of four splenic DC subsets and their capacity to activate naïve CD8+ T cells following Lm infection. These findings offer novel insights as to how individual DC subsets contribute to the overall response to an intracellular pathogen, and indicate that optimal T cell priming is achieved by DC that are both mature and infected.

Materials and Methods


Lm strains 10403s (wild type), DP-L2319 (vacuole-retained), DP-L4056 (wt LmOVA), and DP-L4027 (Δhly LmOVA) were obtained from Dr. Daniel Portnoy (University of California, Berkeley, CA). DP-L2319, which harbors deletions in the genes encoding Listeriolysin O (LLO) and two phospholipases (Δhly ΔplcA ΔplcB), herein referred to as “vacuolar,” cannot enter the cytosol of host cells. DP-L4027 has a single deletion of the hly gene that encodes for LLO. DP-L4056 and DP-L4027, which secrete the ovalbumin protein have been described previously [12].


C57BL/6 mice were purchased from Taconic Farms (Germantown, NY) or Charles River Laboratories (Wilmington, MA). OT-I/Rag1−/− TCR transgenic mice specific for OVA257-264 presented by Kb were purchased from The Jackson Laboratory (Bar Harbor, ME). CD11c-DTR-GFP mice which have been previously described [26] were obtained from The Jackson Laboratory (Bar Harbor, ME).

Infection of mice and bacterial enumeration

Mice were infected with 5×104 or 5×105 cfu wild type Lm or LmOVA (10403s or DP-L4056, respectively), 5×108 cfu vacuolar Lm or vacuolar Lm-OVA (DP-L2319 or DP-L4027, respectively), or mock treated (sterile 1X PBS). For bacterial enumeration ex vivo, sorted splenic DC subsets were lysed in water and dilutions plated on BHI agar to assess bacterial counts. Bacterial counts were normalized per 103 DC or per 106 total splenocytes. GraphPad Prism software was used for statistical analysis.

Staining strategy for flow cytometric identification of splenic DC

At the times indicated post infection, spleens were harvested and digested with Collagenase IV (Worthington Chemical) for 30 min followed by red blood cell lysis, to obtain a single cell suspension. Because the majority of splenocytes are T and B cells, we used a combination of antibodies against CD3 and CD19 labeled with the same fluorophore to exclude these cells from our analysis (referred to as a “dump channel”). Anti-CD3 (145-2C11) and anti-CD19 (1D3; both in PerCP Cy5.5), were used in combination with anti-CD11c APC (HL3) to identify all DC subsets in first step gating. To pinpoint specific cDC subsets, we used: anti-CD8α Pacific Blue (53-6.7), and anti-CD4 FITC (3/23). Anti-CD45R/B220 Pacific Blue (RA3-6B2) and anti-Ly6C FITC (AL-21) were used to define pDC. To determine the maturation status of the specific DC subsets, we used one of the following antibodies as an additional stain: anti-CD86 PE (GL1), anti-CD80 PE (16-10A1), anti-CD40 PE (3/23), or anti-I-Ab PE (AF6-120.1), all from BD Pharmingen. After excluding T and B cells, CD8α+ DC were defined as: CD11c+CD8α+CD4; CD4+ DC: CD11c+CD8αCD4+; DNDC: CD11c+CD8αCD4B220; and pDC: CD11c+ B220+ Ly6C+. Cells were then fixed with 2% PFA and acquired using a BD FACSCanto II with Diva software for acquisition and analysis (BD Biosciences).

Sorting of Splenic DC Subsets

Collagenase-digested splenocytes harvested at the indicated timepoints were first enriched for CD11c+ cells using positive selection with anti-CD11c microbeads from Miltenyi-Biotec. Resulting CD11c-enriched cells were sorted into the 4 DC populations using a BD FACS Aria and the following staining protocol: anti-CD3 (145-2C11) and anti-CD19 (1D3; both labeled with PerCP Cy5.5 to exclude contaminating T and B cells), anti-CD11c APC (HL3), anti-CD8α Pacific Blue (53-6.7), anti-CD4 FITC (3/23), and anti-CD45R/B220 PE (RA3-6B2), all from BD Pharmingen.

