Abstract
Sepsis reduces the number and function of memory CD8 T cells within the host, contributing to the long-lasting state of immunoparalysis. Interestingly, the relative susceptibility of memory CD8 T cell subsets to quantitative/qualitative changes differ after cecal ligation and puncture (CLP)–induced sepsis. Compared with circulatory memory CD8 T cells (TCIRCM), moderate sepsis (0–10% mortality) does not result in numerical decline of CD8 tissue-resident memory T cells (TRM), which retain their “sensing and alarm” IFN-γ–mediated effector function. To interrogate this biologically important dichotomy, vaccinia virus–immune C57BL/6 (B6) mice containing CD8 TCIRCM and skin TRM underwent moderate or severe (∼50% mortality) sepsis. Severe sepsis led to increased morbidity and mortality characterized by increased inflammation compared with moderate CLP or sham controls. Severe CLP mice also displayed increased vascular permeability in the ears. Interestingly, skin CD103+ CD8 TRM, detected by i.v. exclusion or two-photon microscopy, underwent apoptosis and subsequent numerical loss following severe sepsis, which was not observed in mice that experienced moderate CLP or sham surgeries. Consequently, severe septic mice showed diminished CD8 T cell–mediated protection to localized skin reinfection. Finally, the relationship between severity of sepsis and demise in circulatory versus tissue-embedded memory CD8 T cell populations was confirmed by examining tumor-infiltrating and nonspecific CD8 T cells in B16 melanoma tumors. Thus, sepsis can differentially affect the presence and function of Ag-specific CD8 T cells that reside inside tissues/tumors depending on the severity of the insult, a notion with direct relevance to sepsis survivors and their ability to mount protective memory CD8 T cell–dependent responses to localized Ag re-encounter.
Introduction
Sepsis presents a tremendous health care challenge throughout the world, accounting for one third of hospital deaths and affecting almost 2 million people in the United States annually (1–3). The impact of sepsis is more pronounced in the developing world, where it may account for up to 40% of the total deaths in some countries (4, 5). The majority (∼75%) of deaths from sepsis usually occur in patients aged 65 and over, but anyone with an uncontrolled local infection can be at risk for developing sepsis (6, 7). Sepsis is classified as the body’s dysregulated immune response to an uncontrolled systemic infection by a bacterial, viral, fungal, or parasitic pathogen. The septic event can occur from systemic infections, with barrier tissues such as the pulmonary, gastrointestinal, and urinary tracts being common sites of original infection. Following the systemic dissemination of a pathogen from the original nidus of infection, the immune system responds with a massive release of pro- and anti-inflammatory cytokines in the blood (8–10). This “cytokine storm” can lead to host tissue damage and is a period of high risk for acute mortality. Most patients survive the initial septic event, largely through the administration of antibiotics and resuscitation measures, but there are long-lasting lesions in the immune system that compromise the future health of the individual (11, 12). Sepsis survivors are at a significant risk for secondary infection, with 20% of discharged septic survivors being readmitted to the hospital within 30 d (primarily) because of secondary infection complications (13). Also, significant increases in morbidity of septic survivors have been observed due to malignancy and infections that would otherwise be controlled by a healthy, normal immune system (14, 15).
Following the resolution of this acute phase of sepsis, lymphopenia ensues, and a state of systemic immune system paralysis follows thereafter. The immunoparalysis phase is characterized by an overall decrease in immune cell number and function, resulting in an increased susceptibility to viral reactivation, secondary infections, and neoplastic malignancy long after the initial septic event (16–18). Using the experimental murine cecal ligation and puncture (CLP) method of sepsis induction, we and others have observed the diminishment of both naive and memory CD4 and CD8 T cell responses (numbers and Ag-dependent and -independent functions), which leave sepsis survivors with increased susceptibility to newly or previously encountered infections (19–22). These data are consistent with those observed in the human population, in which septic survivors have a marked decrease in T cell number and function (23, 24).
Importantly, the sepsis-induced lesions in CD8 T cells have mostly been studied in the circulating T cell compartment (25–27). However, moderate sepsis (0–10% mortality) does not evoke numerical decline of CD8 tissue-embedded resident memory T cells (TRM), which also retain their “sensing and alarm” IFN-γ–mediated effector functions (28). CD8 TRM are generated at the site of original infection and have been found in many organs, such as the skin, lungs, reproductive tract, and gut. They are phenotypically distinct from their circulating counterparts, and they serve to guard and sense local tissue and provide a robust early response upon cognate Ag encounter (29). Recent reports have also showed their egression to secondary lymphoid organs, suggesting a role in priming a recall (or secondary) response (30, 31). Thus, because of their anatomical positioning outside of the vasculature and in previously infected tissue, CD8 TRM are uniquely poised to provide protection and mediate clearance of reinfection. It is also tempting to speculate that TRM are more resistant to the deleterious effects of sepsis because of their seclusion from circulation and sepsis-induced cytokine storm. In this study, we show that depending on the severity of the insult and magnitude and duration of cytokine storm, sepsis has the capacity to also influence tissue- or tumor-embedded bystander memory CD8 T cells, a concept with direct relevance to sepsis survivors and their ability to mount protective memory CD8 T cell–dependent responses to localized/peripheral Ag re-encounter.
