CD4+ T cell production of IL-10 and regulation of immune responses in aging
A dissertation submitted to the Graduate School of the
University of Cincinnati
in partial fulfillment of the requirements for the degree of
Doctor of Philosophy
Immunology Graduate Program, College of Medicine
2018 by
Maha Almanan M.D., Faculty of Medicine, University of Khartoum, Khartoum, Sudan
Committee Chair: David Hildeman, Ph.D. Abstract
The immune system plays a vital role in the protection against invading pathogens. A successful immune response is dictated by the fine balance between pro- and anti-
inflammatory mediators. With age the immune system undergoes a progressive
dysregulation due to the dramatic reduction in the naïve T cell pool, a relative clonal
expansion of memory T cells as well as accumulation of regulatory T cells. Another
aspect of this immune dysregulation is the persistent, low grade inflammation that
develops with age and is referred to as (inflammaging). Together, dysfunctional immune
responses and inflammaging are a great risk for serious age-related pathologies. Thus,
understanding the mechanisms regulating this age-related immune dysfunction is
critical for healthy aging.
Viral infections including latent cytomegalovirus (CMV) are thought to be one of driving forces of age-related alterations in immune function and inflammaging. While T cell responses are critical against CMV infection, the regulatory mechanisms that control latency are not well understood. Regulatory T cells are known to accumulate with age,
likely due to their enhanced survival. While Treg accumulated with age, their per cell
functionality remains controversial and possibly model dependent. Treg cells are important regulators of acute infections (including CMV). However, the impact of Treg on latent CMV is unclear. In chapter 3 we aimed at investigating the role of Foxp3+ regulatory T cells (Treg) in latent murine cytomegalovirus (MCMV) infection. We show
that Treg played divergent roles in the control of MCMV infection. In the spleen, Treg
antagonize CD8+ T cell effector function and promote viral persistence, while in the
II salivary gland Treg prevent IL-10 production from Foxp3- CD4+ T cells and limit viral
reactivation and replication. Together, work in chapter 3 broadens our understanding of
the homeostasis and functions of regulatory T cells and IL-10 with age.
Our discovery that Treg control IL-10 during latent CMV infection was intriguing. IL-10 is
a known regulatory cytokine, playing a major role in shaping immune responses and
regulating excessive inflammation. Thus, IL-10 may contribute to counter-regulation of
age-driven inflammation. In chapter 4 of this dissertation, we aimed to investigate the
role of IL-10 with age and further understand the cellular and molecular mechanisms
underlying its production. We provide evidence that aging favors the accrual of CD4+
Foxp3- IL-10+ T cells. Additionally, T follicular helper cells (Tfh) were major producers of
IL-10 in aged humans and mice which we have designated as Tfh10 cells. Interestingly, both IL-21 and IL-6 were indispensable for the accumulation of Tfh10 cells with age.
Finally, the loss of BCL6 in Foxp3- CD4+ T cells enhanced the production of IL-10 with
age. Results from this study highlight the dynamic regulatory/counter-regulatory balance
controlling age-related inflammation. More importantly, we show that despite age-driven
intrinsic defects in adaptive immune responses, blockade of IL-10 signaling is largely
sufficient to restore germinal center (GC) B cell responses in aged mice. Together work
from chapter 3 and 4 underscores the complexity of the unique age-related regulatory
networks that shape immune responses to latent infections, as well as vaccines.
III IV Acknowledgments
I would like to express my sincere gratitude to my mentor, Dr. David Hildeman. Thank you for your endless support and patience over the last five years, sense of humor, immense scientific knowledge and for challenging me to think independently. Thank you for believing in me at times when I didn't believe in myself, I cannot think of a better mentor to have. I don’t know how you do it, but I always looked up to you and I aspire to become a great role model and pass on what you've taught me to my own students one day. I just want you to know that I am a much better scientist because of you and for that I will always be grateful for the rest of my life.
I would like to acknowledge members of my committee who provided endless support and guidance throughout my PhD study. Dr. Claire Chougnet, I am grateful for all the invaluable scientific discussions which has greatly impacted my PhD training and I am also thankful for the time you have taken to offer career and life advice. Dr. Kasper
Hoebe, you have always challenged my scientific curiosity and for that I thank you. Dr.
Kris Steinbrecher, thank you for investing your time and personally assisting with my experiments. Dr. Rhonda Cardin, thank you for your patience while introducing me to the world of virology and assisting with experiments, I am truly grateful for all your support.
I would like to thank my lab members Allyson Sholl, Jana Raynor, Kun-Po Li, Matthew
Alder and Pulak Tripathi, who have grown to become my second family. You all were very helpful at various times at the beginning of me joining the lab and I have continued to succeed because of it. Cyd Castro, Sarah Jones, Rachel Walters, and Sharmilia
V Shanmuganad, thank you all for your contributions to my work at different times.
Especially within the last 2 years, we have grown to become one sisterhood in science and I am forever grateful.
I would also like to thank the Immunology Program who accepted me and gave me one of the best opportunities of my life and allowed me to meet some of the most wonderful people and some of my best friends. I especially thank Hesham Shihata, Kate Carroll,
Carolyn Rydyznski, and Courtney Jackson, for always caring, listening and being a shoulder to lean on throughout the years.
Lastly, I would like to thank my family for their continued love and support, especially my mother who was my biggest motivation to pursue my graduate studies. Even though you are no longer in the present, I hope that I have made you proud. Dad, you have taught me to continue to keep my head up and gave me your blessings and because of the way you have raised me, I feel I have become a better parent. To my children,
Yasmeen and Mosaab, at times it was tough to juggle being a student and being your mother. At times people have asked how do I manage, I just say to myself because of you. I am truly blessed to have you in my life.
VI Table Of Contents
Abstract ...... II Acknowledgments ...... V
Table of Contents ...... VII
List of Abbreviations ...... X Chapter 1: Introduction ...... 1 1. Age-dependent immune dysregulation ...... 1
1.1. Defects in innate immune-responses with age ...... 2
1.2. Defects in adaptive immune-responses with age ...... 6
1.2.1. B cells ...... 6
1.2.2. T cells ...... 8
(i) Homeostatic defects in T cells with age ...... 8
(ii) Functional defects in T cells with age ...... 9
1.3. Aging and its impact on vaccine responsiveness ...... 10
2. Regulatory T cells ...... 11
2.1. Thymus derived Treg (tTreg) ...... 12
2.2. Peripherally derived Treg (pTreg) ...... 12
2.3. Treg functionality and specialization ...... 14
2.4. Treg in aging ...... 15
3. Cytomegalovirus (CMV) 16
3.1. Acute CMV ...... 17
3.2. Latent CMV ...... 20
3.2.1. Cellular and molecular mechanisms controlling CMV reactivation
from latency ...... 21
3.2.2. CMV in aging ...... 22
3.2.3. Regulatory mechanisms controlling CMV ...... 23
VII (i) Treg in CMV ...... 23
(ii)IL-10 in CMV ...... 24
4. Interleukin-10 (IL-10) ...... 25
4.1. Transcriptional regulation of IL-10 production in CD4 T cells ...... 26 4.1.1. TCR mediated IL-10 expression ...... 26
4.1.2. Cytokine mediated IL-10 expression ...... 27
4.2. IL-10/IL-10R and signaling pathway ...... 29
4.3. Biological effects of IL-10 ...... 31
4.4. Aging and IL-10 ...... 33
5. T follicular helper (Tfh) cells ...... 34
5.1. Aging and Tfh cells ...... 35
6. Interleukin-6 (IL-6) ...... 37
6.1. IL-6 signaling pathway ...... 37
6.1.1. Classical pathway ...... 37
6.1.2. Trans-signaling pathway ...... 38
6.2. Biological effects of IL-6 ...... 39
6.3. Aging and IL-6 ...... 40
7. Interleukin-21 (IL-21) ...... 41
7.1. IL-21 signaling pathway ...... 41
7.2. Biological effects of IL-21 ...... 42
7.3. Aging and IL-21 ...... 43
Summary ...... 44
References ...... 46
Chapter 2: T-reg Homeostasis and Functions in Ageing ...... 81
VIII References ...... 97
Chapter 3: Tissue-specific control of latent CMV reactivation by regulatory T cells ...... 104
References ...... 126
Figures ...... 128
Chapter 4: IL-10 producing Tfh cells counter inflammaging but suppress humoral responses ...... 151
References ...... 172
Figures ...... 180
Chapter 5: Summary and future directions 200
Figures ...... 220
References ...... 224
IX List of Abbreviations
AID activation-induced cytidine deaminase
APCs antigen presenting cells
Bim Bcl-2 interacting mediator of cell death
CMV cytomegalovirus
CSR class switch recombination cTreg central Treg
DC Dendritic cell eTreg effector Treg
FoxP3 Fork-head box P3
GC germinal center
G-CSF granulocyte colony-stimulating factor
GM-CSF granulocyte-monocyte colony-stimulating factor
ICOS Inducible co-stimulator
IL-10 Interleukin 10
IL-21 Interleukin 21
IL-6 Interleukin 6
LPS lipopolysaccharide
NK natural killer cells
PAMPs pathogen-associated molecular patterns
PRRs pattern recognition receptors pTreg peripherally derived Treg
RTEs recent thymic emigrants
X STAT3 signal transduced and activator of transcription 3
TCR T cell receptor
Tfh T follicular helper cell
TLRs Toll-like receptors
Treg Regulatory T cell tTreg thymus derived regulatory T cell
XI Chapter 1
Introduction
1. Age-related immune dysregulation
The global population aged 60 years and over numbered 962 million in 2017, more than
twice as large as it was in 1980, with predictions of higher rates by 2050. More
importantly, the number of persons aged 80 years and over is projected to increase
more than threefold between 2017 and 2050, rising from 137 million to 425 million (1).
This demographic shift is expected to have a huge economic impact, particularly on health care.
One of the profound consequences of aging is the functional dysregulation in innate and adaptive immune responses (2). Importantly, a significant change favored by this immune dysregulation is the enhanced production of pro-inflammatory mediators resulting in inflammaging (3-6). Together, the immune dysregulation and inflammaging are suggested to be the underlying causes of most of the diseases of the elderly such as cancer, Alzheimer's disease, Parkinson's disease, multiple sclerosis and atherosclerosis (7-10). A major focus of aging research is to understand the cellular and molecular mechanisms underlying immune defects with age with an ultimate goal of improving the quality of life for older individuals and decreasing the costs of health care.
In this introductory section we will summarize the current understanding of the age- driven defects in the immune system and their relevance toward susceptibility to disease.
1 1.1. Defects in innate immune-responses with age
The effect of aging on the innate immune system in humans and mice is an area of
rapidly advancing research. Interestingly, accumulating evidence indicates that aging
has a profound impact on innate immune cell function and homeostasis, which in turn
greatly impacts the adaptive immune response. Innate immunity is mediated by various
cell types such as natural killer (NK) cells, neutrophils, macrophages, dendritic cells
(DCs) and others. In this section, we will summarize the recent knowledge on the
effects of aging on innate immunity.
NK cells are critical for eliminating cancerous and virally infected cells. Several defects
in NK cell maturation and function have been reported with age. For example, previous
studies provided evidence for the defective response of aged NK cell to cytokines like
IL-2, IL-12, IFN-α, IFN-β, and IFN-γ (11). More importantly, cytotoxic activity of human
and murine NK cells declined with age (12-14). Thus, both defects involoving NK- mediated cytotoxicity and/or cytokine production result in lack of functionality with age.
Other studies investigated the homeostasis of NK cells based on their maturation status. Interestingly, there is strong evidence for reduced proportion of mature NK cells with age, which appears to be due to the aged non-hematopoietic environment (15).
Together, these data suggest that aging has profound effects on NK cell functionality and maturation which might compromise immune responses to tumors and viral infections in elderly.
Neutrophils are another type of innate immune cell that plays a critical role in host defense. Several studies have investigated whether there are changes in neutrophil
2
function or homeostasis with age. The data remain controversial. For example, prior
work has shown no difference in neutrophil tissue adherence, migration, and
phagocytosis (16), suggesting marginal defects in neutrophil function with age.
However, another study comparing neutrophil function in young (23–35 years) and elderly (>65 years) individuals reported a reduction in the phagocytic index of neutrophils with age as well as a defect in neutrophil maturation (17).
In addition to their function, neutrophil survival and persistence at sites of inflammation
are also important controllers of infection and immunopathology. Given the increased
inflammation with age, several groups have investigated the survival of aged
neutrophils. Interestingly, aged neutrophils failed to survive under the effect of pro-
inflammatory mediators, such as granulocyte-monocyte colony-stimulating factor (GM-
CSF), granulocyte colony-stimulating factor (G-CSF ), and bacterial lipopolysaccharide
(LPS) (18). Combined, these data suggest that neutrophil function per se is minimally disturbed in aging; however, survival of aged neutrophils is shortened significantly, likely due to their altered response to inflammatory environment. Future studies will be
necessary to better understand the seemingly context-dependent functional changes in
neutrophils with age.
Macrophages are another integral component of the innate immune system. Several studies investigated the phenotype of macrophages and their activity in response to stimuli as readout for their functionality. While some studies reported functional impairment of macrophages with age, this phenotype seems to be reversible. For
example, aged macrophages were defective in their response to IFN-γ stimuli leading to
3
reduced nitric oxide production and increased susceptibility to parasitic infections as well as tumors (19-21). In contrast, other studies have argued that this age-related dysfunction in macrophages is reversible in response to in vitro treatment with multiple cytokines, suggesting that functional defects in macrophages with age are due to the environmental exposure (22). Interestingly, the aging environment favors a shift in adipose tissue macrophages towards an inflammatory phenotype with enhanced production of IL-6 and TNF-α (23). These latter data implicate adipose tissue
macrophages as key contributors to the enhanced inflammation observed with age.
Altogether, aged macrophages seem to acquire substantial plasticity that can shape
their functionality and phenotype in response to the aging environment.
Dendritic cells (DCs) are key participants in the generation of immunity and
maintenance of tolerance. Aging has a profound negative effect on DCs. For example, aged human DCs are less able to prime CD4+ and CD8+ T cell responses relative to young DCs (24, 25). Additionally, aged human DCs showed a reduction in phagocytosis as well as a decline in migration ability (26). Notably, activation of T cells requires the cooperation of DCs expressing MHC class I/II and costimulatory molecules including
CD80, CD86. Interestingly, several studies reported reduced expression of CD80,
CD86, and MHC II on murine DCs with age (27-29), suggesting defective functionality with age. In agreement, recent work has shown that a decline in the phagocytic activity of aged DCs is due to their mitochondrial metabolic dysfunction (30), suggesting that targeting metabolic pathways in DCs is a key for improving their functionality with age.
Together, age-associated functional deficits in DCs might play a substantial role in
increased susceptibility to disease with age.
4 The innate immune response relies on a large family of pattern recognition receptors
(PRRs) expressed on innate immune cells that target pathogen-associated molecular patterns (PAMPs) (31). Three major classes of innate immune receptors have been identified to date. 1) Toll-like receptors, which activate downstream signaling pathways such as (NF-κB) (32). 2) NOD-like receptors mediated by the inflammasome complex activation (33). 3) RIG-I which is critical for the regulation of an antiviral response (34).
Stimulation of innate cells (macrophages, polymorphonuclear cells, natural killer cells and dendritic cells) via these receptors is a critical step for subsequent activation of adaptive immune responses. Indeed, ligation of these receptors by their ligands results in the secretion of cytokines and chemokines that promotes the recruitment and activation/differentiation of adaptive immune cells. Interaction of T cells with antigen presenting cells (APCs) is a key factor for initiating the immune response that normally culminates in the eradication of pathogen and the generation of memory responses for future protection.
There are age-related defects reported in Toll-like receptor signaling and downstream cytokine production which have been mainly attributed to their altered expression. For example, in mice, reduction in the expression of almost all TLRs on macrophages was observed with age. Further, in vitro stimulated splenic macrophages from aged mice secreted lower levels of IL-6 and TNF-α compared to their young counterparts (35).
Similarly, in humans, age-associated decreases in surface expression of TLR1 was associated with deficits in TLR1-induced cytokine production from aged monocytes (36).
Additionally, a study by Panda et al. showed a substantial decrease in old, compared to young individuals in TNF-α production in myeloid (mDCs) and plasmacytoid (pDCs) in
5 response to TLR1/2, TLR2/6, TLR3, TLR5 stimulation. Findings from the study also indicated that the defects in TLR-induced cytokine production with age correlated with poor antibody responses to influenza immunization in elderly individuals (37), suggesting TLR-targeted therapies as a strategy to enhance vaccination efficacy.
Interestingly, poly I:C (TLR3 ligand) given as an adjuvant was sufficient to increase pro-inflammatory cytokines and improve the responsiveness of CD4+ T cells in aged mice (38), suggesting that age-related defects in CD4+ T cell responses are reversible.
Collectively, data emerging from human and mouse studies provide strong evidence for age-related immune dysregulation intrinsic to innate immune cells including aberrant expression and signaling events downstream of TLRs.
1.2. Defects in adaptive immune-responses with age
1.2.1. B cells
B cells and antibody-mediated immunity play an important role against pathogens.
Several studies have suggested defects in B cell homeostasis and function with age
(39). Age-related defects in B cells homeostasis begin with B cell development and persist throughout B cell lifespan. In the periphery, human studies reported that the percentage and absolute number of total B cells, defined by expression of the lineage marker CD19, decrease with age (40). Additionally, mouse studies have reported limited number of naive B cells in the periphery with reduced repertoire diversity due to the clonal expansion of memory B cells with age (41, 42), dominated by antigen- experienced CD5+ B cells (43).
6 A sizable amount of studies suggest that multiple factors during B cell development
contribute to the lack of naïve B cells in aged mice and humans. For example, studies
utilizing mixed bone marrow chimeras suggested that the reduction in B cells with age is
due to reduced hematopoietic stem cell (HSC) commitment to the B lineage, as well as
changes to the bone marrow (BM) microenvironment(44). Additionally, defective B cell
differentiation with age resulted also from reduction in B lineage transcription factors
(E2A and EBF1) as well as reduction in pre-BCR surrogate light chain (45). In contrast, another study showed that the proportions of early B cell precursors are maintained with age in humans (46). Thus, although it is likely that defects during B cell development contribute to altered B cell homeostasis and function with age, the data remain controversial and require verification in subsequent studies.
In addition to their homeostasis, B cell production of antibodies is essential for effective
humoral responses. Functionally, the quality of antibody production from B cells decline
with age resulting in lower-affinity antibodies and reduction in vaccination responses
(47). Indeed, a study by Agematsu et al. provided evidence for a reduction in plasma
cell differentiation and function of IgM memory B cells in response to pneumococcal
vaccine in elderly individuals compared to adults (48). Additionally, in vitro stimulated B
cells from elderly with anti-CD40/IL-4 were profoundly impaired in their capacity to
undergo class switch recombination (CSR) compared to their counterparts (49).
Altogether, B cells encounter major changes in their phenotype and function which contribute to increased risk of infections and loss of vaccination responses with age.
7
1.2.2. T cells
(i) Homeostatic defects in T cells with age
As T cells are key players of the adaptive immune system, extensive research in aging
has focused on their development in the thymus and their homeostasis and function in
the periphery. It is well established that, with age, the size of the thymus declines,
reaching a peak at puberty, followed by a progressive reduction in the output of naïve T
cells (50, 51). Thus, production of naïve T cells is significantly reduced with age (52).
Age-dependent decline in the production of naïve T cells is thought to impair the
response of elderly individuals to new antigens, likely putting the elderly at risk for
infections even with vaccination (53).