Ex vivo T cell priming assays with sorted splenic DC

At 24hpi, sorted splenic DC from uninfected, wt LmOVA-infected or Δhly LmOVA-infected mice were seeded at 2.5×104 cells/well in 96-well V-bottom plates (Corning). Chloramphenicol and gentamicin (each 10μg/ml; Sigma) were added to each well, and mock-treated splenic DC of each subset were loaded with 0.1 ng/ml of preprocessed OVA257-264 peptide (SIINFEKL; from Wake Forest University School of Medicine Peptide Synthesis Facility) or 10 μg/mL of preprocessed OVA323-333 peptide (courtesy of Dr. Steven B. Mizel) for 30 min to 1h prior to the addition of T cells. Peptide was not added to DC infected with LmOVA or Δhly LmOVA.

OVA-specific CD8+ T cells were isolated from the spleens of OT-I/Rag1−/− TCR transgenic mice. T cells were labeled with 5 μM CFSE (BD Pharmingen), according to the manufacturer’s instructions, and added to the DC at a ratio of 2:1 (T cell:DC). T cells were harvested 72 h later and restimulated by incubation with OVA257-264 peptide (2 μg/ml) in the presence of Golgi Plug (BD Pharmingen) for 5 h. The T cells were then stained with anti-CD8α PerCP (53-6.7; BD Pharmingen) then fixed and permeabilized for intracellular cytokine staining using the Cytofix/cytoperm kit (BD Pharmingen), according to the manufacturer’s instructions. Cells were stained intracellularly with anti-IFN-γ Ab (BD Pharmingen). Following staining, cells were resuspended in 2% paraformaldehyde (PFA) and stored at 4°C until acquisition and analysis. CFSE proliferation data were analyzed, as previously described [6], using FlowJo software (TreeStar). Division index indicates the average number of divisions of the starting population of cells (accounting for both divided and undivided cells) and the proliferation index quantitates the average number of divisions of the divided cells.

Cytokine detection in DC by intracellular staining

Mice were infected i.v. with 5×104 Lm-OVA for 24h. Spleens were harvested, collagenase digested, and cell suspensions were incubated for 5h in the presence of Golgi Plug (BD Biosciences). Cells were then surface stained to identify DC subsets as indicated above. Cells were then fixed, permebilized and stained with an antibody against IL-12p40 using the Cytofix/cytoperm kit according to manufacturer instructions (BD Biosciences).


The number of CD8α+ and CD4+ splenic DC decreases after 24 hpi following infection with wild type L. monocytogenes

We first wanted to determine how the number and activation state of splenic DC subsets changed over the first 72 of infection with Lm, as this is a key time for T cell priming. To distinguish among the four previously described resident DC subsets reviewed in [27], we used a staining and gating strategy to examine only CD11c+ cells, then further separated the cells based on expression of CD8α, CD4, B220, and Ly6C. This strategy defined populations of DC that were either CD8α+CD4− (CD8α+ Dc), CD4+CD8α (CD4+ DC), CD8αCD4 (double negative, or DNDC) or B220+Ly6C+ (pDC). In supplementary Fig. 1, we have depicted the gating strategy used to include or exclude cells from the subset analysis.

Using this approach, we now demonstrate that the overall number of CD8α+ and CD4+ DC in the spleen decreased significantly after 24 hours of infection with wild type Lm (Fig. 1A). These decreases were on the order of 50-100-fold loss in the CD8α+ DC by 72h, and an approximate 10-fold loss in the CD4+ population by 48h. Yet, no significant decrease was observed in the DNDC and pDC subsets compared to mock infected animals. A significant decrease in the total number of splenocytes was also observed after 48h infection with WT Lm, but this 2-3-fold decrease could not completely account for the more dramatic losses in the CD8α+ and CD4+ DC. In contrast, the number of CD8α+ and CD4+ DC in vacuolar Lm-infected mice remained steady (Fig. 1A), as did the total number of splenocytes following infection with the vacuolar mutant Lm (Fig. 1B). Surprisingly, the numbers of pDC at 48h and 72h and of DNDC at 48h were significantly increased over mock in the mice infected with vacuolar Lm.

Figure 1
Splenic DC subset numbers decrease 24 hours after Listeria infection

Splenic DC subsets mature to different extents following listerial infection

The extent of DC maturation in specific splenic DC subsets has not been well established following i.v. infection. We therefore determined how splenic DC subsets responded to Lm infection in vivo. We also wanted to determine how the DC maturation response compared with the development of protective immunity in vivo, by assessing the maturation of DC following infection with either bacteria that efficiently induce protection (Wt Lm) or a strain that requires a much higher dose to induce protection (vacuolar Lm). We infected C57BL/6 mice with either 5×104 Wt Lm or 5×108 vacuolar Lm and harvested total splenocytes at 12, 24, 48, or 72 hpi. We examined the maturation state of the DC subsets at this time using the multistep gating strategy described in the methods. The expression of CD86, CD80, and CD40 were quantitated based on median fluorescent intensity compared to the level expressed by DC from mock-infected mice.