Materials and Methods
Mice, pathogens, and memory CD8 T cell generation
Experimental procedures using mice were approved by University of Iowa Animal Care and Use Committee under protocol numbers 6121915 and 9101915. The experiments performed followed Office of Laboratory Animal Welfare guidelines and Public Health Service Policy on Humane Care and Use of Laboratory Animals. Inbred B6 (C57BL/6; Thy 1.2/1.2) mice were purchased from the National Cancer Institute (Frederick, MD) and maintained in the animal facilities at the University of Iowa at the appropriate biosafety level. P14 TCR-transgenic mice (Thy1.1/1.1) were bred and maintained at the University of Iowa (Iowa City, IA). B6.CAG.MRFP1 mice were obtained from The Jackson Laboratory and crossed with B6.P14-Thy1.1 mice at the University of Iowa to yield P14-RFP Thy1.1 B6 mice. P14 mice were crossed with B6 eGFP (The Jackson Laboratory) at the University of Iowa to yield P14-eGFP Thy 1.1 B6 mice.
Lymphocytic choriomeningitis virus (LCMV)–Armstrong (2 × 105 PFU) was injected i.p. Vaccinia virus (VacV)–GP33 infection of the skin was performed by applying 5 × 106 PFU in 5 μl saline to the center of the ear pinna and then poking it 30 times with a 27-gauge needle (32). VacV-GP33 viral titers were quantified using plaque assay on BSC-40 cells, as described previously (33). Naive P14 CD8 T cells obtained from peripheral blood of naive P14 mice (Thy1.1) were adoptively transferred into B6 recipients (Thy1.2; 5 × 103 to 10 × 103 Thy1.1+ P14 T cells per recipient) i.v., followed by infection with a pathogen expressing the gp33-41 epitope of LCMV.
Cell isolation
Peripheral blood was collected by retro-orbital bleeding. Ears, small intestine, and lungs were treated with collagenase type II (100 U/ml; Worthington Biochemical) in RPMI 1640 supplemented with 5% FCS and shaken at 450 rpm for 30–90 min at 37°C. Single-cell suspensions were prepared by mashing tissues through a 70-µm cell strainer (Falcon) with the plunger of a 1-ml syringe (BD Biosciences). Samples were centrifuged and resuspended in RPMI 1640. Single-cell suspensions from spleen, lymph nodes, and tumors were generated after mashing tissue through 70-µm cell strainer without enzymatic digestion.
Flow cytometry, and cytokine detection
Flow cytometry data were acquired on an FACSCanto (BD Biosciences, San Diego, CA) and analyzed with FlowJo software (Tree Star, Ashland, OR). To determine expression of cell surface proteins, mAb were incubated at 4°C for 20–30 min, and cells were fixed using Cytofix/Cytoperm Solution (BD Biosciences) and, in some instances, followed by mAb incubation to detect intracellular proteins. The following mAb clones were used: CD8 (53-6.7; eBioscience), Thy1.1 (HIS51; eBioscience), B220 (RA3-6B2; eBioscience), CD45.2 (104; eBioscience), IFN-γ (XMG1.2; eBioscience), TNF-α (MP6-XT22; eBioscience), IL-2 (JES6-5H4; eBioscience), CD103 (2E7; eBioscience), CD31 (390; BioLegend), CXCL9 (MIG-2F5.5), VCAM-1 (429; eBioscience), CXCR3 (CXCR3-173; eBioscience), CD62L (MEL-14; eBioscience), CD27 (LG.7F9; eBioscience), KLRG-1 (2F1; eBioscience), CD127 (A7R34; eBioscience), CD122 (5H4; eBioscience), CD69 (H1.2F3; eBioscience), T-bet (eBio410; eBioscience), Eomesodermin (Dan11mag; eBioscience), granzyme B (MHGB04; Invitrogen), CD107a (1D4B; BD Pharmingen). IFN-γR was detected using CD119-Biotin (2E2; eBioscience) and streptavidin–PE (eBioscience) after acid stripping, as described previously (34). Plasma concentrations of IFN-γ, IL-1α, IL-1β, IL-2, IL-6, TNF-α, CCL5, IL-10, IL-18, CXCL5, G-CSF, LIF, IL-22, IL-27, CCL11, CXCL1, CXCL10, CCL2, CCL7, CCL3, CCL4, CXCL2, IL-12p70, GM-CSF, and IL-13 were determined using Multiplex Immunoassay (ProcartaPlex; Affymetrix by eBioscience) on a Bio-Rad BioPlex, as performed previously (35). Apoptosis was evaluated using Vybrant FAM Caspase-3 and -7 Assay Kit (Invitrogen) according to manufacturer’s protocol.