With age, both cortical and medullary regions decrease in volume with disorganization
of epithelial cell architecture and of the corticomedullary junction, accompanied by
replacement of lost stromal volume with adipose tissue (54). Furthermore, work from
Hale et al. using GFP to tag recent thymic emigrants (RTEs) in aged mice reported age-
dependent decline in RTE numbers (55). Interestingly, a previous study suggested that reduced thymic output of T cells might be compensated for by a reduction in the pro- apoptotic molecule Bim which enhanced naïve T cell life span in aged mice (56). These
data suggest that the progressive decline in naïve T cell production from the thymus
with age is balanced by enhanced survival in the periphery.
Another concept that has emerged to explain defective T cell responses with age is the
impingement of the naïve T cell repertoire by clonally expanded memory T cells.
8 Although CD4 T cells clonal expansion was observed with age, CD8 T cell clonal expansion is the most prominent (57). One explanation that was suggested for this oligoclonal expansion with age is the repeated exposure to specific viral antigens.
Indeed, expansion of terminally differentiated CD8+ effector cell (TEMRA) populations
has been largely linked to cytomegalovirus (CMV) and Epstein-Barr virus (EBV)
infection (58, 59). Another explanation that was proposed is the bystander proliferation
of CD8 T cell clones in response to Type I IFN production (60). Together, the reduction
in naive T cells in the peripheral pool and predominance of specific memory T cell
clones is thought to contribute to poor responsiveness to new antigens with age and
increased the risk of infections.
(ii) Functional defects in T cells with age
In addition to defects in development and homeostasis, aging also affects T cell
functionality. Indeed, aged T cells experience signaling defects, reduced proliferation
capacity, loss of synapse formation and altered trafficking (61). For example, early work
from Miller et al. documented several defects in TCR signaling including decline in the
activation of the Raf-1/MEK/ERK kinases and in JNK protein kinase with age (62).
Additionally, alteration of lipid raft composition with aging has been shown to contribute
to defective T cell responses (63). Importantly, decreased calcium flux after T cell
receptor engagement or mitogen stimulation along with decreased translocation of the
transcription factor NFATc to the nucleus with age (64), likely contributes to a decline in
T cell functionality with age. Together, aging adversely affect T and B cell homeostasis
9 and function resulting in the loss of ability to fight infections and decline in responses to
immunization.
1.3. Aging and its impact on vaccine responsiveness
Notably, both B and T cell changes with age are associated with a decline in the
antibody response to influenza vaccination (65, 66). Indeed, in young individuals the influenza vaccine provides 65-80% protection while the protection afforded in aged individuals is only 30-50% (67). As a result, 90% of the 10,000–40,000 deaths related to
influenza annually in the United States occur in persons aged ≥65 years. Successful
immunization against influenza is mediated mainly via B cells and T follicular helper
(Tfh) cells that act in concert to produce germinal centers (GCs) where activated B cells
undergo class switch recombination (CSR) and somatic hypermutation (SHM),
producing the high-affinity class-switched antibodies.
Several factors have been attributed to the loss of immunization responses in old
people. Some reports implicated defects in T cells, other reports implicated age-related
B cell defects. For example, work from Lefebvre et al. indicated that aged mice
experienced loss of vaccination responses and protection from influenza infection
compared to young mice. The authors attributed the loss of protection to a shift in aged
CD4 T cells toward Tfh cells rather than Th1 cells in the lungs which are required for
protection from lung infection (68).
B cells also accumulate intrinsic defects that correlate with the reduction in generation
of antibody responses to vaccines with age (65, 69, 70). For example, reduced
10 expression of activation-induced cytidine deaminase (AID) with age results in
diminished class switch recombination (CSR) and loss of production of higher affinity
antibodies in response to vaccination (49). Altogether, the effects of aging on CD4 T
cells and B cells significantly impact vaccination responsiveness. While age-related
intrinsic defects in T and B cell responses is well established in the literature, recent
data has suggested the existence of a suppressive environment that also contributes to
depressed adaptive immune responses with age.
2. Regulatory T cells
Regulatory T cells (Treg), classically defined by expression of high levels of interleukin 2
receptor-α (IL-2Rα/CD25) and transcription factor forkhead winged helix transcription
factor 3 (Foxp3), are indispensable for maintaining immune homeostasis (71). In
humans, mutations in the Foxp3 gene result in profound autoimmune diseases. Indeed,
the immune dysregulation, polyendocrinopathy, enteropathy, X-linked syndrome (IPEX)
is a direct consequence for an X-linked mutation in Foxp3 gene and Treg dysfunction in
humans. The disease was described for the first time in a large family with 19 affected
males across five generations (72). A similar discovery for the disease was made
simultaneously in mice deficient in Treg cells when it was found that these mice also
had defects in the Foxp3 gene (73, 74). Later, it was shown that depletion of Treg cells in mice expressing diphtheria toxin receptor under the control of the foxp3 gene via administration of diphtheria toxin resulted in excessive myelo-proliferative disease and profound inflammation, providing strong evidence that Treg are indispensable for
maintaining peripheral immune homeostasis (75).
11
2.1. Thymus derived Treg (tTreg)
Following their discovery, extensive research was directed towards understanding the development and origin of Treg. Treg cells develop primarily in the thymus (thymus derived Treg, tTreg), or they can also be differentiated in the periphery (peripherally derived Treg, pTreg). During Treg differentiation, thymocytes bearing TCRs with high affinity for self-antigens presented in the thymus are selected into the Treg lineage at the immature CD4SP stage of thymic development (76). As a result of this selection, induction of Foxp3 is initiated leading to the differentiation of tTreg (77). Additionally, both CD80/CD86 and CD28 are required for the differentiation of tTreg cells (78, 79).
Further, cytokines like IL-2 play an essential role in tTreg development and Foxp3 expression (80, 81). Indeed, mice deficient in IL-2 or CD25 can only maintain half of the
Treg pool compared to WT counterpart (80). The incomplete loss of Treg in the absence of IL-2 suggested some redundancy in factors required for their development. Indeed, additional loss of IL-15 and IL-7 resulted in the complete absence of tTreg (82). Thus, evidence from previous research provided great insight into the complex signals that contribute to tTreg differentiation including cytokines and surface expression of co- stimulatory molecules.
2.2. Peripherally derived Treg (pTreg)
In addition to their development in the thymus, Foxp3 expression can be induced in naïve T cells post-thymic development driving their conversion into pTreg. This can occur through two major pathways, one involves TCR stimulation along with TGF-ß and the other involves TCR stimulation and retinoic acid. Differentiation of Treg from
12
CD4+CD25− naive T cell required co-stimulation with T cell receptors (TCRs) and
transforming growth factor β (TGF-ß). These pTreg are functionally similar to nTreg in
suppressing T cell proliferation and Th1 and Th2 cytokine production (83). Further
investigation for the molecular mechanisms involved in TGF-β mediated induction of
Foxp3 in naive CD4+CD25− indicated that IL-2 but not CD28 regulated the induction of
Foxp3 under TGF-β stimulatory conditions. The second pathway of pTreg conversion
occurs primarily in the gut and specifically Peyer’s patches. Here, the induction of pTreg
is mediated by CD103+ dendritic cells which act in the presence of TGF-β and retinoic acid to enhance Foxp3 induction (84). Thus, the peripheral Treg compartment is comprised of nTreg and pTreg both of which likely contribute to immune homeostasis.
As both tTreg and pTreg express CD4, CD25 and Foxp3, a significant amount of
research has investigated potential markers distinguishing tTreg from pTreg. A few
studies have identified the transcription factor Helios and the cell surface marker
neuropilin-1 as being markers that are specifically expressed on tTreg cells (85, 86).
Further complicating the matter in human CD4+ T cells is the fact that normal T cell activation can drive transient increases in Foxp3 expression. Therefore, other markers have been suggested to faithfully identify human Treg cells. For example, a lack of CD127 expression can be used to identify human Treg cells, especially when
combined with CD25. Thus, Low surface expression of CD127 allows for reliable
separation of human Treg (CD25+ CD127−) from effector T cells (CD25+CD127+) (87).
Together, the aforementioned studies describe our current understanding of Treg cells,
their development and their homeostasis. However, a substantial amount of is
beginning to unravel their function in aged mice and humans.
13
2.3. Treg functionality and specialization
Treg suppressive ability is a key factor for maintaining immune homeostasis. Extensive research has uncovered that FoxP3+ Treg utilize both secreted and cell surface molecules to elaborate their immune suppressive effects. Treg suppression can be mediated via expression of several regulatory molecules: CTLA-4, CD39, CD73, LAG-3,
TIGIT and GITR (71). Further, Treg also express several immune-suppressive
cytokines, such as: IL-10, TGF-ß and IL-35 (88, 89). Treg can also act on target cells
inducing their cytolysis via production of Granzyme B (90).
Notably, functionality of Treg not only depends on production and expression of
regulatory mediators but also on their ability to modulate their phenotype in response to immunological stimuli. Interestingly, the concept of functional specialization of Treg cells was recently introduced into the field (91). The concept states that specialization can be
induced in Treg cells through the activation of transcription factors as well as differential
expression of chemokine receptors under the effect of certain biological stimuli and in
conjunction with Foxp3 expression. Thus, leading to the emergence of new distinct
tissue-specific Treg cell which can play a suppressive or an effector role relevant to its
phenotype (92).
For example, the expression of Tbet in Treg under Th1 polarization conditions leads to the induction of CXCR3, which promotes Treg migration to the site of infection (93).
Similarly, the expression of BCL6 and CXCR5 in a subset of Treg supports the existence of T follicular regulatory cells which may function to suppress Tfh and GC B cells during the germinal center reaction (94). Recently, two distinct subsets of Treg with
14
unique homeostatic phenotype were identified by Campbell et al. CCR7(hi) CD44(lo)
CD62L(hi) Treg maintained by the production of IL-2 in secondary lymphoid tissues and
CD44(hi) CD62L(lo) CCR7(lo) Treg localized to non-lymphoid and are maintained via
the co-stimulatory receptor ICOS (inducible co-stimulator) (95). Thus, it is evident that
Treg can acquire functional plasticity in response to environmental stimuli which enable
them to control different types of inflammatory responses.
The role of Treg has been extensively investigated in many inflammatory conditions.
Despite the beneficial regulatory effect of Treg in limiting chronic inflammation and
deleterious immunopathology, Treg can also play a detrimental role promoting pathogen
persistence and limiting effector T cells responses (96). Thus, Treg seems to play a
dual role balancing both protective and aberrant hyperactive immune responses.
2.4. Treg in aging
Aging is associated with increased risk of chronic infections, cancer and loss of
vaccination responses. Although T cell intrinsic defects play a major role in the decline of immune responses with age, accrual of immune-suppressive Treg are suspected to be major contributors as well. Findings from human and mouse studies provide solid
evidence for age related changes in Treg homeostasis (97-99). It is generally accepted that accumulation of Treg cells with age is due to their enhanced survival and specifically the downregulation in pro-apoptotic molecule Bim (100). Additionally, recent work has shown that ICOS contributes to maintenance of Treg with age, likely by
inhibiting Bim-mediated death (101). Functionally, in vitro and in vivo studies have reported similar or increased suppressive capacity of aged Treg compared to their
15 young counterparts which might be model dependent (97, 98, 102). However, many questions about how Treg can regulate immune responses in aged animals need to be answered. Taken together, a significant role for Treg cells in regulating immune responses with age is increasingly appreciated. We will discuss the role of Treg in aging in the upcoming sections entitled Treg homeostasis with age and Treg function with age within chapter 2.
3. Cytomegalovirus (CMV)
Cytomegalovirus (CMV) is a ubiquitous beta-herpesvirus infecting 60 to 90% of adult individuals (103). Margaret Smith was the first to describe murine cytomegalovirus
(MCMV) and human cytomegalovirus (HCMV) in 1954 and 1956, respectively (104,
105). The virus was first called salivary gland virus (SGV) (104). When the virus enters the host target cell, viral genome enters the nucleus, followed by a process of replication of viral DNA and viral gene expression. The process ends in the assembly of a new virion that is ready to spread into a new host cell via lytic replication. CMV can infect many cell types including epithelial, endothelial, smooth muscle, and connective tissue cells, as well as specialized parenchymal cells in multiple tissues.
The virus is known to evade eradication by modulating the antiviral immune system using its own CMV-encoded proteins. For example, MCMV encodes a viral C-C chemokine MCK2 which facilitates viral dissemination. Indeed, MCK2 resulted in the recruitment of myeloid cells to the site of infection during early stages of viral entry.
Patrolling monocytes then become infected with the virus and, following their circulation, naturally disseminate the virus to multiple sites, including salivary glands, enabling
16
widespread distribution of the virus to multiple tissues (106-108). In addition, recruitment of CCR2 expressing monocytes in response to viral MCK2 has been shown to inhibit the differentiation and cytolytic activity of viral specific CD8 T cells via iNOS-mediated
NO production (109). On the other hand, HCMV expresses proteins like CD55 and
CD59 which target the complement system leading to inhibition of complement fixation, cell lysis and enhanced replication in fibroblasts (110). Together, diverse mechanisms are utilized by CMV to support viral replication, spreading, and targeting the immune system at multiple levels including recruitment of immune effector cells, complement activation and effector T cell responses.
3.1. Acute CMV
In general, viral infection typically activates both innate and adaptive arms of the
immune system, whose function is to ultimately contain, control or eradicate the viral
infection. However, in all cases and despite induction of strong immune responses, the virus is never cleared from the host and establishes a life-long latent infection. A major complication for viral persistence is the reactivation from latency which often ends in re- initiation of productive virus replication resulting in life-threatening diseases in immunosuppressed individuals.
The innate immune system is induced rapidly after CMV infection. The detection of virus is carried out by activation of Toll-like receptors (TLRs) which recognize viral products.
After ligation, several innate immune cells are activated including DCs, macrophages and NK cells. For example, TLR3 can recognize double-stranded RNA (dsRNA) and
TLR9 recognizes double-stranded DNA unmethylated at CpG motifs (111). Once
17 activated, a cascade of signaling pathways result in the production of alpha/beta interferon (IFN-α/β) by DCs and macrophages leading to the activation of natural killer
(NK) cells. Notably, using depletion and adoptive transfer approaches it has been shown that NK cells protect against MCMV (112, 113). More importantly, the strain susceptibility of MCMV in BALB/c mice compared to C57BL/6 has been attributed to the lack of Ly49H activation receptor on NK cells which binds the viral m157 protein resulting in potent NK cell activation and lysis of virally infected cells (114).
Together, TLR signaling is critical for NK cell activation during early MCMV infection. In addition, engagement of Ly49H on NK cells by m157 is indispensable for the control of the initial magnitude of viral replication and dissemination.
Several studies investigated the role of antibodies during CMV infection. Their role remains controversial. For example, antibodies seem not to play an important role in the control of acute MCMV replication. However, MCMV-specific antibodies are able to inhibit viral dissemination. Indeed, primary CMV infection results in the production of neutralizing antibodies against gH and gB, two viral proteins that are critical for viral attachment to host immune cells (115, 116). These neutralizing antibodies controlled viral dissemination into endothelial/epithelial cells (117). Additionally, the transfer of antibodies from HCMV sero-positive individual into newborn infants at risk of infection provided protection against HCMV infection (118). Nonetheless, future studies are needed to fully evaluate the role of antibodies in control of acute CMV infection.
T cells are critical for the control of CMV infection. Early work from Reddehase et al. indicated that CD8 T cells can prevent early replication of the virus in the lungs and the
18 adoptive transfer CD8 T cell from MCMV infected into immune-compromised mice was able to limit viral replication of an established lung infection (119). Additionally, epitope- specific CD8+ T cells are able to control acute MCMV infection in the spleen (120).
In humans, transfer of CMV-specific CD8 T cell clones from donors of bone marrow transplants were sufficient to prevent infection in recipients of allogeneic marrow transplantation after transplantation (121, 122). Although CD8 T cells can control the virus in many organs during MCMV infection they have limited control in the salivary gland due to the CMV-induced downregulation of MHC class I molecules in CMV infected cells in this organ (123). This CMV-driven loss of class I MHC expression makes the salivary gland somewhat of an immune privileged site, however, IFN-y production from CD4 T cells can partially substitute for CD8+ T cells and limit viral load in the salivary gland (124).
In contrast, to the role of effector CD4+ IFN-y-producing cells in controlling viral replication, CD4+ regulatory T cells (Foxp3+) and Foxp3- IL-10+ cells seem to have a negative impact. Indeed, depletion of Treg prior to acute infection resulted in enhanced
T-cell activation and reduced viral titers in the salivary glands (125). Additionally, specific deletion of IL-10 in CD4+ and not Foxp3+ T cells resulted in elevated IFN-y production by T cells and reduction in viral load in the salivary glands (125). Thus, CD4
T cells are pivotal for efficient viral control, while both Treg and IL-10 contribute to the ineffective control of infection resulting in CMV persistence in the salivary gland.
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3.2. Latent CMV
CMV latency is defined by the presence of viral genomes in the absence of detectable
replication with the ability of the viral genome to reactivate (126). Latency can be
established in multiple organs and different types of cells including CD34+
hematopoietic progenitor cells, myeloid cells, alveolar macrophages, epithelial cells,
endothelial cells in the kidney, liver, heart, and spleen (127-130). Establishment of latency is a major goal for the virus to ensure persistence and transmission. To achieve such latency, CMV has developed several strategies. For example, HCMV encodes an
IL-10 homologue. Viral IL-10 shares minimum similarity to human IL-10 (27%) (131).
Although v-IL-10 binds to host IL-10R with limited affinity compared to h-IL-10, this binding enables the virus to evade host immune system recognition by modulating expression of MHC II on antigen presenting cells and by downregulating expression of
TNFα by monocytes (132-134). It is important to note that MCMV does not encode its
own IL-10 homolog but instead uses cellular IL-10 to modulate host immunity (135).
Another viral protein encoded by HCMV is gpUS2, which interferes with CD8 and CD4
T cells by targeting MHC class I and II for proteasome degradation (136). Once the
virus has evaded the recognition by the immune system and established latency it starts
to reactivate periodically. These periodic reactivation events are tightly controlled by
viral-specific CD8+ T cells for most tissues, but likely by viral-specific CD4+ T cells in
the salivary gland. Clearly, future studies will be needed to elucidate the cellular and
molecular mechanisms that regulate the maintenance of the viral genome during
latency and those that trigger reactivation.
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3.2.1. Cellular and molecular mechanisms controlling CMV reactivation from
latency
It is widely accepted that chromatin remodeling of the viral major immediate-early
promoter (MIE) under different stimuli (e.g. cytokines) is a major inducer of reactivation
from latency (137). Notably, in vitro allogeneic stimulation of latently infected peripheral mononuclear cells resulted in full reactivation of HCMV. The cellular source in blood for viral reactivation was identified as latently infected monocytes expressing dendritic cell
markers (138). Additionally, Hummel et al. showed that allogeneic transplantation
resulted in the expression of the MCMV ie1 gene which was accompanied by increased
expression of transcripts encoding inflammatory cytokines, including TNF-α, IL-2, and
IFN-y. These data suggested that the induction of these cytokines drives the activation of the major immediate-early (MIE) gene and initiating reactivation from latency.
Interestingly, the authors were able to show that injecting TNF-α alone into latently infected mice was able to induce ie1 gene expression but did not lead to full viral reactivation (139). Thus, factors in addition to TNF-α likely control viral reactivation in vivo. Notably, full reactivation of MCMV was reported in the lungs following allogeneic transplantation combined with immunosuppression (e.g. irradiation) (140).