Among the three subsets of cDC (CD8α+, CD4+, and DNDC) we first examined the CD8α+ DC as these have been reported to be critical for protective immunity to Lm and priming CD8+ T cell responses [3; 23]. We found that these cells strongly up-regulated the expression of CD40, CD80, and CD86 upon infection with wild type Lm (Fig. 2A) by 24 hpi. In contrast, the expression of costimulatory molecules by CD8α+ DC in response to vacuolar Lm was barely above the mock level at all times tested except for an early and transient two-fold increase at 12h. The maturation response of the CD4+ DC elicited by wild type Lm was lower in magnitude (data not shown) but fold induction over basal levels was comparable or slightly greater than the CD8α+ DC (Fig. 2B). However the peak of maturation in this subset occurred at 48 hpi in contrast to the CD8α+ DC, whose maturation peaked closer to 24h. Again, there was no significant increase in costimulatory molecules on CD4+ DC in response to vacuolar Lm. Interestingly, DNDC did not appear to upregulate CD40 or CD80 to a great extent in response to either wild type Lm or vacuolar Lm over DC from mock-infected animals, but did modestly up-regulate CD86 at 24, 48, and 72hpi (Fig. 2C). There was also a small, but significant up-regulation of CD86 by these DNDC in response to vacuolar Lm. Finally, we observed a strong maturation response in the pDC population following exposure to wild type Lm (Fig. 2D) that peaked around 24 hpi. While the level of costimulatory molecule expression on the splenic pDC was substantially lower than the cDC on a per-cell basis (data not shown), the fold induction over background was comparable. Thus, following infection with Wt Lm, we observed different extents of maturation exhibited by the four DC subsets examined and distinct kinetics with which co-stimulatory molecule expression was up- or down-regulated. However, in all DC subsets, the response to vacuolar Lm was weak and transient or virtually non-existent.

Figure 2
Costimulatory molecule expression of splenic DC subsets is largely dependent upon cytosolic entry by Lm and peaks at 24 hours post infection

CD8α+ DC and DNDC had the highest bacterial burdens at 24 hours post infection

Our timecourse analysis of splenic DC numbers and maturation status revealed different patterns of response to infection among the four subsets examined. Thus, we next wanted to assess the level of direct infection of each DC subset ex vivo to determine if infection level correlated with maturation level. For these experiments, we infected groups of C57BL/6 mice i.v. with wild type or vacuolar Lm and measured the numbers of viable bacteria associated with each DC subset in the spleen 24h post infection, (when the peak of CD8α+ DC and pDC maturation occurs). Following infection, spleens were harvested and cDC and pDC selected by cell sorting. The sorted cells were then lysed and lysates plated on BHI agar to measure the number of viable bacteria associated with each subset. CFU in the liver were also enumerated to ensure that mice had similar overall bacterial burden at 24 hpi (Supp. Fig. 2).

In Fig. 3A and B, the data indicate that the CD8α+ DC, DNDC, and CD4+ DC were infected with low, but detectable numbers of bacteria, but we were not able to detect live bacteria associated with the pDC. To address which subsets were infected with the vacuolar Lm, mice were infected with 5×108 cfu vacuolar bacteria and the level of infection determined in splenic cDC and pDC (Fig. 3A). In a pattern similar to the wild type Lm, we observed a low level of infection in each of the cDC subsets but no detectable vacuolar bacteria in the pDC. Thus, although we did observe significant maturation of pDC in response to vacuolar Lm, we could not detect infection of these cells. Because each DC subset constitutes a different proportion of the total splenocyte population (DNDC make up a larger proportion than CD8α+ DC or CD4+ DC), we also normalized the number of cfu to the number of splenocytes. When calculated based on number of bacteria in each subset per million splenocytes, we observed that there were no significant differences between the number of viable bacteria associated with CD8α+ DC vs. DNDC (Fig. 3B). Finally, the CD4+ DC had fewer bacteria than either CD8α+ DC or DNDC per total splenocytes. Thus, when considering the spleen as a whole, the majority of the Lm were associated primarily with the DNDC and CD8α+ DC, but the CD8α+ DC had a higher number of bacteria per cell.