Tissue extraction
At varying times after CLP or sham surgery, ears, lungs, and spleens were collected and added to a 1.5-ml centrifuge tube for processing as previously described (36). Each ear was weighed. To each sample was added 0.5 ml of 0.1% Tween 20 in PBS, and samples were ground for 3–5 min with a pellet pestle attached to a cordless electric drill. Samples were then quick frozen in liquid N2, thawed in a 37°C water bath, and ground again for 3–5 min. Samples were sonicated for 15 s and then centrifuged for 5 min at 13,000 × g. Supernatants were then removed for cytokine measurement by multiplex immunoassay. Blood was also collected from these same mice and then allowed to clot. Serum was collected and run on the same multiplex immunoassay.
Intravascular stain protocol to distinguish circulatory from resident cells
Two minutes after APC-conjugated CD45.2 mAb was injected i.v. into mice, peripheral blood was collected by retro-orbital bleeding and used as a positive control with >99% memory P14 CD8 T cells routinely labeled with CD45.2 mAb (37). After one additional minute, mice were euthanized, and organs of interest were harvested and processed for flow cytometry.
Vascular leakage
Mice were injected i.v. with 200 μl of Evans blue dye (1.0% in PBS) at indicated time points. After 30 min, mice were anesthetized, and ears, lungs, and intestines were harvested and treated with formamide. Quantification of the dye was done using OD reading on a spectrophotometer based on a standard curve with known amounts of Evans blue dye.
CLP model of sepsis induction
CLP procedure was performed (38) on mice that were anesthetized with ketamine/xylazine (University of Iowa, Office of Animal Resources). Briefly, the abdomen was shaved and disinfected with Betadine (povidone iodine; Purdue Products), and a midline incision was made. The distal third of the cecum was ligated with PERMA-HAND Silk (Ethicon) and punctured once (“moderate”) or twice (“severe”) using a 25-gauge needle, and a small amount of fecal matter extruded. The cecum was returned to abdomen, the peritoneum was closed with 641G PERMA-HAND Silk (Ethicon), and skin was sealed using surgical Vetbond (3M). Following surgery, 1 ml PBS was administered s.c. to provide postsurgery fluid resuscitation. Bupivacaine (Hospira) was administered at the incision site, and flunixin meglumine (Phoenix) was administered for postoperative analgesia. This procedure created a septic state characterized by loss of appetite and body weight, ruffled hair, shivering, diarrhea, and/or periorbital exudates with 0–10% mortality rate or 25–50% mortality for moderate and severe sepsis, respectively, similar to our previous reports (39). Sham mice underwent identical surgery excluding CLP. Removal of punctured cecum as source control is not attempted in this study because of the short duration of the experiments—most mice were analyzed/sacrificed in the first 2–4 d after CLP surgery.
Bacterial burden
Livers and spleens were obtained from animals 12 h postsurgery, weighed, and homogenized in antibiotic-free RPMI 1640 (Life Technologies) containing 10% FCS (Life Technologies) with a mechanical disruptor. Serial dilutions of the homogenates were performed, and aliquots were plated on 5-cm petri dishes containing antibiotic-free brain heart infusion agar. Plates were incubated at 37°C, 5% CO2 overnight, and colonies were counted by hand the next morning to calculate the bacterial burden per gram of tissue.
Intravital two-photon microscopy of skin TRM
Fluorescent P14 donor mice were bled, and 2 × 103 to 5 × 103 RFP P14 or 5 × 104 GFP P14 T cells were adoptively transferred to mice by i.v. injection 1 d prior to infection. VacV-GP33 (5 × 106 PFU) was administered to the ear (28), and mice underwent CLP or sham surgery at 30–40 d postinfection. Prior to imaging, mouse ears were treated with Nair (Church and Dwight) for 3–5 min to remove hair. Ears were imaged for RFP or GFP P14 T cells with live intravital two-photon microscopy 48 h postsurgery. The VacV-immune ears that contained RFP or GFP P14 TRM cells of live mice were positioned on the microscope base in a continuously heated enclosed chamber (Leica). A custom suction tissue window apparatus (VueBio) was placed on the ear with 20–25 mm Hg of negative pressure to gently immobilize the tissue against a fixed coverslip. All images were acquired on an upright Leica SP8 Microscope (Leica) using a 25×/0.95 NA water immersion objective with coverslip correction with 1.25× zoom. High-resolution (512 × 512) stacks of 15–38 x-y-sections sampled with a resonant scanning head operating with bidirectional scanning and 5 μm z-spacing were acquired to provide image volumes of 354 × 354 × 75–190 μm and voxel sizes of 0.693 × 0.693 × 5 μm. Line averaging of 16 and a kernel-3 median filter were used in Leica Application Suite X to reduce noise. Images were excited with a tunable InSight laser (Spectra-Physics) with excitation at 1040 nm and collected by two tunable internal Hybrid Detectors with emission collected for secondary-harmonic generation at 515–525 nm and at 560–630 nm for RFP P14s. A tunable Mai Tai HP Sapphire laser (Spectra-Physics) at 940 nm was used for detection of eGFP (500–550 nm) and secondary-harmonic generation (435–485 nm) using a quad Hybrid Detector array equipped with SP565 beam splitter. Tile scans were taken of a large section of the ear (4–12 mm2), and individual fields were merged into a single image. Sequences of image stacks were transformed into volume-rendered, three-dimensional images with Imaris Version 9.1 (Bitplane). The Imaris spot function was used to visualize GFP+ P14s. Tile areas with high levels of autofluorescent hair were trimmed from further analysis with the Imaris crop three-dimensional function, yielding tiles between 4.00–9.25 mm2