Thus, these studies provided evidence that reactivation from latency is potentially controlled via a multistep process which is initiated by induction of ie1 gene expression followed by full lytic replication in the presence of immunosuppressive environment.
While these studies argue that latency is controlled at transcriptional levels, there is also periodic, low-level reactivation from latency that is controlled primarily by CD8 T cells,
21 but also by CD4 T cells and NK cells. Indeed, the combined depletion of T cells and NK cells resulted in the reactivation of MCMV in multiple tissues (141). Thus, it is likely that latency and reactivation from latency is a tightly regulated process regulated by adaptive immune cells and cytokine production which combine to control transcription of immediate early viral genes. Nonetheless, mechanisms regulating CMV reactivation from latency remain poorly understood
3.2.2. CMV in aging
CMV infection is well tolerated by an immune competent host. However, CMV can result in great morbidity and mortality in organ and bone marrow transplant patients and HIV patients (142) (103). In addition, CMV is envisioned as one of the driving forces of the age-related immune dysfunction and inflammaging (143, 144). Also, CMV has been associated with multiple age-related pathologies such as type 2 diabetes, cancer and cardiovascular disease (145).
CMV infection persists latently in the host and reactivates periodically (146). Repeated reactivation of the virus drives the expansion of CMV-specific CD8＋ T cell which can constitute up to 40% of CD8+ T cells in the peripheral blood of HCMV- seropositive individuals (147, 148). Although such expansion is expected to protect against reactivation and perhaps elimination of the virus, these cells have also been thought to be inducers of excessive inflammation. For example, stimulation of human PBMCs with
HCMV antigens resulted in increased production of IFN-y which was eight times greater in very old than in young subjects, suggesting that CMV infection might play detrimental effect with age by contributing to the age-related pro-inflammatory environment.
22
Despite the presence of functional HCMV-specific T cells with age, higher frequency
and greater viral load of CMV was reported in elderly (149). Related to the higher viral
load is that both the frequency and magnitude of CMV reactivation are increased with
age (142). These studies suggested that aging might favor the induction of a
suppressive environment that can modulate T cell responses and facilitate viral
persistence and reactivation. Nonetheless, major gaps remain in our understanding of
the cellular and molecular regulatory mechanisms controlling immune responses and
viral reactivation from latency with age.
3.2.3. Regulatory mechanisms controlling CMV
(i) Treg in CMV
CMV causes a persistent, lifelong latent infection despite a robust immune response. It
is unclear whether Teg cells have an impact on the insufficient host immune response
against the virus and whether they contribute to viral persistence. Clinical studies in
humans have allowed for important insights into the role of Treg in modulating immune
responses to CMV infection. For example, in vitro studies by Aandahl et al.
demonstrated that the depletion of Treg CD25+ cells from peripheral blood of CMV
sero-positive humans resulted in enhanced IFN-y and IFN-α production by syngeneic
cytomegalovirus (CMV)-specific CD8 T cells (150). These data suggested that Treg
might be limiting CD8 T cell responses. In agreement, murine studies indicated that
Treg interfere with an effective anti-mouse immune response and the depletion of Treg resulted in the control of viral persistence in the salivary gland (125). Thus Treg seems to negatively regulate immune responses to CMV favoring viral persistence. However,
23
the role of Treg in viral reactivation from latency is unclear. We will discuss the role of
Treg in latent MCMV in chapter 3.
(ii) IL-10 in CMV
IL-10 is a potent anti-inflammatory cytokine that regulates host immunity to acute as
well as chronic infections. IL-10 can also be exploited by pathogens to modulate the
immune response in favor of persistence in the host. Prior data shows that IL-10 controls CMV viral replication and regulates the establishment of latency (135), this has both benefits and negative consequences. For example, IL-10 has been shown to negatively regulate cellular immunity particularly in the salivary glands in favor of viral persistence (135). Indeed, interfering with IL-10R signaling restricted viral replication in the salivary gland (135). Likewise, the absence of IL-10 promoted the crosstalk between
NK cells, myeloid dendritic cells (mDCs) and in virus-specific CD4+ T cell responses resulting in reduced virus persistence in the salivary glands (151). To further investigate
the cell-specific role of IL-10 in MCMV, a recent study has shown that the specific
deletion of IL-10 in CD4 T cells, but not Foxp3+, cells led to enhanced antiviral T cell
responses and restricted MCMV persistence in salivary glands (125).
In contrast, other studies suggested a beneficial effect for IL-10 during acute MCMV.
For example, IL-10 controlled the weight loss and inflammatory cytokine production in
acute MCMV infection (152). The findings suggest that the role of IL-10 in MCMV is
complicated and might have differential effects on viral load based on IL-10 cellular
sources and the site of infection. We will discuss the role of IL-10 in latent MCMV in chapter 3.
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4. Interleukin-10 (IL-10)
IL-10 is a regulatory cytokine, with potent anti-inflammatory effects. The regulatory
effect of IL-10 could play a beneficial role (e.g. resolution of inflammation and controlling
excessive immunopathology) or a detrimental role (e.g. persistence of pathogens and
inhibition of antitumor immunity) on host immunity. The discovery of IL-10 was first
made in 1989 by Mosmann and co-workers. The cytokine was described first to be produced by Type 2 helper cells (Th2) with a potent inhibitory effect on interleukin (IL-2)
and interferon (IFN-y) production and proliferation of Type1 helper cells (Th1) (153,
154).
IL-10 was initially referred to as “cytokine synthesis inhibitory factor” (CSIF). The gene
encoding IL-10 is located on chromosome 1 in humans and mice and covers a total of
5.1 kb pairs or 4.7 kb pairs, respectively. The IL-10 gene comprises five exons which
encode 178 amino acids. Both human and mouse IL-10 share 75% homology in the
amino acid sequences. As stated before, human IL-10 is known to share homology with some open reading frames in human viruses like Epstein–Barr virus, Herpes type 2
virus, Parapoxvirus, and Cytomegalovirus (155). To date, multiple cytokines have been shown to share structural similarity to IL-10 and collectively are known as IL-10 family
members (e.g. IL-10, IL-19, IL-20, IL-22, IL-24, and IL-26) (156, 157).
Since its discovery, extensive work with age has been focused on the cellular sources of IL-10 and their biological relevance. To date, multiple cells have the capability of producing IL-10, including adaptive immune cells (e.g T and B cells) and innate immune cells ( e.g dendritic cells (DC), γδ T cells, NK cells, mast cells, neutrophils, eosinophils),
25 keratinocytes (158), epithelia and some tumor cells (159). Together, IL-10 has a broad range of cellular sources and its production in subjected to various regulatory mechanisms that vary between tissues and under different inflammatory conditions.
4.1. Transcriptional regulation of IL-10 production in CD4 T cells
IL-10 expression is governed by multiple layers of regulation, including both transcriptional and post-transcriptional mechanisms. These mechanisms are influenced by a myriad of environmental stimuli (e.g. cytokines) as well as by tissues or cell types expressing IL-10. Importantly, the sequence identity and location of transcription factor binding sites in the IL-10 locus are well conserved in both humans and mice.
Understanding the transcriptional regulation of IL-10 is critical given the major roles of
IL-10 in controlling immune homeostasis and the potential therapeutic application of IL-
10 in multiple inflammatory diseases.
4.1.1. TCR mediated IL-10 expression
Following TCR stimulation multiple signaling pathways are initiated in CD4 T cells resulting in the induction of IL-10 expression through the activation of downstream transcription factors including AP-1 and c-Maf downstream of ERK, nuclear factor of activated T cells (NFAT) and NF-κB. After translocating from the cytoplasm into the nucleus NFAT is known to interact with AP-1 (compromised of JunB and c-Jun) and other transcriptional partners to promote IL-10 expression in CD4 T cells (160). In addition to AP-1, c-Maf activation in an ERK-dependent manner induces IL-10 expression in CD4 T cells. Further investigation indicated that c-Maf is able to bind and
26 trans-activate the IL-10 promoter (161, 162). Additionally, a role for NF-κB in IL-10 expression has been demonstrated in different Th subsets, including Th17, Th2 and
Th1 (163), however this pathway has been less characterized compared to NFAT1 and
AP-1.
4.1.2. Cytokine mediated IL-10 expression
The induction of IL-10 expression is complex, it can be controlled by multiple distinct cytokines and different CD4 T cell subsets. For example, IL-6, IL-21, IL-27, type 1 interferons, IL-12 and TGF-β can induce IL-10 production. Interestingly, IL-10 can induce its own expression (164). Further, the regulation of IL-10 expression following cytokine stimulation involves the activation of several STAT molecules and distinct transcription factors. Indeed, STAT3 is essential for IL-6-mediated IL-10 production
(165). Further, IL-27 as IL-12 cytokine family member can induce IL10 gene expression in a STAT3/STAT1 dependent manner (166-168). Similarly, IL-21 signaling via STAT3 is also required for the expression of IL-10 (169). Mechanistically, STAT3 activation under IL-21 and IL-27 has been shown to activate c-Maf expression. Interestingly, AHR synergizes with c-Maf to promote the production IL-10 in Tr1 cells. AHR forms a protein complex with c-Maf, to jointly trans-activate IL-10 promoter (170). Additionally, the expression of IL-10 in Tr1 cells is further regulated by ICOS expression. Indeed, ICOS- deficient T cells show reduced IL-10 production and c-Maf expression under IL-27 stimulatory conditions (171).
Another pathway that regulates IL-10 expression under the effect of IL-27/STAT3 is
Blimp-1. Indeed, Blimp-1 downstream of IL-27 is able to mediate IL-10 induction in CD4
27
T cells (172). In Treg cells, Blimp-1 and IRF-4 jointly induce the expression of IL-10
through direct binding to IL-10 promoter (173). Notably, BCL6 a known transcriptional repressor of Blimp-1, has been shown to negatively regulate IL-10 expression in CD4 T cells (174). Importantly, the negative regulation of IL-10 expression in CD4 T cells by
BCL6 seems to be independent of Blimp-1(175).
Similar to STAT-3, STAT-4 plays a role in the induction of IL-10 production in CD4 T
cells. Indeed, STAT-4 activation results in the development of IL-10/IFN-y double
producing cells under the effect of IL-12 and high antigen load (176). Further, SMAD
molecules in response to TGF-ß can induce IL-10 expression. Indeed, SMAD4 has been shown to activate IL-10 expression in Th1 cells (177), while SMAD3 been shown to induce IL-10 production in response to TGF-ß in Th2 cells (178).
In summary, several mediators are involved in the regulation of IL-10 expression in CD4
T cells including, TCR signaling and cytokine signaling through STAT/SMAD molecules promoting the activation and nuclear localization of several transcription factors
(Figure1). These different circuits allow the accessibility of IL-10 in response to multiple environmental stimuli, likely acting in different cell types or differentiation states to be able to combat various forms of inflammation. However, more work is necessary to understand the environmental contexts that promote IL-10 with age. In addition, as most environments possess multiple cytokine inputs, it is unclear how these multiple inputs contribute to IL-10 production in vivo. Although distinct CD4 T cell subsets require different stimulatory conditions, almost all CD4 T cell subsets have the capability to express IL-10 under differential regulatory mechanisms.
28
promoter
Figure1. Molecular events regulating IL-10 expression in CD4 T cells. IL-10 expression
in CD4 T cells is regulated by signals induced by TCR stimulation and cytokines acting via STAT/SMAD molecules and other transcription factors.
4.2. IL-10/IL-10R and signaling pathway
Similar to mice human IL-10 is a 35 kD homodimer consists of 178 amino acids, with an
N-terminal hydrophobic leader sequence 18 amino acids long (179). IL-10 serum half- life is between 1.1 and 2.6 hours (180). IL-10 signals through IL-10 receptor
(heterotetramer) which is composed of two unique chains of IL-10R1 (ligand binding), expressed on most hematopoietic cells at low levels, with higher levels on CD4+ memory than on naive T cells (181). It is also up-regulated by various stimuli on non- hematopoietic cells, such as fibroblasts and epithelial cells. IL-10R2 chains are constitutively expressed as two IL-10R2 (signal transduction) chains expressed on immune cells in addition to epithelial cells and keratinocytes. IL-10R2 is utilized as a
29
receptor by other cytokines from the IL-10 family, including IL-22, IL-26 and IL-28 and
IL-29 (181).
The activation of IL-10R through the binding of its ligand leads to the activation of downstream signaling involving Janus tyrosine kinases Jak1 constitutively associated with IL-10R1 and Tyk2 which is constitutively associated with IL-10R2 chain (182).
Ligand–driven phosphorylation of Jak1 promotes the phosphorylation and activation of
STAT3 and STAT1. STAT3 homodimers upon activation by IL-10 translocate into the nucleus and bind to STAT elements on promoters of various genes, including IL-10 itself and the suppressor of cytokine signaling (SOCSs) genes, amongst others. Of note, compared to other STAT3-activating cytokines (ie IL-6 and IL-21), IL-10 rapidly activates a more sustained phosphorylation of STAT3 while IL-6 mediated STAT3 activation is transient.
One explanation for this differential activation of STAT3 under IL-10 versus IL-6 stimulation might be the differential induction of SOCS3. Notably, SOCS3 among other
SOCSs molecules plays a major role in the negative regulation of various cytokines including IL-10, IL-6 and IL-21. SOCS3 is induced upon cytokine receptor activation via
STAT3-dependent mechanism. Such induction results in the initiation of a negative feedback loop resulting in the inhibition of the JAK/STAT signaling pathway. However,
SOCS3 can only interfere with JAK/STAT signaling pathway in the presence of receptors, such as gp130. Indeed, binding of SOCS3 to the Src homology phosphatase-
2 (SHP-2) binding domains of gp130 is required for the initiation of SOCS3 regulatory effect (183). Thus, the fact that IL-10 receptor chains do not contain a consensus motif
30
(SOCS box) for suppressor of cytokine signalling-3 (SOCS3) explains the loss of the negative regulatory effect of SOCS3 on IL-10/STAT3 sustained activation compared to
IL-6/STAT3 (184, 185).
The role of STAT3 in the IL-10 signaling pathway is indispensable. Indeed, previous work indicated that the specific deletion of STAT3 in myeloid lineage (macrophages, neutrophils, and dendritic cells) leads to the development of colitis similar to IL-10-/- mice
with enhanced Th1/IFN-y responses (186). The authors in the study concluded that the
increased production of IFN-y from CD4 T cells in STAT3 deficient mice would feedback
on myeloid cells and enhance the severity of colitis in the absence of IL-10/STAT3
signaling (187). In contrast to STAT3, little is known about the role of STAT1 in the
biological effects of IL-10. A study by Meraz et al. indicated that in contrast to STAT3,
deficiency of STAT1 did not result in inflammatory colitis. More importantly,
macrophages from STAT1 deficient mice were completely responsive to IL-10 (188).
Thus, the role of IL-10-driven STAT1 remains to be determined.
4.3. Biological effects of IL-10
IL-10 has a plethora of both negative and positive effects on immune responses.
Indeed, IL-10 can negatively regulate immune responses via inhibition of production of
tumor necrosis factor-α (TNF-α), IL-1, IL-12, IL-6 and granulocyte–macrophage colony-
stimulating factor (189). Additionally, IL-10 can inhibit nitric oxide (NO) production, and
expression of class II MHC and costimulatory molecules such as CD80/CD86 on
dendritic cells and macrophages (190). IL-10 can also inhibit T cells by regulating IL-2,
TNFα, and IL-5 production (191, 192), as well as enhancing the expression of soluble
31
TNFα receptors and IL-1 receptor antagonist (IL-1RA) (193, 194). Together these functions of IL-10 decrease adaptive immune responses.
In contrast, IL-10 has a stimulatory effect on NK cells, enhancing their proliferation, cytokine production, and cytotoxicity (195). Likewise, in vitro studies indicated that IL-10 can promote B cell survival and Immunoglobulin (Ig) class switch (196). Thus, the biological functions of IL-10 are very broad and complex and likely depend on the context of the immune response.
IL-10 plays an indispensable role in the regulation of intestinal homeostasis (197).
Notably, IL-10 production from Foxp3- type 1 regulatory T (Tr1) cells has been shown to
play a critical role in regulating a variety of inflammatory diseases including
development of colitis (198). Additionally, different subsets of T cells like Th1, Th17,
have been shown to experience some plasticity under the effect of inflammatory
environment and acquire the ability to produce IL-10 (199, 200), suggesting that IL-10
production from effector T cells might play a counter-regulatory role against excessive
inflammation.
Tight regulation of IL-10 is also critical to maintain human health. Overproduction of IL-
10 can lead to immune suppression, while underproduction of IL-10 can lead to
autoimmunity. For example, IL-10 produced by Th1 cells prevented excessive
immunopathology upon infection with cutaneous leishmaniasis and toxoplasma gondii
(199, 201). Also, IL-10 from gamma delta T cells prevented excessive tissue injury in
mice infected with Listeria (202). In contrast, excessive IL-10 production can inhibit the
immune responses during infections with Mycobacterium sp. and lymphocytic
32
choriomeningitis virus (LCMV) promoting chronic infection (203-206). On the other hand, IL-10 deficiency resulted in excessive inflammatory colitis by failing to limit Th1/17
responses (197) These data collectively show that dysregulated expression of IL-10 has
serious consequences.
4.4. Aging and IL-10
Aging is associated with low grade systemic inflammation, which contributes heavily to increased mortality and morbidity with age (3, 207, 208). Given its known anti-
inflammatory role, IL-10 might represent a key for balancing inflammaging. However,
the role of IL-10 with age is poorly understood. For example, the possession of -1082G
genotype, which is linked with IL-10 high production, is significantly increased in
centenarians (209), and similarly more in middle-aged controls than in age-matched patients with myocardial infarction (210). While an increase in serum IL-10 levels has been documented in aged humans (211), the cellular sources of IL-10 were not fully addressed. Thus, to date we lack full understanding of the cellular sources of IL-10 with age.
In addition to its role in balancing inflammation, IL-10 limits responses to infections and vaccines (212, 213). Interestingly, a balance between IFN-γ and IL-10 plays a role in vaccination responses with age. Indeed, the IFN-γ:IL-10 ratio correlates with protection against influenza infection. However, the IFNγ:IL-10 ratio in response to influenza virus challenge declined with age resulting in increased risk to influenza infection in vaccinated older adults (214, 215). Thus, strategies aimed at maximizing anti- inflammatory responses to vaccination, enhancing IFN-γ production and antagonizing
33
IL-10 responses are expected to support protection following vaccination with age.
Together, more research is needed to determine the cellular sources and effects of IL-
10 on immune responses with age. We will discuss these potential sources and functions of IL-10 in aging in chapter 4.
5. T follicular helper (Tfh) cells
Robust germinal centre (GC) formation directed by T follicular helper (Tfh) and B cell interaction is absolutely required for the generation of T-dependent humoral immunity
(216). Tfh cells are specialized CD4+ T cell subset that differentiates from naïve cells in response to antigen challenge (217). This differentiation of Tfh cells is indispensable for the activation of B cells, antibody class switching and affinity maturation which results in the production of high affinity circulating antibodies as well as formation of memory B cells (218). The generation of Tfh cells is an integrated process directed by transcriptional program that is including T cell receptor (TCR) signaling, T-B cell interaction and the cytokine environment. Indeed, the transcriptional repressor BCL-6 whose expression is promoted by IL-6 and IL-21 driven STAT-3 signaling is essential for the generation of Tfh cells (219).