Figure 3
Splenic CD8α+ and DNDC both contain live Lm following wild-type infection

CD8α+ DC and CD4+ DC prime naïve OVA specific CD8+ T cells ex vivo

To test the ability of sorted splenic DC subsets to prime naïve OVA-specific T cells, following infection with either wild type Lm-OVA or vacuolar Lm-OVA, we performed T cell priming assays ex vivo. Representative dot plots of proliferation and IFNγ production are shown in Figure 4A. Compiled data quantitating the relative proliferative responses are presented in figures 4B and 4C. We observed that both the CD8α+ DC and, to a lesser extent, CD4+ DC were able to induce OVA specific CD8+ T cell proliferation (Fig. 4A and B). The CD8α+ DC promoted T cells to undergo several rounds of proliferation and to produce high levels of IFNγ. In contrast, the CD4+ DC were also capable of inducing proliferation but to a much lesser extent than the CD8α+ DC and strikingly, CD4+ DC did not activate the T cells to produce substantial IFNγ. Neither the DNDC nor the B220+ pDC induced significant proliferation of the OVA specific CD8+ T cells nor their production of IFNγ. Finally, none of the four DC subsets sorted from mice infected with vacuolar Lm stimulated either significant T cell proliferation or IFNγ production. Intriguingly, all four DC subsets stimulated T cell proliferation and IFNγ production when loaded with exogenous OVA peptide in vitro. Thus, the distinct T cell activation capacities of DC subsets ex vivo likely stemmed from differences in the relative levels of antigen presented. We also wanted to determine which of the four splenic DC subsets could prime CD4+ T cell responses ex vivo following Lm infection i.v. Surprisingly, we were unable to detect activation of OVA-specific CD4+ T cells (OT-II) by any of the DC subsets tested (data not shown).

Figure 4
Splenic CD8α+ DC prime naïve, antigen-specific CD8+ T cell proliferation and IFN-γ production ex vivo and is dependent upon cytosolic entry by Lm

To address the possibility that the level of T cell function induced by the different DC subsets correlated with the relative levels of IL-12 produced by these cells, we measured the frequency of IL-12 producing CD8+ and CD4+ DC following 24h Lm infection in vivo by intracellular cytokine staining. As in the previous experiments, we first excluded T and B cells (CD3+ or CD19+) then examined only the CD11c+ cells among the remaining population. The expression of IL-12 vs. CD8α by the total CD11c+ DC is illustrated in figure 5A. We observed that in Lm-infected mice, the majority of IL-12 production was detected in the CD8+ population both in terms of frequency of IL-12-expressing cells as well as in the amount of cytokine expressed per cell. We also consistently observed a low frequency of IL-12 producing cells in this DC subset even in the absence of infection (mock, figure 5A). However, when we specifically compared IL-12 production by CD8+ vs. CD4+ DC, we found that while both subsets responded to infection by up-regulating IL-12, a much lower frequency of CD4+ DC produced IL-12 in response to Lm infection than CD8+ DC (Figure 5B). Thus, these data indicate that the relative capacity of DC subsets to prime IFN-γ production by T cells (Figure 4) correlates with the frequency and level of IL-12 produced by these cells.

Figure 5
IL-12 production by CD8+ and CD4+ DC at 24h post infection


To our knowledge, this is the first study to evaluate simultaneously the maturation profile, infection level, and T cell priming potency of splenic DC subsets upon systemic infection with Listeria monocytogenes. Previous studies have explored specific aspects of this process, such as how DC numbers and maturation profile change at later times post infection [28], or have measured the T cell priming capacity of specific DC subsets [23], yet, none has comprehensively examined these responses with regard to extent of infection in each subset during the peak of T cell priming activity. Taken together, our data demonstrate that upon infection with Listeria, the priming capacity of individual DC subsets depends on both the maturation state of the cells, their association with bacteria, and correlates with the level of IL-12 produced.

Our study demonstrates that the CD8α+ DC are major players in the initiation of protective T cell responses to Lm because 1) they were the most heavily infected (on a per-cell basis) of the four DC subsets examined, 2) displayed a strong maturation response to infection, and 3) were the most potent at stimulating T cell proliferation and function ex vivo. We also observed that the kinetics of the CD8α+ DC maturation was more transient than the CD4+ DC (Fig 2). This may be due to the high infection level in this population, resulting in death of these cells after 24h. Consistent with this notion, our data in Fig. 1 demonstrate that the greatest loss in number of DC was observed in this population at 48 and 72hpi. To explore the possibility that these DC were dying by apoptosis or by pyroptosis, we examined caspase-3 and caspase-1 activation in these cells at 24h post infection. However, we observed no significant difference in the levels of these markers of cell death in DC from infected vs. uninfected mice (data not shown). Thus, the mechanism through which these cells are depleted remains to be determined.