Tumor induction and monitoring
The B16 melanoma cell line was provided by Lyse Norian (University of Alabama at Birmingham). B16 cells were grown in DMEM with 4.5 g/l d-glucose, l-glutamine, 10% FCS (HyClone Laboratories), and supplementum complementum (made in-house). Cell lines were passaged every 2–3 d and/or when cell confluency was >80% in 75-cm2-tissue culture flask. Cells were not sequentially passaged longer than 3 wk. In vitro and in vivo tumor growth did not vary considerably throughout the study. For implantation, 2 × 104 B16 cells were injected s.c. at 100 μl volume with equal parts B16 medium and Matrigel Matrix (356234; Corning) into the left hind flank of LCMV-infected memory B6 mice that contained P14 memory CD8 T cells (40, 41). Tumor progression was determined by measuring tumor length multiplied by width using an electronic digital caliper. Mice were sacrificed upon reaching any animal protocol threshold including tumor length of >15 mm or tumor ulceration.
Statistical analysis
Unless stated otherwise, data were analyzed using Prism6 software (GraphPad) using two-tailed Student t test, one-way ANOVA, and two-way ANOVA with a confidence interval of >95% to determine significance (*p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001, and ****p ≤ 0.0001). Data are presented as SEM.
Results
The magnitude and kinetics of the cytokine/chemokine storm depend on sepsis severity
Sepsis is an exaggerated host inflammatory response characterized by increased pro- and anti-inflammatory cytokine and chemokine production. This unbridled inflammatory response represents a crucial event in the pathogenesis of sepsis with important implications on the ability of the host to properly respond to ongoing and/or future infections. To firmly establish a correlation between severity of sepsis and cytokine/chemokine storm in our hands, experimental CLP model of sepsis induction was employed to induce moderate and severe sepsis (one or two cecal punctures, respectively). Sham surgery that does not include CLP was included as control (Fig. 1A). Following surgery, mortality and morbidity (weight loss) were measured (Fig. 1B, 1C), and serum was collected at 6, 12, 24, 48, and 72 h postsurgery. Serum cytokine and chemokine concentrations were determined by Bioplex ELISA (Fig. 1D, 1E). Although no changes in morbidity (weight loss) were observed between the severe and moderate CLP groups, severe CLP mice had a significantly higher mortality—0% mortality in the CLP-moderate group compared with 50% mortality in the CLP-severe group (Fig. 1B, 1C). Of note, moderate CLP procedure in our hands evokes <10% mortality (this is historical average mortality for ≥500 mice used in the last 10 y); >90% of mice that succumb to moderate CLP surgery die in the first 48 h, and mice recover their weight loss in the first 7–10 d after surgery (39, 42).
Importantly, the amounts of cytokines/chemokines detected and duration of the cytokine storm were dependent on the sepsis severity, with the extended production of pro- and anti-inflammatory cytokines such as IL-1β, TNF-α, IL-10, and IL-2 observed in the severe sepsis groups at 24 h postsepsis. (Fig. 1D, 1E). The concentrations of cytokines/chemokines detected by BioPlex assays used in this study should only be considered as direct comparison between groups in well-controlled experiments and not as accurate measure of their concentrations in the plasma. In addition, bacterial burden detected in the livers and spleens 12 h after CLP surgery correlated with the severity of sepsis induced (Supplemental Fig. 1A, 1B). Thus, similar to existing reports (43, 44), a direct correlation between the severity of sepsis, bacteremia, and amount and duration of inflammation was established.
Severe sepsis disrupts the barrier function of vascular endothelium and increases the level of inflammation in situ
Vascular dysfunction is another key hallmark in the host response to sepsis (45). To determine if vascular permeability concordantly increases with severity of sepsis, CLP-severe, CLP-moderate, and sham surgeries were performed on naive B6 mice. Two days later, mice were injected i.v. with Evans blue dye (Fig. 2A). Thirty minutes after injection, tissues were harvested (ear, intestines, and lungs) and processed, and the amount of Evans blue dye was quantified using OD reading (46). Morbidity and mortality were assessed (Fig. 2B, 2C), and an increase in mortality with the mice receiving CLP-severe surgery compared with CLP-moderate and sham surgery recipients was observed. Interestingly, the amount of Evans blue dye (calculation based per gram of tissue) was higher in the tissues of the CLP-severe mice compared with both the CLP-moderate and sham mice (Fig. 2D, 2E, Supplemental Fig. 2A–C), in which the amount of dye did not differ. Importantly, and potentially driven by the changes in the vascular permeability, the level of inflammation (e.g., IL-6, IL-10, and IL-1β) detected in the ear (skin), lungs, and spleens correlated with the sepsis severity (Supplemental Fig. 3A–D). Therefore, these data suggest the barrier function of the vascular endothelium in various tissues throughout the body can be disrupted by modulating the severity of sepsis induced, which correlated with the increased inflammatory response(s) detected in situ. Future studies will define the extent to which increase of inflammation inside tissues is driven by the increase in circulatory cytokines/chemokines and/or in situ production by resident cells.