Additionally, the cell surface marker CXCR5 is critical for Tfh localization to the GC and allows for B/T cell interaction (220). CXCL13 production by stromal cells binds to
CXCR5 on B and Tfh cells and promotes their migration to the B cell zone, where Tfh cells provide help in the form of CD40L and IL-21 production to GC B cells resulting in efficient antibody production (221). On the other hand, B cells provide help to Tfh cells through the expression of ICOS-L which is binds to ICOS and promotes Tfh
34
differentiation, BCL-6 expression and maintenance of Tfh cells (222). In addition, lymphocyte activation molecule-associated protein (SAP)/ signaling lymphocyte activation molecule (SLAM) signaling further stabilizes the crosstalk between Tfh and B cells. This interdependent relationship between Tfh and B cells is required for optimal B cell activation and GC formation (223). Given the critical role played by Tfh cells in shaping immune responses towards infections and to support the efficiency of vaccination, any changes that modify the function of Tfh cells or their differentiation could have a major impact on humoral responses. Notably, multiple age-related defects have been reported in Tfh cells which partly contributed to high risk of infections and loss in vaccination responses in elderly individuals. Together, Tfh generation and function are carfeully regulated via interlinked signals including, transcription factors, cytokines as well as expression of suface markers.
5.1. Aging and Tfh cells
Like other cell types, other studies have documented that aged mouse and human Tfh cells are phenotypically and likely functionally different compared to young Tfh cells.
Indeed, detrimental changes to aged Tfh cells have been reported in both human and
mouse studies. Some of these changes are extrinisic caused by the aging environment
and resulting in defective migration, localization and induction of Tfh cells, while other
are intrinsic changes result in impaired differentiation and function and hence poor B
cell help, reducing their ability to produce protective circulating antibodies. For example,
the aged environment failed to support the recruitment of naïve CD4+ T cells to
35
secondary lymphoid organs and hence the differentiation of Tfh cells resulting in poor
vaccination responses in old mice (224).
Additionally, a previous study showed that aged antigen-specific Tfh cells are defective
in their maturation status. The authors argued that aged Tfh cells are defective in their
homing to the GC because of their lower expression of CXCR5 compared to their young
counter parts (225). Further, enhanced production of IL-10 and lower expression of
ICOS on aged antigen -specific CD4+ T cells resulted in impaired B cells reponses
(225). Althogh this regulatory phenotype of CD4+ T cells have been attriabuted to their
Foxp3 expression, the existence of Foxp3- IL-10 producing cells with age is yet to be clear. Notably, a previous study has shown that aged mice demonstrated delayed clearance of influenza virus compared to young mice. The authors attributed this defect to the dyregulated differentiation of Th1 towards Tfh cells in the lungs of aged infected mice which resulted in increased lung viral load and inability to control primary lung infection as well as enhanced suscepitality to secondary infections (68). However, mechanisms by which Tfh cells altered viral clearance were not addressed in this study.
In humans, circulating CXCR5 + CD4 + T cells are increased in elderly individuals and this correlated with higher levels of IL-21 produced by aged PBMCs compared with young subjects (226). Mechanistically, prolonged activation of STAT-4 in response to IL-
12 in aged CD4+ T cells resulted in increased IL-21 with age (227). Together, several immunologic changes have been described in aged Tfh cells, including defective priming, differentiation and cytokine production. Importantly, knowledge of the age- related defects in Tfh cells and the contribution of the inflammatory environment is
36 critical to further advance our understanding of how aging impacts both infections and vaccine responses.
6. Interleukin-6 (IL-6)
6.1. IL-6 signaling pathway
Human IL-6 was first cloned in 1986 and given the name B cell stimulator factor BSF-2 due to its positive effects on promoting plasma cell growth (228). Multiple cell types can produce IL-6, nearly all stromal cells, vascular endothelial cells, and adipocytes at physiological conditions. Current research provided substantial insight into IL-
6 signaling pathways and its versatile biological activities.
Signaling by IL-6 is complex and involves signaling through cells that express IL-6Rα
(classical pathway) and cells that do not express IL-6Rα (trans-signaling). IL-6 signals through IL-6R which is composed of a unique receptor subunit (IL-6Rα) that binds IL-6.
This chain is mainly expressed on, hepatocytes and immune cells like T cells, neutrophils, and monocytes/macrophages (229). The receptor is also composed of two subunits of (gp130) for signal transduction (230). Notably, gp130 is also shared by several other cytokines like IL-11, IL-27, oncostatin-M and expressed ubiquitously on many cells.
6.1.1. Classical pathway
The binding of IL-6 to its membrane-bound receptor (mIL-6Rα) chain results in the formation of a complex with the homodimer gp130 subunits which triggers a
37
downstream signal cascade. Subsequently, kinases of the Jak family (Jak1, Jak2 and
Tyk2) are activated which leads to the activation and phosphorylation of STAT
molecules. As mentioned previously, IL-6 is known to signal primarily through STAT1,
STAT3 and to a lesser extent STAT2 (231). The phosphorylation of STAT molecules
leads to their dimerization and translocation to the nucleus to activate transcription of
genes containing STAT response elements. Notably, STAT3 activation induces the
suppressor of cytokine signaling 1 SOCS1 (232) and SOCS3 (233). SOCS1 binds to tyrosine-phosphorylated JAK, whereas SOCS3 binds to tyrosine-phosphorylated gp130 resulting in the termination of IL-6 cytokine signaling via a negative feedback loop.
6.1.2. Trans-signaling pathway
IL-6 can signal to cells that do not express membrane-bound receptor (mIL-6Rα). This occurs by IL-6 binding to soluble IL-6R (sIL-6Rα), followed by the IL-6/sIL-6R complex binding to gp130 subunits on the surface of cells, including those that do not formally express IL-6Rα on their surface. The binding of IL-6 to its soluble receptor (sIL-6Rα) can activate the dimerization of gp130 subunits similar to the classical pathway. sIL-
6Rα is generated under two mechanisms including, mRNA alternative splicing, and shedding of the membrane-bound IL-6R mediated by metalloproteases ADAM17 and
ADAM10 (234). Interestingly, a soluble form of gp130 is also produced and can act as
an antagonist for IL-6 trans-signaling pathway (235). Together, IL-6 is unique in
signaling via a membrane bound and a soluble receptor and it remains unclear how these distinct forms of signaling impact the biological effects of IL-6.
38
6.2. Biological effects of IL-6
IL-6 is involved in a broad spectrum of biological activities. Indeed, the production of IL-
6 early in the immune response regulates the production of acute phase proteins like
serum amyloid, C-reactive protein and fibrinogen (236, 237). It is generally accepted
that IL-6 activation through the trans-signaling pathway accounts for most of IL-6
biological activity early in the immune response. Additionally, IL-6 regulates
T cell subsets differentiation including Th17, Th2, Th1, Tfh, and Tr1 cells. For example,
IL-6 plays an indispensable role in Th17 differentiation together with transforming growth factor (TGF-β), IL-1β and IL-23 (238) (239). In Th2 cells, IL-6 enhances the expression of IL-4 and IL-13 (240-242), predominantly through an NFATc2 and c-Maf regulated pathway. In contrast to its positive regulatory effect on Th17 and Th2 cell differentiation, in Th1 cells, IL-6 interferes with IFN-y production by CD4 T cells mainly through upregulation of suppressors of cytokine signaling (SOCS1or SOCS3) (243).
One critical role of IL-6 is to induce the differentiation of a unique subset of cells termed
T follicular helper cells (Tfh). Tfh cells are critical for facilitating germinal center
reactions by providing help for B cells. Notably, IL-6 signaling through STAT3 is critical
for the upregulation of BCL6, the signature transcription factor of Tfh cells (244).
Importantly, in a Bcl6-dependent manner, Tfh cells express CXCR5 which enables them
to localize to GCs where they are in close proximity to B cells during the GC reaction
(245) (246). Although most models suggest that IL-6 is required for Tfh differentiation,
there have been a few reports of an IL-6-independent pathway for Tfh differentiation
(246). Current evidence suggests that this IL-6-independent pathway is predominantly
39
mediated by IL-21. Nonetheless, in most models, IL-6 is critical for the initial
development of Tfh cells.
In addition to its role in promoting Th17 and Tfh cells, IL-6 has also been shown to promote IL-10 production from CD4 T cells (247). Indeed, early studies investigating
Th17 cell differentiation have shown that both TGF-β and IL-6 drive the activation of
STAT3 leading to their co-production of IL-10 (162). Additionally, IL-6 was required for the induction of CD4 type 1 regulatory T cells (Tr1) and their production of IL-10 (247).
Further, IL-21/c-Maf expression was indispensable for the induction of IL-10 in Tr1 cells under IL-6 stimulatory conditions (247). Thus, based on the biological context, IL-6 plays a complex role in shaping the immune response.
6.3. Aging and IL-6
As a signature of the “inflammaging” process, several groups, including ours, have
reported increased systemic levels of IL-6 in aged mice and humans (101, 248-250).
Such increases in IL-6 are implicated in the pathogenesis of multiple age-related diseases including atherosclerosis, osteoporosis, sarcopenia, and cardiovascular disease, resulting in high mortality in the elderly (251-254). Importantly, increased
production of IL-6 with age is a major risk for age-associated frailty resulting in
functional decline and serious adverse health problems. A previous study has shown
that frail elderly had higher levels of IL-6 than non-frail, age-matched individuals (255).
Additionally, using IL-10 deficient mice as a model of frailty, old IL-10-/- mice with a frailty
phenotype had elevated IL-6 levels compared to the control WT mice (256). This latter
study also suggested a counter-regulatory role for the anti-inflammatory cytokine IL-10
40
in regulating IL-6 inflammation with age. Thus, findings from both human and mouse
studies provided evidence for the strong association of IL-6 to frailty and its adverse
outcome in elderly.
Interestingly, available data suggest that IL-6 and IL-10 may play important roles in
balancing inflammaging. The fact that centenarian males had an IL-10 polymorphism
associated with high IL-10 production in one study (209) and IL-6 serum levels were
increased among the same age group in another study is very intriguing (248). Although
measurements in both studies were independent, the findings suggested the possibility
that enhanced IL-10 levels might be a compensatory mechanism driven by increased
IL-6 to guard against excessive inflammation with age. Thus, investigating the role of IL-
6 in the induction of IL-10 with age may shed light on the development of more specific
therapeutic approaches targeting age related diseases, especially those where IL-6
plays a dominant role. Our data in chapter 4 will shed light on the relationship between
IL-6 and IL-10 in aged mice.
7. Interleukin-21 (IL-21)
7.1. IL-21 signaling pathway
Both IL-21 and its receptor were discovered in 2000 (257). IL-21 is produced primarily by activated CD4 T cells, natural killer T cells, and follicular T helper cells. The
heterodimeric receptor for IL-21 is composed of (IL-21R αβ) and the common cytokine
receptor (γc) chain, which is also a shared chain for IL-2, IL-4, IL-7, IL-9, and IL-15. IL-
21R can be expressed on many cells including, dendritic cells, macrophages, epithelial
41 cells, and keratinocytes. In addition to B, T, and NK cells (258). Signaling through IL-
21R is well known to function primarily by activating STAT transcription factors as well as ERK and PI3-kinase-Akt. IL-21 is known to activate STAT3 and STAT1 and to lesser extent STAT5 (259).
7.2. Biological effects of IL-21
As expected from the broad expression of its receptor, IL-21 is a pleiotropic cytokine with multiple actions on B cells, NK cells and CD4 T cells. On B cells, IL-21 seems to play opposing roles. For example, IL-21 plays an inhibitory effect on human and mouse
B cell proliferation and survival (260). However, it can enhance the differentiation of B cells into plasma cells in a STAT3 dependent manner (261). Additionally, mice with genetic loss in IL-21 receptor expression had reduced levels of serum IgG1 and high levels of IgE (262), similar to what has been reported in IL-21-deficient humans (263), suggesting an indispensable role for IL-21 in promoting immunoglobulin production.
Together, effects of IL-21 on B cells may be modulated by the differentiation status of B cells. Further, IL-21 plays a positive regulatory effect on NK cells. Indeed, NK cells experienced reduced cytolytic activity in humans deficient in IL-21R (264). Additionally,
IL-21 promoted survival and upregulated the expression of several surface receptors, including NKG2A/C/E on human NK cells (265), suggesting that IL-21 is enhancing survival and functionality of human NK cells.
In CD4 T cells, IL-21 can contribute to Th17 differentiation along with transforming growth factor (TGF-β) and IL-6 (266). Similarly, IL-21 together with IL-6 has been shown to be fundamental in Tfh development (245). Mechanistically, IL-21 is required for high
42 level expression of BCL6, which is important for Tfh differentiation and their sustained expression of CXCR5. Indeed, IL-21-KO mice showed reduced expression of BCL6
(267) and CXCR5 surface expression on CD4+ T cells was reduced in the absence of
IL-21 (268). In addition to its effect on BCL6 in Tfh cells, IL-21 can regulate other transcription factors involved in Tfh differentiation. For example, c-Maf, another transcription factor involved in the differentiation of Tfh cell is regulated by the expression of IL-21. Indeed, mice lacking c-Maf in the T cell compartment had reduced numbers of Tfh cells with limited secretion of high-affinity antibodies (269). Thus, IL-21 plays a significant role in promoting Tfh and Th17 differentiation.
In contrast to its role in the differentiation of inflammatory Th17 cells, IL21 can exert immunosuppressive effects via induction of IL-10-producing cells. Indeed, IL-21 is known to induce the expression of IL-10 in CD4 T cells (169). Several cytokines were shown to govern the production of IL-21 from T cells and subsequently the induction of
IL-10 in an autocrine manner. For example, IL-27 was shown to stimulate the production of IL-21 from type 1 regulatory T cells. The autocrine effect of IL-21 was then required to enhance and maintain the production of IL-10 from Tr1 cells (270). Similar effect of IL-
21 on IL-10 production was reported in IL-10-secreting type 1 regulatory T (Tr1) cells stimulated under IL-6 (247). Thus, the effects IL-21 on CD4 T cells differentiation can be complex and vary depending upon potential environmental stimuli.
7.3. Aging and IL-21
Aging is associated with severe alterations in immune function which contribute to increased susceptibility to infections and reduced response to vaccination. B and Tfh
43 cells are major players in regulating immune responses to vaccination with age. Given the role played by IL-21 in dictating both B and Tfh cell responses, some studies investigated the production of IL-21 with age. The data remains controversial. For example, a previous study demonstrated that aged PBMCs produced significantly higher levels of IL-21 compared with young individuals (226). In agreement, another study reported that CD4+ T cells from aged individuals secreted significantly higher levels of IL-21 on priming with dendritic cells (DCs) (227). Further examination revealed that the response of aged CD4+ T cells was due to alteration in IL-12/STAT4 pathway with age. In contrast, aged memory CD4 + T cells expressed lower levels of IL-21 after stimulation with anti-CD3/ anti-CD28 (271). The discrepancy in the findings might relate to approaches utilized and different experimental conditions For example, CD4 T cells were cultured with dendritic cells for 6 days, while the stimulation of CD4 T cells with anti-CD3/anti-CD28 lasted for 48hrs only. Nonetheless, whether IL-21 levels are altered with age or not remains unclear.
Summary
Aging is associated with profound immune dysregulation that contributes to the increased morbidity and mortality in elderly. Viral infections including CMV are implicated in accelerating age-related immune dysfunction. It is well established that
Treg and IL-10 play critical roles in regulating immune responses to acute CMV.
However the interplay between Treg and IL-10 producing cells during latent CMV infection is not clear. To date, we lack full understanding of factors contributing to latent
MCMV. More importantly, there is still a gap in knowledge about the role of Treg and IL-
44
10 on controlling the reactivation of latent MCMV. Our data suggest the existence of a
crosstalk between Treg and Foxp3- IL-10 producing T cells that differentially regulates the replication/reactivation of MCMV. Further, we show that the biologic outcome of this
dynamic between FoxP3+ CD4+ Treg cells and IL-10-producing FoxP3- CD4+ T cells depends upon the infected tissue. The data supporting this concept are the subject of chapter 3.
Inflammaging is characterized by increased production of inflammatory cytokines (e.g.
IL-6). On the other hand, production of anti-inflammatory cytokines such as IL-10 is envisioned as a strategy to counterbalance inflammaging. However, the role of IL-10 in aging is not clear. Notably, IL-6 and IL-21 normally promote Tfh development. Further, both cytokines are strong inducers of IL-10 production from CD4 T cells. However, it is unclear how Tfh development and IL-10 production proceed when the expression of IL-
6 and/or IL-21 becomes dysregulated. Our data provides strong evidence supporting the accrual of Tfh10 cells with age. Further, our data provide strong biological relevance for their existence through counterbalancing inflammaging on one hand and on the other hampering vaccination responses with age. The data supporting this concept are the subject of chapter 4.
45
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Chapter 2
T-reg Homeostasis and Functions in Ageing
Treg and Aging
Maha Almanan, Claire Chougnet, and David A. Hildeman
Abstract Aging has a profound impact on immune responses. Both innate and adaptive compartments of the immune system undergo fundamental changes with age. It is well recognized that intrinsic defects, imprinted on T cells, contribute signifi- cantly to age-related diseases. However, recent data show a profound accumula- tion of regulatory T cells in aged mice and humans. The accumulation of these regulatory T cells, and their production of immune suppressive cytokines, likely contributes to immune dysregulation in aging. On the other hand, a great deal of work has shown substantially enhanced inflammation with age, with IL-6 being one of the hallmark inflammatory cytokines associated with this age-driven inflammation. Importantly, recent data coming from our labs and others suggest a novel paradigm, e.g., that the “inflammaging” environment also contributes to the accumulation of regulatory T cells with age. Indeed, while it is well known that the lack of IL-10 enhances inflammation, our recent data show that the lack of a key inflammatory cytokine, IL-6, actually reduces the accumulation of regula- tory T cells. Thus, we think that during aging there is an interrelated, mixed inflammatory/anti-inflammatory environment that is attempting to maintain homeostasis. Recent findings regarding the homeostasis of regulatory T cells and their role in controlling immune responses with age will be highlighted in this chapter.
# Springer International Publishing AG 2018
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Claire Chougnet and David A. Hildeman contributed equally to this work.