CD8α+ DC have been identified in several studies as being key to the development of protective immunity to a number of infectious organisms [23; 25; 29; 30]. Several studies have employed the CD11c-DTR model for depleting DC to assess the role of DC, (including CD8α+ DC) in establishing protective immunity to Lm [31; 32]. However, these studies revealed that in the Listeria model, CD8α+ DC are also required for seeding of the spleen with bacteria [25]. Thus, distinguishing between the role of DC in T cell activation vs. transport of bacteria has been difficult.

Intriguingly, we also observed a strong maturation response of the CD4+ DC upon infection. These DC were also infected, although at a lower level than the CD8α+ DC. One potential explanation for the failure of CD4+ DC to induce IFNγ production by CTL is that they don’t express sufficient IL-12 (Fig 5) or other Th1 polarizing cytokines [33]. Data in support of this idea have come from several studies of DC responses in which the majority of IL-12 producing DC are found within the CD8α+ subset [34; 35; 36; 37; 38]. An alternative explanation for these findings is that CD4+ DC are not thought to be optimal for priming CTL responses because their antigen processing machinery favors presentation of antigen via MHC class II [39]. However, we were unable to detect any stimulation of CD4+ T cells in our priming assays ex vivo, likely due to a low level of antigen presented at the doses used for infection. Thus, the role of CD4+ DC in the priming of protective T cell responses to Listeria remains to be determined.

The DNDC represent a more heterogeneous subset of cells that expresses CD11c, as well as CD11b (data not shown) but none of the other subset markers examined here. These are also the most numerous of the DC examined. While these cells were associated with a high proportion of the total bacteria, on a per-cell basis they had fewer bacteria than the CD8α+ DC and thus, were not preferentially infected. Additionally, these DC displayed only a modest maturation response, in the form of CD86 upregulation alone. This may be due to the differentiation state of these cells. DC with this phenotype have been identified in the spleen and are thought to be the precursors of several cDC subsets (pre cDC) [17; 40; 41]. Thus, their ability to mature fully in response to bacterial infection may not be completely developed. DC lacking CD8 and CD4 have also been associated with suppressor activity in cancer as well as infectious model systems [42; 43; 44; 45]. While it is not clear if the DNDC in our studies had inhibitory activity, it is likely that the failure of this population to prime CTL activation likely stems from their poor maturation level, in spite of their association with bacteria.

While pDC did strongly upregulate costimulatory molecule expression in response to wild type Lm, their association with viable bacteria was below the level of detection. These findings indicate that either pDC are capable of “bystander” maturation without direct infection, or perhaps pDC are associated with very low numbers of Lm. Because we were unable to detect T cell activation by these cells following Lm infection, we favor the hypothesis that these cells were not directly associated with bacteria, and did not present bacterial antigens. This is not surprising given that pDC are not highly phagocytic and likely unable to take up intact bacteria [20]. However, cross-presentation capacity has recently been reported in pDC [20; 21]. As a control, we determined that when splenic pDC were given pre-processed peptide antigen they were indeed capable of priming naïve, antigen specific CD8+ T cells (Fig. 4).

Perhaps as a consequence of their inability to induce DC maturation, vacuolar Listeria are poor inducers of protective T cell responses in vivo [4-12]. This failure to induce maturation may also affect the localization of the bacteria within the lymphoid organs. One study that addressed this issue [4] demonstrated that while wild type bacteria localized to the T cell zones of the spleen following infection, both killed Lm and Δhly Lm were absent from the T cell zones and instead localized to the red pulp and marginal zones. This finding has been confirmed and extended in three more recent publications examining the formation of innate immune cell clusters in the T cell zones of the spleen following i.v. Lm infection [32; 46; 47]. They observed that Lm were transported by DC into the T cell zones of the spleen, but this transport occurred only upon infection with WT Lm, not vacuolar Lm. Thus, the level of infection and T cell priming capacity of a given DC subset likely depends on its access to bacteria as well as its ability to gain access to the T cell zone.

In summary, our findings provide a clear illustration that individual DC subsets have distinct roles in initiating immunity against Lm. Our study indicates that activation of T cells following the first 24h of infection depends on the DC having both infection and a strong maturation response. This study further provides important insights for targeting DC subsets in the design of vaccines against intracellular pathogens and invites further investigation into the impact of individual DC subsets on protective immune responses to other infectious organisms.