Severity of sepsis influences the susceptibility of skin CD8 TRM to numerical alterations
Our published data show moderate sepsis (0–10% mortality) does not evoke significant numerical decline of skin CD8 TRM (generated in response to VacV) when compared with pathogen-specific CD8 circulatory memory T cells (TCIRCM). Furthermore, these skin CD8 TRM retained their effector functions (e.g., Ag-dependent IFN-γ production) (28). Because of their localization within the tissue parenchyma, we hypothesized the CD8 TRM were “shielded” from the systemic apoptosis-inducing cytokine storm. In moderate sepsis, the barrier function of the vascular endothelium remains intact, which we posit helps facilitate CD8 TRM survival. However, data in (Fig. 2 show the barrier function of the skin vascular endothelial cells was compromised during severe sepsis. As the endothelial cells lose their barrier integrity, we reasoned that circulating factors (such as cytokines and chemokines) could now leach into the otherwise shielded tissue parenchyma and affect the cellular and/or functional integrity of the CD8 TRM. Thus, to define the extent to which disruption of endothelial barrier under more severe sepsis conditions affects the numbers of skin-embedded CD8 TRM, a physiologically relevant number (5 × 103 cells per mouse) of naive Thy1.1 TCR-transgenic P14 gp33-specific CD8 T cells were adoptively transferred into naive B6 Thy1.2 mice prior to skin VacV-GP33 infection (Fig. 3A) (28). Surgery was performed on VacV-immune mice once the CD8 TCIRCM and TRM compartments were firmly established (day 30 postinfection). A fluorescent intravascular label was administered 3 min prior to tissue collection on day 2 postsurgery to distinguish circulatory (i.v. positive [i.v.+]) and tissue-resident (i.v. negative [i.v.−]) Thy1.1 P14 memory CD8 T cell subsets (Fig. 3A) (37). CD8 TRM are not positively stained using this technique under normal conditions, but the possibility exists that the disruption of the endothelial barrier during severe sepsis could enable the i.v. administered mAb to reach the tissue-embedded Thy1.1 P14 CD8 T cells and mark them as “circulatory” rather than “tissue resident.” To control for this possibility, the frequency of i.v.− (tissue resident), i.v.+ (circulatory), and CD103+ i.v.+ cells in the skin was determined in all groups of mice. Importantly, no changes in the percentage of i.v.− and i.v.+ P14 CD8 T cells in the ear skin was observed after severe sepsis compared with moderate sepsis or sham controls (Fig. 3B, 3C). In addition, the frequency of CD103+ i.v.+ P14 CD8 T cells remained low and indistinguishable between groups, suggesting the i.v. administered mAb does not label bona fide TRM even in the severe sepsis group (Fig. 3D).
As expected, both sepsis modalities led to significant reductions in number of P14 TCIRCM in the blood and spleen (Fig. 3E, 3F). In line with previous data (28), moderate sepsis did not influence the number of CD103+ i.v.− P14 TRM in the skin (Fig. 3G). In contrast, severe sepsis led to a 2.4-fold decrease in the total number of P14 TRM (Fig. 3G). Thus, the data in (Figs. 2 and (3 collectively show CD8 TRM, which are protected in nonlymphoid tissues during a moderate septic event, become potentially susceptible to the detrimental effects of the cytokine storm following a severe septic event as a result of breeches in the vascular endothelium.
In vivo imaging of skin CD8 TRM confirms numerical decline after severe sepsis
To formally prove CD8 TRM in the skin undergo numerical decline during a severe septic event, two-photon microscopy—an approach that does not rely on labeling cells with i.v. injected mAb—was employed to visualize and quantitate bona fide CD8 TRM. To achieve this, naive Thy1.1+ P14-RFP T cells (1 × 104 cells per mouse) were adoptively transferred into naive B6 recipients, followed by VacV-GP33 infection the next day. Sham, moderate CLP, and severe CLP surgery was performed 30 d later, and the total number of RFP+ cells was determined in infected ears 2 d postsurgery using in vivo two-photon microscopy (28, 47). We observed a marked decrease in the number of RFP+ P14 CD8 T cells in CLP-severe mice compared with CLP-moderate mice (Fig. 4A, 4B). Moreover, we saw no difference in the number of RFP+ P14 CD8 T cells when comparing the sham and CLP-moderate groups (Fig. 4C). To increase the resolution of the in vivo imaging, the experiment was repeated using GFP+ P14 CD8 T cells transferred at 5-fold higher numbers (5 × 104 cells per mouse) prior to VacV-GP33 infection (Fig. 4F). Importantly, severe sepsis significantly diminished the number of ear-resident GFP+ P14 CD8 T cells compared with sham and/or moderate sepsis groups (Fig. 4G, Supplemental Fig. 4). Thus, these data confirm the number of tissue-embedded memory CD8 T cells can be differentially affected depending on the severity of the septic insult.