M. Almanan • C. Chougnet (*) • D.A. Hildeman (*) Department of Pediatrics and Division of Immunobiology, MLC 7038, University of Cincinnati College of Medicine and Children’s Hospital Medical Center, Cincinnati, OH, USA Division of Infectious Diseases, Children’s Hospital Medical Center, Cincinnati, OH, USA e-mail: [email protected]; [email protected]; [email protected]
# Springer International Publishing AG 2018 T. Fulop et al. (eds.), Handbook of Immunosenescence, https://doi.org/10.1007/978-3-319-64597-1_82-1
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T-reg Homeostasis and Functions in Ageing
Keywords CD25 • CD8+ T cells • Cytotoxic T-lymphocyte-associated antigen4 (CTLA-4) • Foxp3+ Treg cells • Myeloid-derived suppressor cells (MDSC) • Peripherally derived terg (pTreg) • Thymic-derived treg (tTreg) • Tr1 cells • Transforming growth factor-β (TGF-β) • Treg homeostasis
Contents Introduction ...... 2 Foxp3+ Treg Characterization ...... 3 Foxp3+ Treg Function ...... 5 Foxp3+ Treg Homeostasis with Age ...... 5 Environmental-Mediated Regulation of Treg Homeostasis with Age ...... 6 Inflammation Regulates Tissue-Specific Treg Accrual with Age ...... 8 Treg Function with Age ...... 8 CD8+ Treg Homeostasis and Function with Age ...... 11 Tr1 Homeostasis and Function with Age ...... 12 Functional Cross Talk between Regulators with Age ...... 14 Conclusion ...... 14 References ...... 16
Introduction
The world’s population over age 60 is predicted to increase to 9.7 billion by 2050 (UN). With age, the immune system is subjected to deleterious changes that shape its phenotype and functionality (Lopez-Otin et al. 2013; Goronzy et al. 2007; Arnold et al. 2011; Lefebvre and Haynes 2012; Linton and Dorshkind 2004; Eaton et al. 2004; Miller 1996, 2000; Miller et al. 1997; Weng 2006; Nikolich-Zugich 2014; Garcia and Miller 2002). As a result of this decreased functionality, higher incidence of infections, cancer, and poor vaccination contribute significantly to increased morbidity and mortality reported among the elderly (Lord 2013; Foster et al. 2011; Cevenini et al. 2013; Niccoli and Partridge 2012). A substantial amount of work has already characterized age-driven deleterious changes in adaptive immune cells (B, T) (Linton and Dorshkind 2004). Among these changes are (i) thymic involution and lack of naïve T cell production (Aspinall and Andrew 2000; Hale et al. 2006), (ii) intrinsic defects in T and B cell receptor-signaling (Miller 2000; Scholz et al. 2013; Tamir et al. 2000), (iii) terminal differentiation and acquisition of a replicative senescence phenotype (Effros et al. 2005), and finally, (iv) the expansion of sup- pressive cells in aged hosts (Nishioka et al. 2006; Sharma et al. 2006; Lages et al. 2008; Agius et al. 2009). Over the past 20 years, there has been a rebirth of “suppressor cell” immunology. Initially described in the 1970s, the phenomenon of “suppressive” T cells made a splash and then eventually went by the wayside as the cells were difficult to clone and characterize (Gershon and Kondo 1970; Sakaguchi et al. 2007). Further, a putative MHC cell surface marker that some suppressor cells reportedly expressed, I–J, was later found not to be encoded by an actual gene (Kronenberg et al. 1983).
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It was not until Shimon Sakaguchi described a small population of CD25-expressing T cells must be present in mice in the neonatal period to prevent fatal autoimmunity, that “suppressive” T cells were reborn as “regulatory” T cells (Sakaguchi et al. 1995). Now equipped with modern genetic tools, these cells have been well defined and are characterized by the expression of Foxp3, a master transcription factor that is essential and sufficient for their intrathymic development and acquisition of function (Fontenot et al. 2003; Hori et al. 2003; Wildin et al. 2001; Brunkow et al. 2001; Yagi et al. 2004; Sakaguchi et al. 2010). Extensive work has gone into understanding how they develop, how they are maintained, and the mechanisms they utilize to suppress immune responses (Stephens et al. 2001; Jonuleit et al. 2001; Dieckmann et al. 2001; Baecher-Allan et al. 2001; Miyara and Sakaguchi 2007; Sakaguchi et al. 2009; Schmidt et al. 2012). Moreover, in addition to these Foxp3+ regulatory T cells, many other flavors of “regulatory” T cells have been shown to exist. Notably, CD8+ regulatory T cells (Lu and Cantor 2008), IL-10-producing Foxp3 negative regulatory T cell type 1 (Tr1) (Roncarolo et al. 2006), transforming growth factor-β (TGF-β)- secreting T-helper cell type 3 (Th3) (Wan and Flavell 2007), γδ T cells (Caccamo et al. 2013), and iNKT cells (Ikehara et al. 2000) were shown to temper adaptive and innate immune responses. Thus, the adaptive compartment not only promotes protective immunity but also employs cellular mechanisms to temper those responses. While most of the regulatory mechanisms utilized by these “regulatory” T cell subsets are still controversial, their major role is to maintain tolerance to self- antigens to prevent autoimmune diseases in some situations (Sakaguchi et al. 1995; Sakaguchi 2000) and to guard against exaggerated inflammatory responses to parasites and viruses in other situations (Belkaid 2007). New research also suggests some specialization in their role. There is growing evidence supporting the existence of nonlymphoid tissue specific Foxp3+ Treg cells, with distinct functionality and phenotype that regulate specific tissue resident immune cells (Burzyn et al. 2013; Cipolletta 2014). Importantly, regulatory T cells seem to play differential roles in modulating innate immune responses, for example, IL-10+ non-Treg cells appear more efficient at controlling inflammasome activation than regular Foxp3+ cells (Yao et al. 2015). Importantly, most of the characterization of regulatory subsets with age has focused on Foxp3 + CD4+ Treg and CD8+ Treg in addition to some data that suggest quantitative changes in IL-10 producing CD4+ T cells with age. However, we note that this does not imply that age-driven changes in other regulatory subsets does not occur, it just has not been reported yet to the best of our knowledge.
Foxp3+ Treg Characterization
Initially, most of the characterization of Treg in humans and mice lacked consen- sus on the reliable markers for their identification. For example, cytotoxic T- lymphocyte-associated antigen4 (CTLA-4), glucocorticoid-induced tumor necrosis factor receptor (GITR), Helios, programmed cell death-1 (PD-1), and ICOS (induc- ible costimulator) are all expressed on Treg, but also on activated T cells (Baecher- Allan et al. 2001; Dhuban and Piccirillo 2014). Further, although Foxp3 is a marker
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T-reg Homeostasis and Functions in Ageing for Treg in mice and humans, one caveat is that, in humans, it is induced with T cell activation (Tran et al. 2007; Pillai et al. 2007; Walker et al. 2003). Also, as Foxp3 is an intracellular molecule and therefore intractable to functional analyses after intra- cellular staining, the lack of expression of CD127 (IL-7Rα) and the high expression of CD25 (IL-2Rα) have been used as an isolation strategy for human Treg allowing greater than 95% purity in most cases (Shevach 2004; Liu et al. 2006; Seddiki et al. 2006). However, this strategy leaves out the CD25lo CD127lo Foxp3+ cells that may be enriched in memory Treg (Raynor et al. 2013). In mice, the development of several Foxp3 reporter strains has greatly aided in their isolation and purification for adoptive transfer studies and biochemical analysis (Wan and Flavell 2005; Fontenot et al. 2005a). However, it is important to point out that, one of these reporter mice that generates Foxp3-GFP fusion protein appears to have altered Treg function, so studies using these mice have to be interpreted with caution (Bettini et al. 2012). Nonetheless, the ability to have distinct markers has greatly facilitated our under- standing of the development, homeostasis, and function of these cells. Classification of Treg cells has been an area of active research for years. Based on their development and site of generation, Treg were initially categorized into two main subpopulations of thymic-derived (tTreg) and peripherally derived (pTreg) Treg (Feuerer et al. 2009; Huehn et al. 2009). Both populations express CD4, CD25, and Foxp3. While tTreg constitutively express Foxp3, Foxp3 expression in pTreg is acquired in vivo in CD4 + Foxp3- Tcells in response to a variety of stimuli. However, both tTreg and pTreg have been shown to exert comparable suppressive abilities and in some experimental models they also synergized to obtain optimal regulation (Haribhai et al. 2009). Early insight into the mechanisms involved in the induction of pTreg came from in vitro studies. Both TGF-β1 and IL-2 were required for in vitro induction of Foxp3 expression in naïve CD4 + Foxp3- T cells, leading to a population of in vitro-induced Treg (iTreg) (Davidson et al. 2007; Chen et al. 2003). Following these initial studies, different murine models have confirmed the in vivo generation of pTreg in different tolerogenic settings. For example, pTreg develop following intra- venous administration of antigen in the absence of optimal costimulatory signals (Thorstenson and Khoruts 2001), or oral administration of antigen (Mucida et al. 2005). In the gut, CD103(+) mesenteric lymph node DCs induce pTreg in a TGF-β and retinoic acid (RA)-dependent manner (Coombes et al. 2007). More recently, additional phenotypic and functional analysis identified Treg as heterogeneous cells with distinct chemokine receptor expression that governs their migration to different tissues resulting in regulation of different inflammatory conditions. The transcriptional machinery regulating these diverse Treg phenotypes is believed to involve multiple ancillary transcription factors that act in conjunction with Foxp3 (Liston and Gray 2014; Campbell and Koch 2011). Importantly, work by Campbell et al. provided significant insight into the molecular mechanisms involved in Treg homeostatic localization. The study identified fundamental sub- divisions in Foxp3+ Treg cells, notably, lymphoid homing central Treg (CD44loCD62LhiCCR7+CD25hi) maintained by IL-2 and circulating tissue-homing effector Treg (CD44hiCD62LloCCR7-CD25lo) maintained by continuous signaling through the costimulatory receptor ICOS (Smigiel et al. 2014). Interestingly, similar phenotypic analysis in humans also identified two distinct Treg subpopulations:
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CD45RA+Foxp3lo resting Treg cells (rTreg cells) and CD45RA-Foxp3hi activated Treg cells (aTreg cells) (Miyara et al. 2009). Taken together, work from humans and mice supports the existence of significant heterogeneity within Foxp3+ Treg.
Foxp3+ Treg Function
Treg are known to utilize multiple mechanisms for suppression, some of those mechanisms require contact and others do not require contact with conventional T cells or other cells such as dendritic cells (Sakaguchi et al. 2009). For example, as mentioned above, Treg express surface molecules like cytotoxic T-lymphocyte associated protein-4 (CTLA-4) that can modulate DC maturation or function by inhibiting the interaction of CD80/86 on DCs with CD28 on conventional T cells (Oderup et al. 2006). Treg also express other inhibitory molecules, such as lymphocyte-activation gene 3 (LAG3) (Huang et al. 2004), Programmed cell death protein 1 (PD-1), T cell immunoglobulin mucin 3 (TIM-3) (Gupta et al. 2012), which can all directly interfere with T cell activation. Additionally, Treg can also exert cytotoxic ability via production of granzyme A in humans and granzyme B in mice (Velaga et al. 2015; Cao et al. 2007). Importantly, while many of the factors that Treg utilize to exert their suppressive function have been identified, it is likely that some remain undiscovered, particularly under inflammatory contexts in different tissues. Additionally, Treg can indirectly inhibit effector cells via the induction of regu- latory biochemical pathways via the expression of ectoenzymes CD39 and CD73. High expression of both ectoenzymes on Treg controls the conversion of ADP/ATP to AMP and AMP to adenosine. Notably, extracellular adenosine is known to inhibit T cell proliferation, activation, and cytokine production (Deaglio et al. 2007; Bono et al. 2015). Moreover, Treg can induce the upregulation of IDO in DCs which facilitates the differentiation of naïve CD4(+) T cells toward a Foxp3+ (inducible Treg) (Fallarino et al. 2003) and can disrupt T cell metabolic functions by IL-2 deprivation and induction of apoptosis (Pandiyan et al. 2007). Treg can also produce immunoregulatory cytokines such as TGF-β, IL-10, and IL-35, which can all provide contact independent suppressive effects (Nakamura et al. 2001; Asseman et al. 1999; Collison et al. 2007). Thus, Treg deploy different suppressive weapons from their arsenal, depending upon the context and environment.
Foxp3+ Treg Homeostasis with Age
In aged mice, Treg are increased in both frequency and number in the lymphoid tissues (spleen, lymph nodes), but not in the blood (Sharma et al. 2006; Lages et al. 2008; Raynor et al. 2013; Zhao et al. 2007; Chougnet et al. 2011; Williams-Bey et al. 2011; Chiu et al. 2007; Han et al. 2009). Importantly, age-driven Treg accrual was observed in multiple strains of mice, making it a generalizable phenomenon. Further, the accumulation of Treg is progressive with middle-aged mice displaying interme- diate levels of Treg relative to young or aged mice (Raynor et al. 2013; Chougnet et al. 2011). In humans, an increased frequency of Treg within circulating mononuclear
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T-reg Homeostasis and Functions in Ageing cells was observed in most studies, but these increases were modest (Lages et al. 2008; Rosenkranz et al. 2007; Gottenberg et al. 2005; Gregg et al. 2005). However, one study showed a significant expansion in the proportion of Foxp3+ Treg cells in the skin of old subjects compared with young ones (Agius et al. 2009), suggesting that Treg proportion is also increased with age in humans, but these cells are largely sequestered in the tissues. While Treg accumulate with age, the mechanisms underlying their accrual have just begun to be investigated. Given the massive thymic involution with age, it is unlikely that thymic output contributes significantly to Treg accrual (Chougnet et al. 2011; Aw and Palmer 2011). Another possibility is that aging enhances the conver- sion of conventional T cells into Treg. However, several pieces of data argue against this possibility. First, compared to young mice, conventional T cells from aged mice have less of an ability to undergo peripheral conversion (Chougnet et al. 2011). Second, the vast majority of Treg from aged mice express both Helios and Neuropilin, two markers of nTreg cells (Raynor et al. 2015). Third, using RNAseq analysis, we showed that Treg from aged mice are most closely related to young Treg, rather than to aged or young naïve or memory T cells (Raynor et al. 2015). As Treg accrual did not appear to be due to enhanced production or conversion, the other possibilities were proliferation or survival. We showed that, if anything, Treg from aged mice proliferated less, not more, than their young counterparts (Chougnet et al. 2011; Raynor et al. 2015). Instead, we found that both in vitro and in vivo, aged Treg survived longer than young Treg (Raynor et al. 2013, 2015; Chougnet et al. 2011). Such enhanced survival was due to their progressive decrease in expression of the key proapoptotic molecule Bim (Raynor et al. 2013; Chougnet et al. 2011). Further, mice deficient in Bim (even Treg-specific loss of Bim) accumulated periph- eral Treg rapidly, the peripheral Treg levels in a 4–6-month-old Bim-/- mouse resembled a 1.5-year-old wildtype mouse (Raynor et al. 2013; Chougnet et al. 2011). Thus, aging drives a loss of Bim expression in Treg, which likely contributes to their preferential survival and accumulation.
Environmental-Mediated Regulation of Treg Homeostasis with Age
In young mice, CD25 signaling plays an important role in promoting the develop- ment and peripheral survival of Treg (Bayer et al. 2007; D’Cruz and Klein 2005; Fontenot et al. 2005b; Burchill et al. 2007a, b; Mahmud et al. 2013). However, while Treg are reduced in young CD25-deficient mice, they are not completely absent. One potential reason is that Treg can use other common gamma chain cytokines (e.g., IL-7 and IL-15), when CD25 is missing (Burchill et al. 2007b; Soper et al. 2007; Vang et al. 2008), although, in the normal setting IL-2 seems to be the primary driver of Treg development and peripheral survival. The aging environment is associated with an altered cytokine milieu (O’Mahony et al. 1998; Bruunsgaard et al. 2003; Ershler and Keller 2000) and we recently showed that circulating IL-2 levels are significantly decreased with age (Raynor et al. 2013). Consistent with this, we found
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T-reg Homeostasis and Functions in Ageing that roughly half of the Treg that accumulate with age express low levels of CD25 (Raynor et al. 2013). These data suggested that, in contrast to Treg in young mice, Treg in aged mice are likely less reliant on IL-2, and may rely more on other cytokines. Indeed, we found that Treg from aged mice have higher levels of CD122 (IL-2Rß, the receptor for IL-15) (Raynor et al. 2013). Neutralization of IL- 15, but not IL-2, led to a significant reduction of Treg in aged mice and the additional loss of Bim in IL-15-/- mice rescued the loss of Treg in middle-aged IL-15-/- mice (Raynor et al. 2013). Further, in-depth characterization of old Treg showed significant selective accumulation of Bimlo CD25lo (Raynor et al. 2013). Of note, although aged Treg express increased levels of CD127, IL-7 neutralization studies had no effect on Treg accrual (Chougnet et al. 2011). Thus, it appears that IL-15 favors age-driven Treg accrual perhaps as a compen- satory mechanism for the loss of IL-2 with age. However, since Treg are not completely absent in aged IL-15 deficient mice, it was possible that other cytokines or molecules can also promote Treg accrual with age. Interestingly, similar to earlier characterization of Treg by Campbell et al. (Smigiel et al. 2014), we showed that the vast majority of accumulated Treg in aged mice were of an “effector” phenotype (CD44hi CD62LloICOShiBlimp-1hi) and these were partially maintained by ICOS (Raynor et al. 2015). In contrast, there were less “central” Treg (CD44lo CD62Lhi), which fits the overall model as central Treg cells are largely maintained by IL-2 (Smigiel et al. 2014). Thus, IL-15 and ICOS contribute to Treg accumulation with age. As IL-6 is significantly increased with age (Mysliwska et al. 1998), and IL-6 transgenic mice have elevated numbers of tTreg (Fujimoto et al. 2011), we hypoth- esized that IL-6 increased IL-6 with age might have a modulatory effect on Treg homeostasis with age. Interestingly, with age there was a significant loss of total Treg and more precisely effector Treg in IL-6 deficient mice, likely through the regulation of ICOS expression which in turn likely contributes to the expression of Bim (Raynor et al. 2015). The role of IL-6-ICOS-Bim axis on Treg accrual with age was a novel finding considering the inflammatory role of IL-6. It suggests that the increase in effector Treg population with age acts as a counterbalance to augmented inflammatory IL-6 production. How this IL-6-ICOS-Bim axis signals to control Treg homeostasis with age remains unclear. ICOS signals through the activation of the PI3K/Akt pathway which is known to modulate FOXO3a transcription factor acti- vation, which has been shown to alter Bim expression (Gigoux et al. 2009; Tang et al. 1999; Brunet et al. 1999; Salih and Brunet 2008; Stahl et al. 2002). However, our preliminary data suggest that PI3K/Akt pathway is not solely responsible for regulating Bim expression (unpublished data). Another mechanism that could con- tribute to altered Bim expression includes epigenetic modification to the Bcl2L11 gene (the gene encoding Bim) (San Jose-Eneriz et al. 2009; Paschos et al. 2009). This mechanism is intriguing because it would be a stable and heritable way to ensure lowered Bim expression in a population of cells that periodically divides. Future studies are needed to better elucidate the mechanisms involved in the control of Treg/effector Treg survival with age.
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Inflammation Regulates Tissue-Specific Treg Accrual with Age
In addition to the role played by IL-6, ICOS, and IL-15 in promoting the accumu- lation of Treg with age in the lymphoid tissues, other factors may regulate (both positively and negatively) Treg accumulation in the tissues. Interestingly, a recent study showed that induced inflammation during muscle injury can lead to a Treg deficit with age via an IL-33-ST2 dependent mechanism (Kuswanto et al. 2016). Treg have high expression of ST2 (IL-33R) which controls their specific migration to injured muscle as loss of ST2 on Treg in young mice compromised their accrual in injured muscles but not in the spleen. In aged mice, Treg had a muscle specific gene signature that was unique compared to other lymphoid tissues and hence regulated their migration to/retention in injured muscles. In-depth investigation of the role of IL-33-ST2 axis on accrual of muscle Treg with age showed that reduced IL-33 production upon muscle injury led to diminished accumulation of Treg in injured muscle and resulted in poor repair of skeletal injury of old mice compared to young mice. Importantly, IL-33 supplementation resulted in enhanced Treg accumulation in injured muscle and tissue regeneration suggesting that muscle repair during aging is impaired due to reduced Treg muscle retention. In contrast, prior work showed that Treg accumulate in the skin of aged humans which may be triggered by local inflammatory stimuli or other factors (Agius et al. 2009). Further, the accumulation of Treg correlated with decreased TNF-α secretion by cutaneous macrophages in aged individuals. The implication is that Treg accumulation in human skin with age might contribute to increased risk of cutaneous malignancy and infections. None- theless, the accumulation or lack of accumulation of Treg in nonlymphoid tissues suggests their tissue-specific roles in age-driven immune dysregulation. Combined, these intriguing data support further investigation of mechanisms regulating Treg tissue homeostasis and their role in driving immune-suppression with age.