Supplementary Material


Supplementary Figure 1. Splenic DC subset gating strategy. Whole splenocytes were stained for CD3 and CD19 to exclude T and B cells, CD11c to delineate dendritic cells, then CD8α and CD4, or B220 and Ly6C to delineate CD8α+ DC, CD4+ DC, DNDC and pDC (B220+Ly6C+). Cells were excluded for dead, autofluorescent cells and gated on (A and C) CD3 and CD19 cells. (B) Cells then gated on CD8α+ (bottom right gate), CD4+ (top left gate), or CD8αCD4 (bottom left gated) or in a separate gating strategy (D) B220+Ly6C+.


Supplementary Figure 2. Bacterial cfu in Liver. Livers were harvested from mice infected with 5×104 wild type Lm, 5×108 cfu vacuolar Lm, or mock treated with sterile PBS for 12-72h and weighed. Livers then homogenized in sterile water and homogenate serially diluted and plated on BHI for 48h at 37°C. Colony forming units per gram of homogenate then calculated. Vacuolar Lm depicted with open circles. Wild type Lm depicted with black diamonds. Students t test was used to determine significance between vacuolar Lm and wild type cfu.


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[1] Steinman RM. Dendritic cells: understanding immunogenicity. Eur J Immunol. 2007;37(Suppl 1):S53–60. [PubMed]
[2] Jung S. In vivo depletion of CD11c+ dendritic cells abrogates priming of CD8 T cells by exogenous cell-associated antigens. Immunity. 2002;17:211–220. [PMC free article] [PubMed]
[3] Zammit DJ, Cauley LS, Pham QM, Lefrancois L. Dendritic cells maximize the memory CD8 T cell response to infection. Immunity. 2005;22:561–70. [PMC free article] [PubMed]
[4] Muraille E, Giannino R, Guirnalda P, Leiner I, Jung S, Pamer EG, Lauvau G. Distinct in vivo dendritic cell activation by live versus killed Listeria monocytogenes. Eur J Immunol. 2005;35:1463–71. [PubMed]
[5] Feng H, Zhang D, Palliser D, Zhu P, Cai S, Schlesinger A, Maliszewski L, Lieberman J. Listeria-infected myeloid dendritic cells produce IFN-beta, priming T cell activation. J Immunol. 2005;175:421–32. [PubMed]
[6] Brzoza KL, Rockel AB, Hiltbold EM. Cytoplasmic entry of Listeria monocytogenes enhances dendritic cell maturation and T cell differentiation and function. J Immunol. 2004;173:2641–51. [PubMed]
[7] Berche P, Gaillard JL, Sansonetti PJ. Intracellular growth of Listeria monocytogenes as a prerequisite for in vivo induction of T cell-mediated immunity. J Immunol. 1987;138:2266–71. [PubMed]
[8] Barry RA, Bouwer HG, Portnoy DA, Hinrichs DJ. Pathogenicity and immunogenicity of Listeria monocytogenes small-plaque mutants defective for intracellular growth and cell-to-cell spread. Infect Immun. 1992;60:1625–32. [PMC free article] [PubMed]
[9] Pamer EG. Immune responses to Listeria monocytogenes. Nat Rev Immunol. 2004;4:812–23. [PubMed]
[10] Hamilton SE, Badovinac VP, Khanolkar A, Harty JT. Listeriolysin O-deficient Listeria monocytogenes as a vaccine delivery vehicle: antigen-specific CD8 T cell priming and protective immunity. J Immunol. 2006;177:4012–20. [PubMed]
[11] Bahjat KS, Meyer-Morse N, Lemmens EE, Shugart JA, Dubensky TW, Brockstedt DG, Portnoy DA. Suppression of cell-mediated immunity following recognition of phagosome-confined bacteria. PLoS Pathog. 2009;5:e1000568. [PMC free article] [PubMed]
[12] Bahjat KS, Liu W, Lemmens EE, Schoenberger SP, Portnoy DA, Dubensky TW, Jr., Brockstedt DG. Cytosolic entry controls CD8+-T-cell potency during bacterial infection. Infect Immun. 2006;74:6387–97. [PMC free article] [PubMed]
[13] Anjuere F, Martin P, Ferrero I, Fraga ML, del Hoyo GM, Wright N, Ardavin C. Definition of Dendritic Cell Subpopulations Present in the Spleen, Peyer’s Patches, Lymph Nodes, and Skin of the Mouse. Blood. 1999;93:590–598. [PubMed]