Severe sepsis leads to increase in apoptosis of CD8 TRM
We next aimed to determine potential mechanisms responsible for the numerical decline in CD8 TRM during severe sepsis. To determine if increased apoptotic death was occurring, mice were seeded with P14 CD8 T cells, infected with VacV-GP33 to generate CD8 TRM, and then underwent sham, moderate CLP, or severe CLP surgery 30 d later (Fig. 5A). On day 2 postsurgery, mice were injected with anti-CD45.2 mAb i.v., and tissues were collected and processed to determine the extent of caspase-3/7 activity (Fig. 5A). When examining P14 CD8 TCIRCM cells from the spleen and peripheral blood, we noted a stepwise increase in caspase-3/7+ cells from CLP-moderate and CLP-severe mice compared with sham controls (Fig. 5B, 5C). Consistent with our previous data (28, 47), moderate sepsis did not result in an increase in caspase-3/7+ CD8 TRM in the skin (Fig. 5D), which is supported by the observation of no measurable decline in numbers of these cells (as shown in (Figs. 3G and (4E). There was, however, a significant increase in frequency of caspase-3/7+ CD8 TRM in the severe sepsis group (Fig. 5D). Thus, these data show severe sepsis leads to an increase in apoptotic CD8 TRM.
The severity of sepsis influences the capacity of skin CD8 TRM to protect against homologous skin infection
Following the observation that skin CD8 TRM decrease in number during severe sepsis, we next examined the functional consequences of this phenomenon. We generated P14 CD8 TRM and performed sham or CLP surgery 30 d later. Groups of naive, nonimmunized B6 mice were included at the time of surgery as infection controls. Mice were then infected with VacV-GP33 in the ear 2 d after surgery. Ears were excised 2 d later, and viral load in the skin was measured (Fig. 6A). It is important to note that although sepsis does increase the susceptibility of naive mice to localized VacV skin infection, the initial (up to day 2 postinfection) viral titers were similar in the naive sham or CLP-moderate groups after primary infection (28). All of the groups of nonvaccinated mice had similar viral titer, indicating the existence of memory CD8 T cells could mediate local protection against viral reinfection (Fig. 6B). However, within the vaccinated hosts, CLP surgery reduced the level of protection, as both the CLP-moderate and CLP-severe groups had significantly higher viral titers than sham mice (Fig. 6B). Furthermore, CLP-severe surgery resulted in even greater loss of protection than CLP-moderate surgery (Fig. 6B), confirming the relationship between the severity of the septic event, number of CD8 TRM, and protection they mediate.
The severity of sepsis controls the numerical loss of tumor-infiltrating and nonspecific memory CD8 T cells
The experiments described so far have focused on understanding the impact of sepsis severity on skin CD8 TRM within the context of virus infection. To expand the relevance of our data showing how sepsis, depending on the severity, has a capacity to influence the number and function of CD8 T cells outside of the vasculature, we used the B16 melanoma model to interrogate role of sepsis on tumor-infiltrating, virus-specific memory CD8 T cells (tumor-infiltrating lymphocytes [TILs]). Specifically, naive Thy1.1+ P14 CD8 T cells (1 × 104 cells per mouse) were transferred into naive Thy1.2+ B6 mice prior to LCMV-Armstrong challenge. Once memory had been firmly established (30 d postinfection), all immune mice were s.c. challenged with B16 melanoma in the hind flank (40, 41) and then underwent sham or CLP surgery 15 d later (Fig. 7A). Of note, tumor-infiltrating, virus-specific memory CD8 T cells residing inside tumor mass are considered to be bystander, tumor nonspecific, memory CD8 T cells (48), enabling an additional model in which demise in memory CD8 T cell subsets can be interrogated by modulating severity of septic insult.
Tumors were collected after i.v. injection of anti-CD45.2 mAb and processed to enumerate the virus-specific memory P14 CD8 TILs by flow cytometry (Fig. 7A). To determine how the severity of the septic event influences the ratio of memory P14 CD8 T cells in the circulation and within the tumor, the ratio of i.v.+/i.v.− P14 CD8 T cells was analyzed across the groups. We observed a trending decrease in the ratio of i.v.+/i.v.− P14 CD8 T cells in mice receiving CLP surgery compared with mice who received sham surgery, highlighting the correlative loss of TCIRCM with the increased severity of the septic event (Fig. 7B). Next, we determined the number of i.v.+ and i.v.− P14 CD8 T cells in the tumor. In both cases, mice that experienced CLP-severe surgery had significantly fewer i.v.+ and i.v.− P14 CD8 T cells compared with sham surgery recipients. By comparison, moderate CLP surgery did not lead to a significant decline in i.v.− P14 CD8 TIL numbers (Fig. 7C, 7D). Collectively, these data show the number of tumor-infiltrating, virus-specific bystander memory CD8 T cells present after sepsis is dependent on the severity of the septic insult, which was similar to what was seen for pathogen-induced CD8 TRM present at the site of the infection.