Treg Function with Age
The increased numbers of Treg with age implies that they have a role in age-driven immune suppression in mice and humans. Recent work has begun to dissect the functionality of aged Treg at the population and individual cell level, and although there are a few exceptions, overall the data suggest increased Treg function with age at both levels. Further, the data suggest a role for Treg in controlling susceptibility to autoimmunity, infections, and tumors in aged mice and humans. For example, recent work showed that Treg from patients with Alzheimer’s as well as Parkinson’s disease had enhanced suppressive activity on a per cell basis, compared to age matched controls (Rosenkranz et al. 2007). In another study, an association was found between an increased frequency of Foxp3-expressing CD4+CD25+CD127lo Treg cells in elderly patients with metastatic primary nonsmall cell lung cancer, suggesting a potential suppressive effect of Treg population on antitumor immunity (Pan et al. 2012). In mice, Treg depletion using α-CD25 antibodies restored the antitumor immune response and led to a significant reduction in tumor burden,
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T-reg Homeostasis and Functions in Ageing suggesting that aged Treg reduced antitumor immunity (Sharma et al. 2006). Sim- ilarly, we showed that same depletion strategy in aged mice reduced the lesion size in a Leishmania major infection model (Lages et al. 2008). Additionally, aged Treg inhibited antigen-specific IFN- γ, but not IL-10 production from effector T cells (Lages et al. 2008). Notably, this latter study was the first to show the similar if not enhanced suppressive activity of old Treg relative to young mice by using the Foxp3-GFP knock-in mice. This is important because, with age, many Treg are CD25lo (Raynor et al. 2013) and therefore, responses measured based on CD25+ Treg cells are likely to be an underestimate of Treg function. In another infection model, a significant expansion of Treg was reported in the spleen of old compared to young mice (Williams-Bey et al. 2011). Although the study implicated Treg expan- sion in diminished CD8 T cell responses to influenza infection with age, unfortu- nately the authors did not test the role of Treg in modulating the anti-influenza responses. Given the findings that supported enhanced functionality of Treg with age, a few studies have investigated the molecular mechanism underlying this enhanced func- tionality. For example, CD40 and CD86 expression on myeloid DCs was enhanced upon depletion of CD25+ T cells in aged mice, suggesting Treg reduce DC function with age (Chiu et al. 2007). Another study showed high suppressive function of Treg from old mice that correlated with an increase in Foxp3 expression driven by the hypomethylation of the upstream Foxp3 enhancer with age (Garg et al. 2014). The authors demonstrated that aged Treg limited the availability of extracellular cysteine which led to imbalanced redox potential that was unfavorable for T cell activation and proliferation. Additionally, Treg modulated the CD86 expression on APCs in an IL-10-dependent manner thus further limiting effector T cell proliferation. Other studies show an equivalent suppression of T cell proliferation in response to Phytohaemagglutinin (PHA) by Treg from young and old individuals (Gregg et al. 2005) as well as a similar inhibitory effect on effector cytokine production (e.g., IFN-γ, TNF-α, IL-6 and IL-2) (Hwang et al. 2009). Similarly, studies in mice have shown that CD4 + CD25+ sorted Treg cells had a similar suppressive effect on the proliferation (Zhao et al. 2007) and IFN- γ production by effector T cells (Sun et al. 2012) compared to their young counterparts. Nonetheless, even if certain models show similar Treg function in aged and young animals, the elevated numbers of Treg would suggest an enhanced overall function. In contrast, in a few other studies, Treg function seemed to decline with age. In some studies, examination of Treg functionality in humans reported a decline in the ability of aged Treg (relative to young) to suppress the proliferation of CD4 + CD25- T cells stimulated by a combination of anti-CD3 and anti-CD28 mAb, independent of the gender of the aged individual (Tsaknaridis et al. 2003). Old Treg cells in mice also had diminished suppression of IL-2 and IFN- γ production in vitro as well as a delayed hypersensitivity reaction (DTH) in vivo (Zhao et al. 2007). Additionally, Treg from old mice were defective in suppressing IL-17 production from T cells compared to young Treg in a model of colitis, probably due to a reduction in STAT3
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T-reg Homeostasis and Functions in Ageing activation (Sun et al. 2012). This suggests that the inability of aged Treg to restrain Th17 cells might contribute to the higher incidence of IL-17-driven, age-related autoimmunity. Collectively, the data strongly suggest increased Treg function with age, however, we do acknowledge, in some instances, decreased Treg function is observed. We think there are a few explanations for these differences. For example, in vitro suppression assays have been used to assess Treg function, there are some major caveats regarding their use and interpretation. First, they may not represent the in vivo mechanisms by which Treg control effector T cell responses. For example, soluble factors like IL-10 or TGF-ß are dispensable for Treg in vitro but not in vivo suppression (Sakaguchi et al. 2009; Asseman et al. 1999; Thornton and Shevach 1998; Suri-Payer and Cantor 2001; Piccirillo et al. 2002). Second, accumulation of CD8 + CD25 + Foxp3+ Treg has also been reported with age, suggesting that the results obtained through the use of α-CD25 antibodies in some of the previous studies might not be CD4 + CD25+ Treg-specific (Sharma et al. 2006). Third, the use of young or aged responder cells might contribute to the observed differential results. Thus, the responsiveness of CD4 + CD25- target cells might differ based on the type and strength of stimuli used, independent of the regulatory properties of the Treg or their number in the assay (Kozlowska et al. 2007). Fourth, aged Treg might differ in function with respect to the type of in vivo models used (e.g., infectious disease, autoimmune, or tumor). Each model adds another layer of complexity because the local inflammatory environment would influence not only cellular effector responses, but also the phenotype, function, and homing of Treg (Wei et al. 2006). For example, the expression of CD62L and CD103 both change with age and can affect migration of Treg cells into secondary lymphoid organs and tissues (Lages et al. 2008; Zhao et al. 2007; Braun et al. 2015; Anz et al. 2011; Suffia et al. 2005). Additionally, CD4 + Foxp3 + CD25lo Treg from aged naïve mice were functional but less suppressive than CD4 + Foxp3 + CD25hi Treg in vitro. However, the two Treg subsets were equally suppressive in aged tumor-bearing mice, suggesting that the functionality of a specific subset of Treg can be altered by the local inflammatory environment (Hurez et al. 2012). More importantly, in humans, functional studies require sampling CD25+ Treg cells from the blood, and as mentioned, CD25 as a surface marker is not sufficient to isolate all Treg, particularly in the aging population as many of these cells are CD25lo and the function of CD4 + CD25+ T cells do not necessarily correlate with CD4 + Foxp3+ Treg cells. Also, Treg from blood are poorly suppressive as they express lower levels of CTLA- 4. Similar tissue specific expression of CTLA-4 was also reported in mice (Lages et al. 2008). Thus, in addition to the potential lack of relevance of the functional assay to their in vivo function, sampling Treg activity in human blood might not be representative of Treg that accumulate in the tissues. Thus, more work is required to determine the heterogeneity within the Treg compartment and how this changes with age so that similar populations can be compared between young and aged mice. Nevertheless, these prior studies have begun to pave the road toward better under- standing of Treg functionality with age.
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CD8+ Treg Homeostasis and Function with Age
CD8+ T cells with immunosuppressive functions were first identified by Gershon and Kondo in the 1970s (Gershon and Kondo 1970, 1971). Regulatory CD8+ T cells are a functionally and phenotypically distinct subset from the classical cytotoxic CD8+ T cells, and play an essential role in fine-tuning immune responses to prevent autoimmunity and inflammation (Konya et al. 2009). A fair amount of work aimed at investigating the development, characterization, and functions of CD8+ Treg using different models. Interestingly, similar to CD4 + Foxp3+ Treg, CD8+ Treg that express Foxp3 have been shown to be produced naturally in the thymus or induced in the periphery through antigen-dependent or antigen-independent mechanisms (Kapp and Bucy 2008; Tang et al. 2005; Churlaud et al. 2015). They are character- ized by the expression of several markers such as CD8aa, CD25, CD38, CD45RA, CD45RO, CD56, Foxp3, CXCR3, LAG-3, CD103, and CD122, as well as by the lack of expression of CD28 and CD127 (Tang et al. 2005; Pomie et al. 2008; Dinesh et al. 2010). Clearly, CD8+ Treg are a heterogeneous population composed of a variety of subsets, each with a distinct phenotype. CD8+ Treg can directly inhibit immune responses via cell contact-dependent or -independent mechanisms. For example, surface expression of CTLA-4 and/or production of soluble regulatory cytokines such as IL-10, TGF-ß can all induce a tolerogenic state in APCs and modulate effector T cell responses (Vukmanovic-Stejic et al. 2001). Notably, much of what is now known about regulatory T cells in aging has been learned from studies investigating CD4 + Foxp3+ Treg, with limited information available on CD8+ Treg cells. With age, qualitative and quantitative changes in CD8+ Treg have been reported in both humans and murine models. While some studies have shown that both the percentage and number of CD8+ Treg were significantly increased in the blood of older individuals and in aged mice (Sharma et al. 2006; Simone et al. 2008), others have shown reduced frequencies of CD8+ CCR7+ Treg induced under low-dose of anti-CD3 and IL-15 (Suzuki et al. 2012) or similar frequencies of circulating CD8 + Foxp3 + CCR7+ Treg in older individuals compared to their young coun- terparts (Wen et al. 2016). However, there remains considerable uncertainty in increased CD8+ Foxp + Treg with age and it needs to be mentioned that with age CD8+ Foxp3+ Treg cells remain of extremely low frequency (<2% of all splenic CD8+ T cells, unpublished data). Likewise, there has also been disagreement as to the functionality of these cells with age; some human studies reported equivalent regulatory function with age, where both old and young CD8+ Treg showed similar abilities in suppressing proliferation and cytokine production from responding cells (Simone et al. 2008). However, others have shown that CD8 + CCR7+ Treg derived from aged individuals had reduced expression of Foxp3+ suggesting reduced sup- pressive abilities (Suzuki et al. 2012) (Nakagawa et al. 2010; Singh et al. 2007). The reduced functionality of CD8+ Treg, however, may not be due wholly to reduced Foxp3 expression; other molecular defects have also been identified in relation to age-related failure of CD8 + Treg functionality. For example, CD8+ Treg from older individuals with giant cell arteritis (GCA) a disease characterized
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T-reg Homeostasis and Functions in Ageing by inflammatory vasculopathy, suffered from inefficient induction of NADPH oxi- dase and shedding of NOX2-containing exosomes. NOX2-containing exosomes are known to regulate the activity of effector CD4+ T cells by abrogating the phosphor- ylation of the upstream signaling molecules ZAP70 and linker of activated T cells (LAT) (Wen et al. 2016). The study suggested that aged patients with autoimmune diseases might have less functional CD8+ Treg, thus highlighting the possible therapeutic applications in manipulating CD8+ Treg with age. Notably, variations in reporting CD8+ Treg percentages or functions with age were clearly related to the lack of consensus on specific markers for CD8+ Treg identification which vary among reports. To date, there is considerable uncertainty about the molecular mechanisms regulating the homeostasis and function of CD8+ Treg with age. Surprisingly, IL-6 positively regulates the Foxp3 + CD8+ Treg development and function, suggesting that the effects of IL-6 in the development of regulatory T cells are not likely to be selective for CD4+ Foxp3+ T cells over CD8+ Foxp3+ T cells. In F759 mice, which carry a Y759F mutation in the gp130 signal transducer of the IL-6 receptor complex, the Foxp3+ CD8+ Treg number was significantly increased with age via enhanced IL-6 signaling (Nakagawa et al. 2010). However, this study only looked at Foxp3+ CD8+ Treg up to 6 months of age. The role of IL-6 and IL-6- dependent molecular mechanisms in modulating the Foxp3+ CD8+ Treg population with age thus requires further investigation. Collectively, whether the aged inflam- matory environment plays a role in Foxp3 + CD8+ Treg accrual and whether conversion, excessive proliferation, or survival mechanisms play a role in main- taining aged CD8+ Treg similar to their CD4 + Foxp3+ counterparts remains largely unknown.
Tr1 Homeostasis and Function with Age
CD4 + Foxp3- type 1 regulatory T (Tr1) cells play a major role in peripheral immune tolerance (Roncarolo and Battaglia 2007). Tr1 cells were first described by Roncarolo et al. in 1988 in the blood of a severe combined immunodeficiency (SCID) patient, who developed long-term tolerance after HLA-mismatched fetal liver hematopoietic stem cell transplant (HSCT) (Roncarolo et al. 1988). Tr1 cells are characterized by their ability to produce high levels of IL-10 and to mediate immune suppression in humans and mice despite their lack of Foxp3 expression (Roncarolo et al. 2006). Several cytokines are involved in the generation of Tr1 cells. For example, IL-10, IL-6, IL-21, IL-27, and type-1 interferons (Vasanthakumar and Kallies 2013; Ziegler- Heitbrock et al. 2003; Stumhofer et al. 2007; Levings et al. 2001; Jin et al. 2013; Pot et al. 2009). Additionally, Tr1 cells can also secrete variable amounts of TGF-ß, IFN-γ, and IL-5 (Battaglia et al. 2006; Groux et al. 1997). Although Tr1 cells lack expression of a specific lineage tracing marker, the cell-surface markers CD49b, LAG-3, and CD226 were recently identified as reliable markers for the characteriza- tion of Tr1 cells in humans and mice (Gagliani et al. 2013). Several different mechanisms have been identified in Tr1 immune-suppression (Gregori et al. 2012). Tr1 cells can modulate T cells and APC function via IL-10 and
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TGF-β secretion (Fiorentino et al. 1991; de Waal Malefyt et al. 1991; Cavani et al. 2000). Additionally, the production of granzyme B (GZB), perforin, and the inter- action of CD226 on Tr1 cells with CD155 on APCs can lead to the death of myeloid APCs (Magnani et al. 2011). More importantly, CTLA-4 signaling represents a major mechanism of Tr1 suppression (Meiler et al. 2008; Akdis et al. 2004). Tr1 cells have been shown to play significant roles in the control of autoimmune diseases and organ transplantation. Thus, there are ongoing clinical trials using human Tr1 cells in various immune-related disorders like hematologic malignancies and Crohn’s disease (Roncarolo and Battaglia 2007; Zeng et al. 2015). Despite the well-established role of Tr1 cells in immune tolerance, little information is known about their homeostasis and function in aging. Increased IL-10 production associated with aging has been reported in both aged mice and humans (Corsini et al. 2006) (Hobbs et al. 1994). However, few studies have determined the cellular source of this increased IL-10. In mice, the capacity of CD4+ T cells to produce IL-10 increased significantly with age, both at the level of gene expression as well as cytokine release upon TCR stimulation. Further analysis showed that CD4 + CD44+ were the predominant source of IL-10 in old mice (Hobbs et al. 1994). Additionally, one study provided evidence for an increase in CD3lo CD5hi (high avidity) aged T cells that expressed higher levels of IL-10 at both the mRNA and protein level (Tatari-Calderone et al. 2012). However, it was unclear whether the increase in IL-10 producing cells reflects a specific increase in CD4+ Foxp3- IL-10+ (Tr1) cells with age. The functional consequences of increased IL-10 in aging is also largely unknown, with suggestions that it could be either protective or harmful, depending on the circumstances. For example, in vitro stimulation of whole blood from aged individuals vaccinated for influenza resulted in a significant increase in IL-10 production which correlated with reduced responses to vaccination compared with their young counterparts (Corsini et al. 2006). In contrast, HIV- specific IL-10-producing CD4 + Foxp3- T cells were significantly more frequent in peripheral blood of older AIDS patients treated by antiretroviral therapy (HAART) than in their younger counterparts. Furthermore, in vitro neutralization of IL-10 correlated with enhanced viral replication and higher IL-1β and IL-6 production. The results suggested that in aged HIV-infected individuals, IL-10-producing CD4 + Foxp3- T cells help better control virus replication by diminishing the release of pro-inflammatory cytokines (Andrade et al. 2012). Also interesting was the fact that an IL-10 polymorphism associated with IL-10 high production was frequent in male centenarians and was suggested to be associated with longevity (Lio et al. 2002). Interestingly, a recent study provided evidence for a role of the inflammatory aged environment in the regulation of IL-10-producing CD4+ T cells. This study suggested that the tolerogenic environment permissive of tumors seen in aged mice is not due to T cell cell-intrinsic differences; rather, it is due to the aged environ- ment’s conditioning of these cells, as tumor specific CD4+ T cells from young mice transferred into old mice had defective Th1 differentiation, acquired a Tr1 like phenotype and were unable to inhibit tumor growth (Tsukamoto et al. 2015). The data raises the question about the effect of IL-6 or other aspects of the aged
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T-reg Homeostasis and Functions in Ageing environment on the induction of Tr1 cells with age. Thus, future studies are required to investigate the homeostasis and function of Tr1 cells with age.
Functional Cross Talk between Regulators with Age
A realistic picture of both Treg homeostasis and functionality with age is based on the presence of crosstalk between different regulatory subsets as well as effector T cells. For example, one study showed that Treg as well as myeloid-derived suppres- sor cells (MDSC) are both increased in with age (Hurez et al. 2012). Depletion of Foxp3 + Treg using diphtheria toxin (DT) further increased the numbers of MDSC in aged melanoma tumor-bearing mice and abrogated the antitumor effects of Treg depletion observed in young mice. These results in contrast with enhanced tumor control in aged mice upon Treg depletion in a BM-185-EGFP tumor model com- pared to young mice (Sharma et al. 2006). Thus, with age Treg might have a differential regulatory effect on effector T cells and other regulatory subsets. Like- wise, we observed a profound increase in CD4 + Foxp3-IL-10-producing regulatory T (Tr1) cells upon Treg depletion in aged mice (manuscript in preparation), and Treg depletion in aged mice latently infected with MCMV resulted in viral reactivation, likely through enhanced accumulation of these CD4 + Foxp3-IL-10-producing regulatory T cells and (Almanan et al. 2017). Our data is consistent with previously work that suggested enhanced capacity of aged CD4 + CD25+ Treg in regulating IL-10 production from responding cells compared to their young counterparts (Hwang et al. 2009). This suggests that, with age, Foxp3+ Treg might play not only their classical role in regulating effector cells but also in regulating other regulatory populations to ensure protective immunity and limit immunopathology. Thus, the ways in which this interconnected network of regulatory and effector T cells evolves with age could present both advantages and disadvantages, depending on its context.