[14] Henri S, Vremec D, Kamath A, Waithman J, Williams S, Benoist C, Burnham K, Saeland S, Handman E, Shortman K. The dendritic cell populations of mouse lymph nodes. J Immunol. 2001;167:741–8. [PubMed]
[15] Kamath AT, Pooley J, O’Keeffe MA, Vremec D, Zhan Y, Lew AM, D’Amico A, Wu L, Tough DF, Shortman K. The Development, Maturation, and Turnover Rate of Mouse Spleen Dendritic Cell Populations. J Immunol. 2000;165:6762–6770. [PubMed]
[16] Ardavin C. Origin, Precursors and Differentiation of Mouse Dendritic Cells. Nat Rev Immunol. 2003;3:582–591. [PubMed]
[17] Liu Y-J. IPC: Professional Type 1 Interferon-Producing Cells and Plasmacytoid Dendritic Cell Precursors. Annual Review of Immunology. 2005;23:275–306. [PubMed]
[18] Shortman K, Heath WR. The CD8+ dendritic cell subset. Immunol Rev. 234:18–31. [PubMed]
[19] Dudziak D, Kamphorst AO, Heidkamp GF, Buchholz VR, Trumpfheller C, Yamazaki S, Cheong C, Liu K, Lee HW, Park CG, Steinman RM, Nussenzweig MC. Differential antigen processing by dendritic cell subsets in vivo. Science. 2007;315:107–11. [PubMed]
[20] Villadangos JA, Young L. Antigen-presentation properties of plasmacytoid dendritic cells. Immunity. 2008;29:352–61. [PubMed]
[21] Lui G, Manches O, Angel J, Molens JP, Chaperot L, Plumas J. Plasmacytoid dendritic cells capture and cross-present viral antigens from influenza-virus exposed cells. PLoS One. 2009;4:e7111. [PMC free article] [PubMed]
[22] Gray RC, Kuchtey J, Harding CV. CpG-B ODNs potently induce low levels of IFN-alphabeta and induce IFN-alphabeta-dependent MHC-I cross-presentation in DCs as effectively as CpG-A and CpG-C ODNs. J Leukoc Biol. 2007;81:1075–85. [PubMed]
[23] Belz GT, Shortman K, Bevan MJ, Heath WR. CD8alpha+ dendritic cells selectively present MHC class I-restricted noncytolytic viral and intracellular bacterial antigens in vivo. J Immunol. 2005;175:196–200. [PMC free article] [PubMed]
[24] Neuenhahn M, Busch DH. Unique functions of splenic CD8alpha+ dendritic cells during infection with intracellular pathogens. Immunol Lett. 2007;114:66–72. [PubMed]
[25] Neuenhahn M, Kerksiek KM, Nauerth M, Suhre MH, Schiemann M, Gebhardt FE, Stemberger C, Panthel K, Schroder S, Chakraborty T, Jung S, Hochrein H, Russmann H, Brocker T, Busch DH. CD8alpha+ dendritic cells are required for efficient entry of Listeria monocytogenes into the spleen. Immunity. 2006;25:619–30. [PubMed]
[26] Probst HC, Tschannen K, Odermatt B, Schwendener R, Zinkernagel RM, Van Den Broek M. Histological analysis of CD11c-DTR/GFP mice after in vivo depletion of dendritic cells. Clin Exp Immunol. 2005;141:398–404. [PMC free article] [PubMed]
[27] Shortman K, Naik SH. Steady-state and inflammatory dendritic-cell development. Nat Rev Immunol. 2007;7:19–30. [PubMed]
[28] Tam MA, Wick MJ. Differential expansion, activation and effector functions of conventional and plasmacytoid dendritic cells in mouse tissues transiently infected with Listeria monocytogenes. Cellular Microbiology. 2006;8:1172–1187. [PubMed]
[29] Reinicke AT, Omilusik KD, Basha G, Jefferies WA. Dendritic cell cross-priming is essential for immune responses to Listeria monocytogenes. PLoS One. 2009;4:e7210. [PMC free article] [PubMed]
[30] Kerksiek KM, Niedergang F, Chavrier P, Busch DH, Brocker T. Selective Rac1 inhibition in dendritic cells diminishes apoptotic cell uptake and cross-presentation in vivo. Blood. 2005;105:742–9. [PubMed]