Discussion
Sepsis is a serious medical condition that accounts for ∼20% of deaths around the world. Sepsis is characterized by an overreactive immune response to a systemic infection and leads to uncontrolled inflammation, vascular dysfunction, acute lymphopenia, and long-term immunoparalysis. It is important to note that a spectrum of disease severity exists for septic patients: some patients are able to overcome the septic event, whereas others succumb and die. Our laboratory and others have employed the CLP model of sepsis because of its similarities in the clinical manifestation and presentation of the disease (49–52). In this report, we used CLP to model the spectrum of disease severity seen in patients. Specifically, we modulated the severity of the septic event by altering the “puncture” portion of the CLP protocol—a single puncture of the cecum was used to induce a moderate septic event, whereas two punctures induced a more severe sepsis. The increased duration and magnitude of the cytokine storm observed in CLP-severe mice models the disease pathogenesis seen in human populations, with more septic patients having increased amounts of cytokine present in the serum (53). Furthermore, CLP severe resulted in increased vascular dysfunction and more pronounced lymphopenia when compared with CLP moderate, again better modeling the spectrum of disease severity.
Of the patients that survive a septic event, they display increased susceptibility to secondary infections and viral reactivation due to long-term immunoparalysis (16). Studies have detailed the impact of sepsis on naive and Ag-experienced CD4 and CD8 T cells in the circulation (25, 39, 54), but the influence of sepsis specifically on CD8 TRM has been neglected until recently (28). Previous data from our laboratory have demonstrated CD8 TRM are numerically stable and maintain their function in the context of a moderate septic event (0–10% mortality). The maintenance of CD8 TRM number and retention of their function during moderate sepsis is most likely due to their localization in tissue parenchyma, where they are likely shielded from the apoptosis-inducing cytokines circulating in the vasculature. However, vascular junction integrity is lost during a more severe septic event (i.e., one resulting in ∼50% mortality), and cytokines (and likely other factors) are now able to leach into the tissue. In this setting, we were now able to detect increased CD8 TRM apoptosis and subsequent numerical decline (which is intent of our experimental design), but we expect other pathophysiological effects were being experienced in the tissue parenchyma in the CLP-severe mice. Our use of Evan blue dye to quantify vascular dysfunction supports this hypothesis, but this technique does not give us any indication which anatomical locations (e.g., blood vessel versus capillary bed) are experiencing endothelial barrier disruption. Supernatants from tissues sections may be examined for cytokine levels to confirm elevated levels present in the parenchyma.
A number of laboratories, including ours, have begun to include two-photon microscopy to visualize CD8 TRM in situ (28, 55). We used this methodology to confirm the loss of CD8 TRM in nonlymphoid tissue after a septic event, and our data show significant reduction in the overall TRM cellularity of a tissue. Interestingly, some areas appeared to become completely devoid of CD8 TRM during sepsis, whereas other areas were spared of this loss. This difference in CD8 TRM susceptibility may be due to their location with respect to blood vessels and capillary beds, allowing parts of the tissues to be bathed in apoptosis-inducing cytokines and other factors, whereas the localized concentrations in other parts of the tissues did not reach critical levels to trigger apoptosis in those CD8 TRM.