Conclusion
Several important questions remain unanswered: what are the molecular mecha- nisms of aged Treg-mediated suppression? Can specific Treg cells be targeted therapeutically in order to boost antitumor immunity or to control chronic infections to improve the quality of life of the elderly population? The role of IL-6 in Treg homeostasis with age is a novel finding. Although IL-6 plays a central role in the inflammaging process leading to frailty, morbidity, and mortality with age (Maggio et al. 2006), IL-6 also counterbalances excessive inflammation by promoting several Treg subsets accrual with age. These data collectively suggest a reevaluation of the “inflammaging” hypothesis to include its natural anti-inflammatory counterbalance. In all, the aging environment reflects a disturbed equilibrium that is attempting to maintain homeostasis (Fig. 1). It is in this mixed inflammatory/anti-inflammatory environment that immune responses are
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Fig. 1 Age-related mixed pro- and anti-inflammatory environment. Regulatory T cells that are Foxp3+, CD8+ Foxp3+, and Foxp3- IL-10+ (Tr1) accumulate dramatically in aged hosts along with a profound increase in the levels of IL-6 and IL-10. With age IL-6 plays not only its classical inflammatory role but also promotes regulatory T cells accrual and increased production of IL-10, perhaps as a mechanism to counterbalance the dysregulated inflammatory environment and main- tain homeostasis. While Foxp3+ Treg cells have been shown to produce higher levels of IL-10 with age, enhanced production of IL-10 from Tr1 cells with age still unclear. The homeostasis of the aged mixed inflammatory/anti-inflammatory environment is also related to the cross talk between different regulatory and effector T cells. While the accumulation Foxp3+ T cells and perhaps Tr1 cells with age clearly would have a negative impact on naïve/effector T cells or on APCs to inhibit T cell responses, Foxp3+ Treg accrual might also restrain undesirable consequences of Tr1 accumu- lation and accompanied increase in IL-10 production to maintain protective immunity. However, the cross talk between the regulators with age remains unclear instigated and propagated. Thus, a better understanding of mechanisms underlying tissue-specific Treg homeostasis and function will hopefully lead to strategies tailored to manipulate local Treg accumulation and function to selectively enhance (e.g., antitumor immunity, infections) or inhibit (e.g., autoimmunity) effector T cell responses. However, our data also suggest that those manipulations may have unforeseen consequences, due to the intricate network of regulatory cells that act synergistically or antagonistically, depending on the tissue context.
Acknowledgments We thank Kate Carroll for critical reading of the manuscript and the funding from NIH -RO1AG033057 to C.A.C. and D.A.H
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Chapter 3
TISSUE-SPECIFIC CONTROL OF LATENT CMV REACTIVATION BY REGULATORY T CELLS
Maha Almanan1, Jana Raynor1, Allyson Sholl1, Mei Wang2, Claire Chougnet1,
Rhonda D. Cardin2, 3¶*, and David A. Hildeman1¶*
Running title: Treg and latent MCMV infection
1Department of Pediatrics, University of Cincinnati College of Medicine, Division of
Immunobiology, Children’s Hospital Medical Center, Cincinnati, OH, United States of America.
2Department of Pediatrics, University of Cincinnati College of Medicine, Division of Infectious
Diseases, Children’s Hospital Medical Center, Cincinnati, OH, United States of America.
3Department of Pathobiological Sciences, School of Veterinary Medicine, Louisiana State
University, Baton Rouge, LA, United States of America.
*Corresponding authorsEmail:[email protected](DH),[email protected] (RC)
¶These authors contributed equally to this work
Keywords: Foxp3+ Treg, IL-10, MCMV, virus, tissues
Published in PLOS Pathogens
August 10, 2017
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Abstract:
Cytomegalovirus (CMV) causes a persistent, lifelong infection. CMV persists in a latent state
and undergoes intermittent subclinical viral reactivation that is quelled by ongoing T cell
responses. While T cells are critical to maintain control of infection, the immunological factors
that promote CMV persistence remain unclear. Here, we investigated the role of regulatory T
cells (Treg) in a mouse model of latent CMV infection using Foxp3-diptheria toxin receptor
(Foxp3-DTR) mice. Eight months after infection, MCMV had established latency in the spleen,
salivary gland, lung, and pancreas, which was accompanied by an increased frequency of Treg.
Administration of diphtheria toxin (DT) after establishment of latency efficiently depleted Treg
and drove a significant increase in the numbers of functional MCMV-specific CD4+ and CD8+
T cells. Strikingly, Treg depletion decreased the number of animals with reactivatable latent
MCMV in the spleen. Unexpectedly, in the same animals, ablation of Treg drove a significant increase in viral reactivation in the salivary gland that was accompanied with augmented local
IL-10 production by Foxp3-CD4+T cells. Further, neutralization of IL-10 after Treg depletion significantly decreased viral load in the salivary gland. Combined, these data show that Treg have divergent control of MCMV infection depending upon the tissue. In the spleen, Treg antagonize CD8+ effector function and promote viral persistence while in the salivary gland
Treg prevent IL-10 production and limit viral reactivation and replication. These data provide new insights into the organ-specific roles of Treg in controlling the reactivation of latent MCMV infection.
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Author Summary:
Cytomegalovirus (CMV) infection in both mice and humans is normally initially contained by a vigorous adaptive immune response that drives the virus into latency in multiple tissues.
However, the immunologic mechanisms that control latency are not well understood. In this report, we have examined the role of regulatory T cells (Treg) in a mouse model of CMV infection. Interestingly, depletion of regulatory T cells had profound consequences on MCMV latent infection, depending upon the tissue. In the spleen, Treg depletion enhanced CD8+ T cell responses and reduced reactivatable latent infection from the spleen. In striking contrast, in the salivary gland, Treg depletion enhanced the production of IL-10 from CD4+ T cells as well as viral reactivation. Thus, Treg play divergent and tissue specific roles in controlling MCMV reactivation from latency.
106
Introduction:
The immune system has evolved multiple innate and adaptive strategies to control pathogens[1]. Likewise, in order to ensure their persistence, pathogens have developed sophisticated and elaborate mechanisms to avoid the host immune system establishing latent infections that are never cleared from the host [2-7]. Human cytomegalovirus (HCMV) and its murine homolog (MCMV) are well-studied examples of pathogens that have developed multiple means to establish latency [8-10]. MCMV is a reasonable model for HCMV as it shares multiple biological characteristics and significant homology to the genome of HCMV[11]. A large number of HCMV and MCMV genes are involved in modulating innate and adaptive host immune responses [12-15]. During primary infection, these viruses vigorously replicate and disseminate by infecting many cell types, including epithelial, endothelial, smooth muscle, and connective tissue cells, as well as specialized parenchymal cells in multiple tissues [16].
Primary CMV infection is well controlled by a robust early NK cell response followed by
CD4+ and CD8+ T cell responses that ultimately results in control of virus replication, although the virus is not eliminated and persists for the lifetime of the host [17-20]. Interestingly, prior work shows that the control of lytic virus in different tissues requires distinct immune cell populations [21-25]. In the spleen, epitope-specific CD8+ T cells are sufficient to control acute
MCMV infection [26]. Whereas in the salivary gland (SG), CD4+ T cells and, in particular their production of IFN-γ, are crucial for terminating lytic viral replication. Further, IL-10R blockade or the absence of CD4+ cell-derived IL-10 enhanced the accumulation of IFN-γ–producing
CD4+ T cells and inhibited MCMV persistence in the SG [21, 22, 25, 27-29]. This role of CD4+
T cells in the SG is critical, as there appears to be a reservoir of MCMV in non-hematopoietic cells within the SG that downregulate MHC class I and become resistant to CD8+ T cell killing
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[22]. However, recent work showed that CD8+ T cells can play a role in controlling MCMV in the SG after local re-infection [30]. The likely explanation for this differential role of CD8+ T cells during re-infection is because of the higher expression of class I MHC in acutely infected cells in contrast to cells harboring latent virus. Regulatory T cells (Treg) have also been shown to contribute to immune-mediated control of acute MCMV infection. In vitro, Treg were shown to suppress the function of MCMV-specific CD8+T cells via secretion of TGF-ß [31]. In vivo, during acute MCMV, Treg depletion resulted in enhanced MCMV-specific T cell responses and decreased viral load [32]. Nonetheless, following the termination of lytic infection, the virus persists in a latent state in which viral genomes are present in the absence of replicating virus.
MCMV establishes latency in a number of tissues similar to HCMV, including the spleen, lungs, and bone marrow [33-36]. This latent form of HCMV drives a substantial amount of morbidity when it reactivates in immune suppressed individuals (e.g. aged individuals, persons with HIV infection, transplant patients) [37-39]. During latent infection, HCMV persists in CD34+ hematopoietic stem cells as well as more committed myeloid lineage progenitor cells, monocytes, and macrophages [40-44]. Both myeloid lineage cells [33, 45], as well as non- hematopoietic cells such as endothelial cells in many organs are considered cellular sites of latent infection for MCMV [33, 46, 47].While a fair amount is known regarding immune mechanisms controlling acute MCMV infection, significantly less is known about immune mechanisms contributing to the control of latent MCMV infection. It has been reported that once latency is established, the cooperative function of lymphocytes including NK, CD4+, CD8+ T cells and
CMV-specific antibodies prevent the production of lytic virus from latent pools in the spleen and lungs and SG [48]. In most visceral organs, CD8+ T cells play a crucial role in preventing the emergence of lytic viral replication. For example, CD8 T cells maintain latency by epitope-
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specific sensing of transcriptional reactivation in the lungs and killing these cells [49]. Similar to
their role in controlling acute infection in the SG, CD4+ T cell production of IFN-γ is critical to
prevent lytic virus production from latently infected cells [22, 50, 51].
During latency, there is substantial epigenetic suppression of viral immediate-early genes
that must be overcome by cellular signals to exit from latency [42, 44, 52]. Two models have been proposed for MCMV reactivation during latency. First, a two-step model proposed by
Hummel et al. [53] in which an inflammatory immune response can drive the activation of the major immediate-early (MIE) gene, initiating reactivation from latency. The second step requires an immune-suppressive environment, such as that which occurs during γ-irradiation or immune- deficiency. This immune-suppressive environment allows the virus to actively replicate, facilitating the production of lytic virus. In one example of this two-step scenario, the production of inflammatory mediators like tumor necrosis factor alpha (TNF-α), interleukin-2
(IL-2), and gamma interferon (IFN-γ) following allogeneic transplantation of kidneys in mice initiated IE gene expression [54]. However, full lytic reactivation of MCMV was only found in immune-deficient recipients [55]. Similarly, in human transplant patients, HCMV reactivation correlated with increased inflammatory responses [56, 57]. A second model focusing on latency and reactivation in the lung suggests that virus is reactivating frequently in immunocompetent hosts as shown by detectable levels of the immediate-early ie1/ie3 transcripts but that checkpoints exist which prevent full production of infectious virus following transcriptional activation [58-60]. Increased TNF-α levels were not enough to fully reactivate virus, although five-fold higher levels of IE transcripts were detected. Presumably memory T cells, specific for the IE proteins, lyse the reactivating cells before virus is produced [9, 58-61]. Multiple immune cells such as CD4+ and CD8+ T cells and other factors such as antibody and IFN-γ have also
109 been shown to be important for maintaining latency [48, 62]. However, the immune cells and molecular factors that control viral latency and whether these cells and factors are the same between different tissues remains unclear.
In recent work, recurrent virus reactivation was accompanied by a concomitant increase in Treg frequency and suppressive functionality [63]. Thus, while Treg have been associated with viral reactivation, a causative role in control of latent CMV/MCMV has not been established. Here, we investigated the role of Treg in controlling latent MCMV infection.
Strikingly, we found that Treg had opposing effects in distinct tissues harboring latent virus. In the spleen, Treg promoted viral persistence and suppressed local MCMV-specific effector T cell responses, while in the SG Treg were required to prevent viral reactivation/replication and IL-10 production by Foxp3- CD4+ T cells. These data show that Treg play divergent and tissue- specific roles in controlling viral reactivation from latency depending upon the type of T cells
(effector or regulatory) they inhibit.
Results:
Activated Treg are increased in the spleen during latent MCMV.
Foxp3+ regulatory T cells (Treg) can suppress effector T cells and promote MCMV replication in the spleen and salivary gland (SG) during acute MCMV infection [32]. However, there are no studies examining the role of Treg during latent MCMV infection. To investigate the role of regulatory T cells (Treg) during latent MCMV infection, we infected cohorts of C57BL/6 and
Foxp3-DTR mice with MCMV and waited for 8 months. Notably, in the C57BL/6 background, low level virus replication continues in the SG and spleen for months longer than is observed in mice on a BALB/c background. Thus, it took 7 to 8 months following infection for the
110 establishment of latency, as measured by the inability to detect replicating virus in multiple tissues, including spleen, lung, liver, pancreas, and SG (Supplementary Table.1). Next, we examined the levels and activation status of Treg in the spleen of latently infected mice.
Interestingly, compared to uninfected mice of the same age, the frequency of Treg (Fig.1A, B), and in particular “effector” Treg (as assessed by CD69 expression) were significantly increased in MCMV infected mice (Fig.1C, D), although the total number of these cells was not different
(Supplementary Fig.1). Thus, latent MCMV infection favors the preferential accrual of activated
Treg in the spleen.
Treg inhibit effector T cell responses and promote latent MCMV infection in the spleen.
Next, we determined the role of Treg in latent MCMV infection using Foxp3-DTR mice, which allows the specific depletion of Treg using diphtheria toxin (DT) [64]. Before DT administration the levels of MCMV-specific T cells were not significantly different between WT and Foxp3-
DTR mice (Supplementary Fig.2A). After DT administration, the frequency and total numbers of Treg were substantially reduced relative to DT-treated controls (Supplementary Fig.2B). As
MCMV-specific T cells control viral replication, we determined the role of Treg in suppressing
MCMV-specific T cell responses during latent infection. Treg depletion resulted in a significant increase in MCMV-specific CD8+ T and CD4+ T cells, relative to control animals (Fig.2A).
MCMV-specific CD8+ T cells in Treg depleted mice expressed markers of differentiation
(KLRG-1) and proliferation (Ki67) (Supplementary Fig.2C, D). Functionally, the frequency of
TNF-α or IFN-γ single producers and the numbers of single and double producers was significantly increased in Treg depleted mice (Fig.2B). Similarly, CD107α expressing MCMV-
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specific CD8+ T cells were significantly increased after Treg depletion (Fig.2C). Thus, Treg
suppress effector T cell responses during latent MCMV infection.
Given the differences in effector T cell responses following Treg depletion, it was
important to evaluate if loss of Treg would modulate the latent MCMV viral load. Seven days
after initial DT treatment there was no actively replicating virus in the spleen in either controls or
Foxp3-DTR mice as assessed by plaque assay (Supplementary Table.2). Using a spleen explant
assay [65], we examined the impact of Treg on the latent viral pool. The spleen explant assay
provides a functional assay to assess reactivation from latency, and thus, indirectly provide
information on viral load levels since a lower latent viral load is expected to be less efficient at
reactivation [65]. As detected in the explant assay, for the WT control mice, 6 out of 9 mice had
evidence of viral reactivation by 28 days post explant culture, whereas MCMV reactivated only
in 3 out of 9 Foxp3-DTR mice (Fig.3A). Further, titers of reactivating virus in WT mice were
significantly higher in the first two weeks than those detected in the few Treg-depleted mice that
had reactivated MCMV (Fig.3B). To further evaluate whether reduced virus reactivation from
the spleen also correlated with reduced levels of viral DNA following Treg depletion, we
performed qPCR on total splenic DNA to quantify viral genomes. Although there was variation
from one experiment to the next, we saw a consistent decrease in viral genome levels in spleens
from Foxp3-DTR mice relative to C57BL/6 controls across four independent experiments
(Supplementary Fig.3, average percent decrease of 62.72% +/- 9.4; p<0.007). Thus, Treg are critical to maintain the reactivatable latent pool of MCMV in the spleen. However, the increase in overall spleen cellularity (and hence DNA content) following Treg depletion could also contribute to a decreased detection of viral load.
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Treg are required to prevent MCMV reactivation and MCMV-specific CD4+ T cells in the
SG.
Given the role of Treg in promoting viral persistence in the spleen, we next determined the role
of Treg in control of latent MCMV infection in the SG. Similar to the spleen, latency in the SG
was established by 8 months as evidenced by the lack of replicating virus prior to DT
administration (Fig.4A). However, in stark contrast to the reduction in viral load in the spleen
upon Treg depletion at day 7, Treg depleted mice had detectable replicating virus in the SG.
While 3 out of 11 WT mice had a barely detectable viral reactivation, 9 out of 10 DT-treated
Foxp3-DTR mice had a significant increase in viral reactivation and viral titers in the SG
(Fig.4A, B). Indeed, in several experiments, virus was not detected in the WT SG whereas actively replicating virus was consistently detected in the SG of the Foxp3-DTR mice (Fig.4A,
B). Additionally, reactivation occurred as early as day 4 after Treg depletion with evidence of increased, albeit low, viral titers in reactivating mice observed at that time (Supplementary
Fig.4A, B). Thus, in stark contrast to the spleen, Treg are critical to prevent lytic viral reactivation in the SG.
The SG provides a site for MCMV viral persistence and is a major site for accrual of tissue resident MCMV-specific memory CD8+ and CD4+ T cells [30, 66], although MCMV- specific CD8+ T cells are unable to drive viral clearance in this organ [22]. Given that Treg inhibited viral reactivation in the SG, we next investigated the SG T cell responses following
Treg depletion in latently MCMV-infected mice. First, we established that DT administration efficiently depleted Treg in the SG (Supplementary Fig.5A). In contrast to the spleen, we did not observe any change in the MCMV-specific CD8+ T cells (Fig.4C) or tissue resident MCMV-
specific CD8+ T cells (CD103+CD69+) (Supplementary Fig.5B) in Treg depleted mice.
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However, depletion of Treg resulted in an increase in the number of MCMV-specific CD4+ T cells in SG (Fig.4D). Thus, Treg suppress CD4+ but not CD8+ T cells in the SG during latent infection.
Treg suppress CD4+ Foxp3- IL-10+ cells in the SG and IL-10 is critical for limiting viral load in the SG after Treg depletion.
Prior work showed that IL-10 is critical for promoting viral replication in the SG [27, 28].
Indeed, we found that depletion of Treg drove a significant increase in SG IL-10 mRNA levels
(Fig.5A). Given the increase in CD4+ T cell responses in the SG, we next examined the potential cellular sources of IL-10. Importantly, CD4+Foxp3-cells were the predominant source for IL-10 production compared to CD8+ T cells and non–T cells (Fig.5B). Combined, these data show that
Treg suppress a population of CD4+ Foxp3- IL-10+ cells, consistent with a potential role of these cells in MCMV replication in the SG. As the effects of Treg depletion on viral reactivation/latency were quite different in the SG compared to the spleen, we examined IL-10 levels in the spleen as reduced IL-10 could potentially explain the different viral loads. Similar to the SG, IL-10-producing FoxPp3- CD4+ T cells were increased in the spleen (Supplementary
Fig. 6A). However, unlike the SG, total IL-10 mRNA was not increased in the spleen, suggesting that the total amount of IL-10 in the spleen is not substantially increased
(Supplementary Fig. 6B).
To further investigate the role of IL-10 in the SG after Treg depletion, we blocked IL-
10R signaling by administration of a blocking IL-10R antibody with and without Treg depletion using Foxp3-DTR mice (Fig.5C). As expected, Treg depletion was again accompanied by an increase in the number of mice harboring reactivating virus as measured by plaque assay
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(Fig.5D). Strikingly, neutralization of IL-10 during Treg depletion significantly reduced the total number of mice with MCMV reactivation (9/13 to 3/13) (Fig.5D). Thus, our data show that Treg limit IL-10 production in the SG and that IL-10 is essential for viral reactivation/replication in the SG after Treg depletion.
Discussion:
Investigating the role of regulatory T cells in latent MCMV has been hampered by the lack of
appropriate tools. Our ability to assess the role of Treg in latent MCMV infection was greatly
facilitated by two essential tools. First, Foxp3-DTR mice, which express diphtheria toxin
receptor under the Foxp3 promoter allowed for depletion of Treg upon DT administration[64].