[31] Jung S, Unutmaz D, Wong P, Sano G, De los Santos K, Sparwasser T, Wu S, Vuthoori S, Ko K, Zavala F, Pamer EG, Littman DR, Lang RA. In vivo depletion of CD11c(+) dendritic cells abrogates priming of CD8(+) T cells by exogenous cell-associated antigens. Immunity. 2002;17:211–20. [PMC free article] [PubMed]
[32] Kang SJ, Liang HE, Reizis B, Locksley RM. Regulation of hierarchical clustering and activation of innate immune cells by dendritic cells. Immunity. 2008;29:819–33. [PMC free article] [PubMed]
[33] Henry CJ, Ornelles DA, Mitchell LM, Brzoza-Lewis KL, Hiltbold EM. IL-12 produced by dendritic cells augments CD8+ T cell activation through the production of the chemokines CCL1 and CCL17. J Immunol. 2008;181:8576–84. [PMC free article] [PubMed]
[34] Farrand KJ, Dickgreber N, Stoitzner P, Ronchese F, Petersen TR, Hermans IF. Langerin+ CD8alpha+ dendritic cells are critical for cross-priming and IL-12 production in response to systemic antigens. J Immunol. 2009;183:7732–42. [PubMed]
[35] la Sala A, He J, Laricchia-Robbio L, Gorini S, Iwasaki A, Braun M, Yap GS, Sher A, Ozato K, Kelsall B. Cholera toxin inhibits IL-12 production and CD8alpha+ dendritic cell differentiation by cAMP-mediated inhibition of IRF8 function. J Exp Med. 2009;206:1227–35. [PMC free article] [PubMed]
[36] Hao X, Kim TS, Braciale TJ. Differential response of respiratory dendritic cell subsets to influenza virus infection. J Virol. 2008;82:4908–19. [PMC free article] [PubMed]
[37] Kim TS, Braciale TJ. Respiratory dendritic cell subsets differ in their capacity to support the induction of virus-specific cytotoxic CD8+ T cell responses. PLoS One. 2009;4:e4204. [PMC free article] [PubMed]
[38] Hochrein H, Shortman K, Vremec D, Scott B, Hertzog P, O’Keeffe M. Differential production of IL-12, IFN-alpha, and IFN-gamma by mouse dendritic cell subsets. J Immunol. 2001;166:5448–55. [PubMed]
[39] Dudziak D, Kamphorst AO, Heidkamp GF, Buchholz VR, Trumpfheller C, Yamazaki S, Cheong C, Liu K, Lee H-W, Park CG, Steinman RM, Nussenzweig MC. Differential Antigen Processing by Dendritic Cell Subsets in Vivo. Science. 2007;315:107–111. [PubMed]
[40] Liu K, Nussenzweig MC. Origin and development of dendritic cells. Immunol Rev. 234:45–54. [PubMed]
[41] Liu K, Victora GD, Schwickert TA, Guermonprez P, Meredith MM, Yao K, Chu FF, Randolph GJ, Rudensky AY, Nussenzweig M. In vivo analysis of dendritic cell development and homeostasis. Science. 2009;324:392–7. [PMC free article] [PubMed]
[42] Nagaraj S, Schrum AG, Cho HI, Celis E, Gabrilovich DI. Mechanism of T cell tolerance induced by myeloid-derived suppressor cells. J Immunol. 184:3106–16. [PMC free article] [PubMed]
[43] Ilarregui JM, Rabinovich GA. Tolerogenic dendritic cells in the control of autoimmune neuroinflammation: an emerging role of protein-glycan interactions. Neuroimmunomodulation. 17:157–60. [PubMed]
[44] Bronte V. Myeloid-derived suppressor cells in inflammation: uncovering cell subsets with enhanced immunosuppressive functions. Eur J Immunol. 2009;39:2670–2. [PubMed]
[45] Dugast AS, Vanhove B. Immune regulation by non-lymphoid cells in transplantation. Clin Exp Immunol. 2009;156:25–34. [PMC free article] [PubMed]
[46] Aoshi T, Carrero JA, Konjufca V, Koide Y, Unanue ER, Miller MJ. The cellular niche of Listeria monocytogenes infection changes rapidly in the spleen. Eur J Immunol. 2009;39:417–25. [PMC free article] [PubMed]
[47] Aoshi T, Zinselmeyer BH, Konjufca V, Lynch JN, Zhang X, Koide Y, Miller MJ. Bacterial entry to the splenic white pulp initiates antigen presentation to CD8+ T cells. Immunity. 2008;29:476–86. [PubMed]
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