The molecular driver(s) of T cell apoptosis during sepsis has been difficult to ascertain, whether this is for circulating T cells or those resident in nonlymphoid tissues (such as the TRM specifically investigated in this study). Data generated from a number of papers have suggested sepsis-induced T cell apoptosis is regulated by a complex network of pro- and antiapoptotic proteins within the T cells, which can be further influenced by a number of extracellular factors. Similarly, administration of mAb specific for TIGIT, 2B4, OX40, and PD-1 (among others) can modulate the number of T cells in the circulation or primary lymphoid organs by changing the frequency of apoptotic cells (56–60), but it remains to be determined if these same proteins contribute to the apoptosis of TRM during severe sepsis. Interestingly, recent data by Nedeva et al. (61) identified the importance of TREML4 receptor in regulating the inflammatory cytokine response and innate immune cell (i.e., neutrophils) after cecal slurry injection. It would be interesting to use the same CRISPR-screening approach to confirm the importance of the various proteins that have been suggested to control T cell (both circulating and tissue resident) apoptosis during sepsis, as well as identify other new regulatory proteins
To examine the functional consequences of local CD8 TRM decline after a septic event, mice were given a homologous VacV challenge. The extent of viral clearance was measured 2 d postinfection via plaque assay. We observed higher viral load in the CLP-severe mice compared with CLP-moderate mice, suggesting the local loss of CD8 TRM resulted in reduced protection from homologous infection. This observation also has important translational implications, in which patients who survived a septic event may have lost the memory immune cells responsible for protection against common pathogens at barrier tissues—a primary function of CD8 TRM. Although others have documented the effects of sepsis on B cells and Ab production (62, 63), our focus was primarily on the CD8 T cell arm of adaptive immunity. We cannot rule out the contribution of B cells and/or vaccinia-specific Abs in this homologous reinfection model, and further interrogation may elucidate additional mechanisms that contribute to the loss of protection. Furthermore, these data raise the question of whether septic survivors would benefit from revaccination. Our data make it tempting to speculate secondary (booster) immunizations would help to restore CD8 TRM in barrier tissues of survivors of a severe septic event, as their prior immunity may have waned as a result of sepsis-induced CD8 TRM attrition. To extend our findings to other relevant models, we examined how the severity of sepsis influenced the maintenance of TILs. CD8 TILs control tumor growth, both in a nonspecific (bystander) and Ag-specific manner (41, 64, 65). Because these CD8 TILs are thought to be embedded in the tumors and outside of the circulation, we interrogated how a severe septic event may influence the numbers of this important population of cells. We observed a significant decrease in the number of tissue-infiltrating (i.v.−) P14 CD8 TILs present in the tumor. Because these CD8 T cells are specific for gp33, an Ag not expressed by the B16 melanoma tumor cells, these cells model bystander CD8 TILs. It would be interesting to observe the effect of a severe septic event on tumor-specific CD8 TILs, and the consequence of the loss of these CD8 TILs on tumor growth, in future studies.
In summary, our data collectively show that increasing the severity of the septic event increases the length and magnitude of cytokine production, impairs the barrier function of vascular endothelium, and results in the correlational numerical decline of both the CD8 TCIRCM and TRM compartments. Following a severe septic event, the amount of a number of cytokines (such as IL-10 and TNF-α) more than doubled at their peaks when compared with a moderate septic event. Moreover, other cytokines (such as IL-1β and IL-2) were present at 24 h postsepsis in CLP-severe mice, whereas they were almost undetectable in CLP-moderate mice. This finding indicates a prolonged and more intense cytokine storm is present in CLP-severe mice, which in turn may lead to a more profound numerical decline in a number of CD8 T cell memory populations. Furthermore, we showed increased caspase activity indicates increased levels of CD8 TRM apoptosis, likely driving the numerical decline of these cells. As a result of the loss of the CD8 TRM, CLP-severe mice were less protected from viral rechallenge.
Finally, we extended our findings to cancer immunology to show severe sepsis can also lead a numerical decline of tumor-infiltrating bystander (i.e., virus-specific) memory CD8 T cells. These data, together with previously published findings, further elucidate how CD8 TRM can be affected during a septic event and the functional consequences of their numerical decline. These data also give us insight into how blood vessel physiology during sepsis can shape immunological landscapes within tissues. Highly vascularized tissues may experience increased loss of CD8 TRM compared with tissues with lower vascularization because of the number of apoptotic factors that flow into these tissues during sepsis.
Overall, this report adds important information to the current body of literature surrounding the impact of sepsis on memory CD8 T cells subpopulations and specifically dissects how the severity of sepsis concordantly influences the numerical decline of CD8 TRM. The translational implication of this study relates directly to the spectrum of disease severity observed in septic humans. It is integral to determine how variation in sepsis severity dictates the extent of impairment in memory CD8 T cell subpopulations, as lesions in this population render the host susceptible to secondary, repeated, and/or nosocomial infections. Furthermore, our study sheds light on some of the potential mechanisms behind the increase in susceptibility to reinfections among sepsis survivors and informs future studies designed to boost the recovery of protective circulatory and TRM after sepsis. Future human studies that analyze the recovery of these memory T cell populations will benefit from the newfound understanding into how the severity of the initial septic insult influences the numbers and function of pre-existing infection and/or vaccine-induced memory CD8 T cells.
Acknowledgements
We thank members of our laboratories for technical assistance and helpful discussions.
Footnotes
This work was supported by National Institutes of Health (NIH) Grants GM134880 (to V.P.B.), AI114543 and AI151183 (to V.P.B. and J.T.H.), GM115462 (to T.S.G.), R35GM140881 (to T.S.G.), T32AI007485 (to D.B.D. and I.J.J.), and AI42767, AI085515, and AI100527, (to J.T.H.). Support was also received from Foundation for the NIH T325T32HL007 (to S.M.A.) and the Holden Comprehensive Cancer Center at The University of Iowa and its National Cancer Institute Award P30CA086862 (to V.P.B.). This work was also supported by Veterans Administration Merit Review Award I01BX001324 (to T.S.G.).
The online version of this article contains supplemental material.
References
Disclosures
The authors have no financial conflicts of interest.