Second, the use of a sensitive tissue explant assay and qPCR allowed us to assess the reactivatable latent viral pool in mice with Treg depletion [65, 67]. Herein, we found that
depletion of Treg during latent MCMV infection had profound consequences. In the spleen, Treg
restrained MCMV-specific CD4+ and CD8+ T cells and promoted the latent viral pool. In stark
contrast, in the SG, Treg were essential to limit viral reactivation because they prevented the
emergence of IL-10- secreting Foxp3- CD4+ T cells. Thus, we demonstrate unique tissue-
specific functions of Treg in control of latent MCMV infection.
During latent MCMV infection, we found a significant increase in activated Treg in the
spleen. We speculate that the increase in Treg maybe a direct consequence of the chronic
stimulation due to the periodic low-level viral reactivation during latency. In agreement, in other
chronic infections like Leishmaniasis, hepatitis C virus (HCV), hepatitis B virus (HBV) and
human immunodeficiency virus (HIV), Treg frequency was substantially increased in the spleen
and other tissues [68-70]. Interestingly, human herpes virus 6 (HHV-6) infection, another herpes
115 virus closely related to HCMV, induces virus-specific CD4+ and CD8+ regulatory T cells [71].
Thus, chronic infections appear to promote their own persistence by driving Treg accrual.
Strikingly, Treg depletion resulted in markedly reduced latent virus in the spleen as measured by two assays. Some mice had undetectable viral DNA levels in the spleen, suggesting that viral load had been reduced to levels that were below the limit of detection in our assay. We acknowledge that the increase spleen size and cellularity could contribute to the observed reduction in latent viral load. However, in Treg-depleted mice where viral DNA load was undetectable, the spleen size and cellularity was similar to mice with detectable levels of viral DNA. Nonetheless, future experiments will investigate this in more detail, examining the number of cells harboring latent virus and the levels of virus within such cells. The spleen explant assay allowed us to assess one aspect of latent MCMV infection, the ability to reactivate from latency as a measure of whether reactivatable latent viral loads in the spleen were affected by Treg depletion. In our hands, if virus is replicating in the spleen at the time of isolation, this is detected by seven days post explants [65, 72]. However, if virus is latent in the tissue at the time of isolation, it takes longer for the virus to be detected in the explant assay, usually by day
14, and then rapidly spreads and replicates within the culture. Although this assay is not quantitative per se, we recently demonstrated that a mutant virus was unable to reactivate from the spleen by explant culture and had reduced levels of viral DNA [65], (Cardin, manuscript in preparation). Here, our results show that the Foxp3-DTR mice were less efficient at reactivation in this assay, suggesting lower levels of latent virus in the spleen. Importantly, we found that both the number of Foxp3-DTR mice with reactivating virus and also the levels of virus replication per mouse in the spleen explant cultures were reduced. Decreased reactivation from the spleen was highly reproducible between studies, and indeed, qPCR analysis of viral DNA in
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the spleen could indicate lower levels of latent virus in mice depleted of Treg, with the caveat of
spleen size as mentioned earlier. Alternatively, we cannot rule out that the increased numbers of
virus-specific T cells in the Treg-depleted spleens are functional after placement into the explant
culture and thus elimination of latently-infected cells could also occur in vitro; however, these T cells likely have a very limited survival in vitro. It is important to note that we were unable to carry out long-term Treg depletion (greater than 14 days), because of the rampant and lethal autoimmunity that ensues with long-term Treg depletion in these mice [64]. Further, the impact of Treg depletion on latent viral genomes likely only impacted those genomes that underwent a reactivation event and became detectable by the immune system due to viral antigen expression.
Given that Treg depletion was accompanied by a substantial increase in cytotoxic MCMV- specific CD8+ T cells in the spleen, it is likely that Treg depletion led to the reactivation of some viral genomes, which was likely quickly quelled in the spleen by an MCMV-specific CD8+ T cell response, possibly even before lytic virus was produced [49]. Additional studies are needed to address this further.
One important question raised by our study is the mechanism(s) by which Treg suppress
CD8+ T cell responses to MCMV. Prior in vitro work suggested that Treg utilize TGF-β to suppress MCMV- specific T cell responses [31], although CTLA-4 and IL-10 may also have a regulatory role [73, 74]. In a chronic Friend virus (FV) infection model, FV-specific CD8+ T cells were also partially restrained by Treg, although the mechanism(s) appear to be independent of PD-1 and Tim-3 [75]. Interestingly, our previously published data show that Treg from aged mice have significantly increased levels of IL-35p19, suggesting that IL-35 may contribute to suppression of MCMV-specific CD8+ T cell responses [76]. However, a recent paper showed that IL-35 produced by NK cells during acute MCMV infection promotes, rather than inhibits
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IL-10 production [77]. Nonetheless, further studies are required to determine the mechanism by
which Treg restrain MCMV-specific CD8+ T cell effector functions and thus promote latent
infections.
It was very surprising that, while the depletion of Treg resulted in reduction in viral load
in the spleen, it was accompanied by an augmented reactivation and/or replication of the latent
viral pool in the SG. We envision three major potential mechanisms that may explain these
results. First, it is possible that under chronic infection such as that observed in MCMV infected
SG, Treg could lose immunosuppressive abilities and acquire the phenotype of a more
pathogenic or anti-viral Treg. There is precedence for Treg metamorphosis in tumor models and
under chronic inflammatory conditions [78-81]. This makes teleological sense as chronic
inflammation may favor Treg with more effector function to replace exhausted effector T cells.
However, we looked for Treg expression of pathogenic markers (i.e. T-bet, RORγt) on SG Treg
and failed to find expression of these markers. Second, it is possible that the kinetics of latency
establishment between the two tissues explains the differential effects on latent virus (clearance
& reactivation) in the two tissues. For example, because the virus established latency in the
spleen after three months but was latent in the SG after 7 months, perhaps clearance was easier to
achieve in the spleen due to a lower level of latent viral load. However, we performed several
experiments examining Treg depletion 2-5 months after primary infection and found that, similar to 8 months after infection, that loss of Treg drove significant decrease in MCMV replication in the spleen (Supplementary Fig.7). Thus, it is likely that the tissue microenvironment rather than the timing controls the tissue-specific role of Treg. Third, it is possible that Treg maintain their
suppressive capacity and instead of their “classical” role of inhibiting pro-inflammatory T cells,
it is possible that they similarly control anti-inflammatory T cells. If so, the result of suppressing
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anti-inflammatory cells would be the promotion of an immune-suppressive environment which is permissive for viral reactivation/replication. Our data are most consistent with this latter possibility. In this regard, CD4+ T cell production of IL-10 was increased in both the spleen and
SG after Treg depletion. One obvious question is why didn’t this increase in IL-10 drive viral reactivation in the spleen? One likely explanation is that, in the spleen, where CD8+ T cells are critical to prevent lytic virus production, their levels of IL-10R are significantly decreased, making them insensitive to IL-10 [82].
Our data also show that Treg depletion drives viral reactivation in the SG and implicates
IL-10 in the process. However, current data in the literature suggest that IL-10 contributes more to viral replication than outright reactivation [83]. Indeed, prior work has shown that the specific production of IL-10 from Foxp3- CD4+ cells attenuates acute antiviral immune-responses and leads to persistent viral replication in the SG [27]. Thus, while our data clearly show that inhibition of IL-10R signaling restored viral control, more work is required to conclusively determine whether IL-10 promotes outright reactivation or promotes viral replication. However, in our study, we initiated IL-10R antibody treatment following the initiation of Treg depletion, thus, if virus had started to reactivate, it could have been effectively controlled. The effect of IL-
10 on viral control could be direct or indirect. IL-10 can directly affect CD4+ T cell production of effector cytokines like IFN-γ [84]. Indeed, IFN-γ is indispensable for the control of viral load in the SG [25, 85]. However, there was no diminution in IFN-γ production upon Treg depletion, instead IFN-γ production was actually increased (Supplementary Fig.8). Alternatively, IL-10 can indirectly compromise CD4+ T cell responses in the SG by interfering with the responsiveness of APC to IFN-γ. For example, it is well known that IL-10 inhibits the effect of
IFN-γ by interfering with IFN-γ induced genes, preventing the phosphorylation of STAT-1
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molecules and activation of monocytes [86]. Thus, while our data clearly show a role for IL-10,
more work is required to determine the cellular targets of IL-10 that regulate
reactivation/replication. Notably, the cellular site(s) of MCMV latency in host tissues is a long
debated issue. Like HCMV, MCMV can establish a latent infection in cells of the myeloid
lineage and these cells are able to respond to IL-10. However, other, non-hematopoietic targets,
like endothelial cells cannot be excluded.
This divergent and tissue-specific role in controlling MCMV viral latency by Treg raises
the question as to whether Treg manipulation is a useful therapy in latent HCMV infection. In
HCMV, manipulating Treg is viewed as a promising therapeutic approach to control latent viral
reactivation in immune-compromised hosts like organ transplant patients. In a prior study, the
use of immune-suppressive drugs like daclizumab (anti-CD25), steroids and calcineurin
inhibitors in CMV sero-positive renal transplant patients, led to reduction of Treg levels that
correlated with enhanced levels of CMV-specific effector T cells, suggesting that modulation of
Treg favors maintenance of CMV-specific immunity [87]. However, Treg depletion may also
promote viral reactivation in sites such as SG that are not assessed in treated patients. Thus,
manipulating Treg could be a double-edged sword. Treg in CMV infection might exert different
functional activity depending on their localization within the infected host and the T-cell
responses that they regulate. Our data, suggesting that Treg could regulate another suppressor
CD4+Foxp3-IL-10+ cells in the SG and limit viral reactivation opposed to their conventional role in regulating effector T cells as observed in the spleen, is a novel concept. It is unclear whether the mechanisms employed by Treg to inhibit effector T cell responses vs. other regulatory cell populations are distinct. Understanding such mechanisms might be crucial however to enhance the suppressive effects of Treg on other immune suppressive cell
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populations in some instances (chronic infections) or block the suppressive effect of Treg on
others (i.e. autoimmunity).
Methods:
Cells and Virus
NIH 3T3 cells (ATCC CRL1658) were grown in Dulbecco’s modified Eagle’s medium (DMEM,
Media tech, Herndon, VA) supplemented with 10 % fetal bovine serum (FBS, Hyclone, Logan,
UT), 7.5% Sodium Bicarbonate, 4 mM HEPES, 2 mM L-glutamine, and gentamicin in a
humidified 5% CO2 incubator at37ºC. Parent stocks of the wild type MCMV K181 (originally a kind gift from Dr. Ed Mocarski, Stanford University) were prepared in NIH 3T3 cells from a SG-
derived virus stock as previously described [65]. Virus titers were determined by plaque assay on
NIH 3T3 cells. All virus stocks were stored at -70oC and re-titered before use in experiments.
Mice and infection
Young C57BL/6 mice were purchased from Taconic Farms (Germantown, NY)
Foxp3-IRES-DTR-GFP knock-in C57BL/6 mice were a generous gift from Dr. A. Rudensky.
(Rockefeller University, NY). For depletion of Foxp3 Treg cells, 1µg DT was injected at day 0.
Followed by 0.25µg DT on day 3, 6. Mice were sacrificed on day 7. For Treg depletion and IL-
10R neutralization, mice were injected with 1µg DT at day 0, followed by 0.25µg DT on day 3, 6 and 8. In addition, mice received antiIL-10R blocking antibody (Clone: 1B1.3A BioXcell) or ratIgG1 isotype control (Clone: HRPN BioXcell) 500µg on day 2, 5, 8 and 250µg at day 8. Mice were sacrificed on day 10. All mice were injected by intraperitoneal route (i.p.).
For MCMV infection, five to six week-old mice were infected with MCMV K181 strain, via i.p.
inoculation with 1 × 106 PFU. Mice were sacrificed upon establishment of latent MCMV at
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various times (> 6-7 months post infection in C57Bl/6 mice). Mice were maintained under
specific-pathogen-free conditions at Cincinnati Children's Hospital Medical Center.
Ethics Statement
All animal protocols were reviewed and approved by the Institutional Animal Care and Use
Committee at the Cincinnati Children’s Hospital Research Foundation(CCHRF) under
IACUC2016-0087 (1D03023).
The care and use of laboratory animals at CCHMC is in accordance with the principles and
standards set forth in their Principles for Use of Animals (NIH Guide for grants and Contracts),
the Guide for the care and Use of Laboratory Animals (Department of Health, Education, and
Welfare DHEW, Public Health Service PHS, National Institutes of Health NIH Publ. 8th edition,
Rev.2011). The provisions of the Animal Welfare Acts (P.L. 89-544 and its amendments), and other applicable laws and regulations.
Virological Methods
For plaque assays, dilutions of virus stocks, 10% (w/v) mouse tissue sonicates, and sonicated
leukocyte cell suspensions were adsorbed onto 70% confluent NIH 3T3 monolayers for one hour
at 37oC, and then overlaid with 1:1 carboxymethyl cellulose (CMC): 2X modified Eagle’s
medium as previously described[85]. At 6 days, the overlay was removed and the cells were
fixed with methanol and stained with Giemsa to determine the number of plaques. For
measurement of tissue virus replication following mouse infection, tissues were placed in pre-
weighed tubes or were homogenized in media to prepare cell suspensions, followed by
sonication on ice to disrupt cells and release free virus. Tissue homogenates were titered by
plaque assay similar to virus stocks as described above.
Reactivation Assays
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Explant reactivation assays of tissues from latently infected mice were established as previously
described [45, 65]. After infection when virus was no longer replicating and establishment of
latency (> 6-7 months post infection in C57Bl/6 mice) and following DT treatment, the SGs and spleens were collected. In some experiments, other tissues such as lungs and liver were also collected for plaque assays. The SGs were sonicated and titered to detect persistent replicating virus as the SG is a major site of viral persistence. In some experiments, to analyze populations of infiltrating leukocytes in the SGs, the SGs were first homogenized to prepare a cell suspension, an aliquot was removed for sonication and plaque assay analysis, and the remaining sample was treated as described below under ‘cell isolation’. For the explant reactivation assays, the spleens were minced and primary spleen cultures were established as previously described
[45].In some studies, SGs and lungs were also analyzed by explant reactivation assay. Briefly, the spleen explant assay was established by dividing the spleens into three parts, with each part placed into a well of a 6-well tissue culture plate containing 5 ml of media. The cultures were followed for up to 6 weeks and culture media from each well were collected weekly, sonicated and titered in plaque assays to detect the presence of reactivating or replicating virus. Plaques detected in any wells were counted as a reactivation event.
PCR Analysis
Tissues were isolated from infected and uninfected mice using dissection tools pre-treated with
DNA Away (Molecular BioProducts). DNA was isolated from cell suspensions following tissue homogenization using QIAamp DNA Mini Kit (Qiagen #51306) according to manufacturer’s protocol. Quantitative PCR was performed as previously described[88] using the following primers: E1 forward primer 5’-TCGCCCATCGTTTCGAGA-3’ ; E1 reverse primer 5’-
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TCTCGTAGGTCCACTGACGGA-3’ to yield an amplified 106bp product. The TAMRA
Taqman E1 probe was purchased from Applied Biosystems.
Cell isolation
Spleens were harvested and crushed through 100-µm filters (BD Falcon) to generate single-cell
suspensions. For the SG, tissue was digested with collagenase (4) or with liberase in media
(RPMI 1640 containing 0.5 mg/ml digestion enzyme (Sigma), 5mM CaCl2, 0.2 mg/ml of DNase
I (Sigma) and 5% FBS) incubated for 45minutes at 37°C, followed by a two-step
discontinuousPercoll gradient (Little Chalfont, UK). Gradient samples were centrifuged at
25000rpm, room temperature, for 25 min with the brake off. The lymphocytes were harvested at
the interface between 30% and 70% Percoll layers.
Flow Cytometry
A total of 1 -2 million single-cell suspensions from either spleen or SG surface stained with a
combination of the following tetramers and antibodies:
M45 –HGIRNASFI, m139 – TVYGFCLL, M38 – SSPPMFRV, IE3 – RALEYKNL andM25 –
NHLYETPISATAMVI. Tetramers were all synthesized by the NIH tetramer core facility. Cells were then incubated with Fc block and surface stained with
Surface Abs: αCD4, αCD8α, αTCRβ (BD Biosciences, San Diego, CA, USA), αCD44,
αKLRG1, αCD127, αCD69, αCD103, αCX3CR1, αCD16/32 and αLy6C (eBioscience, San
Diego, CA, USA). For CD8 T cell intracellular staining, cells were stained in media and fixed with 2%Methanol free formaldehyde (MF FA) for 1 hour and then intracellularly stained with
αKi67 (eBioscience) using eBio permeabilization buffer. For CD8 T cell peptide stimulation using M38, M139 at 2µg/ml (a gift from Dr.Edith Janssen, Cincinnati Children's Hospital
Medical Center, Cincinnati,OH) cells were stimulated for 5 h in the presence of anti-CD107a
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antibody and in the presence of brefeldinA for the final 4 h. Cells were then fixed with 2%MF
FA for 1 hour and then in 0.05% saponin. Cells were stained for IFN-γ and TNF-α production
(all from eBioscience). For CD4 T cells, cells were stimulated with 50ng/ml PMA and 1µg/ml
ionomycin for 5 hours, in the presence of brefeldinA for the final 4 h and fixed with 2%MF FA
for 1 hour followed by intracellular staining for IL-10, IFN-γ, TNF-α (all from eBioscience) and
Foxp3 (eBioscience) staining was done according to eBioscience Foxp3 staining kit and protocol. Data were acquired on an LSRII flow cytometer (BD Biosciences) and analyzed using
FACSDiva software (BD Biosciences).
RT-PCR
Samples were homogenized and total cellular RNA was extracted and quantified. DNase-treated
RNA was then used to synthesize cDNA. The primer sequences used for detection of IL-10 were: 5′- GCTCTTACTGACTGGCATGAG -3′ and 5′- CGCAGCTCTAGGAGCATGTG -3′.
The primer sequences used for detection of β-actin as an internal control were
5′-GGCCCAGAGCAAGAGAGGTA-3′ and 5′-GGTTGGCCTTAGGTTTCAGG-3′.
Quantitative real-time PCR was performed with Roche LightCycler 480 SYBRGreen 1 Master
Mix using the Roche LightCycler 480 II instrument (Roche Diagnostics).
Statistical Analysis
Data were analyzed using GraphPad Prism software and Excel software. Statistical analysis was performed using either a Student’s t-test or aFisher’s exact test in different experiments. A p- value of <0.05 was considered significant.
125
Acknowledgements
Thanks to Dr. Ian Humphreys (Cardiff University) for helpful advice on the IL-10R
neutralization strategy. This work was supported by NIH grants AG033057 (to C.A.C and
D.A.H), AG033057-S1 (to M.A.), and NIH R01 AI087683-01A1, Steinmann Family Foundation
CMV Award (R.D.C).
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Figures
Fig.1. Activated Treg are increased in the spleen during latent MCMV. Splenocytes were isolated
from naïve (9.5months, white bars), or aged matched MCMV-latently infected mice (black bars, 8 months post- inoculation with1× 10 pfu of MCMV). Cells were stained for CD4, CD69, and Foxp3 and analyzed by flow cytometry. Dot plots⁶ (A) and bar graph (B) show the frequency of Foxp3+ cells in total CD4+ cells from naïve and MCMV infected mice as described above (mean+SEM). Representative histograms (C) and bar graph (D) show the frequency of CD69+ on gated Foxp3+ cells from naïve and MCMV infected mice as described above (mean+SEM). Data is representative of two independent experiments. Naïve
(N=4), WT MCMV infected (N=6). Statistical analysis (**p ≤ 0.01, Student’s t-test).
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Fig.2. Treg inhibit effector T cell responses during latent MCMV infection in the spleen. 5-6 week old WT C57BL/6 (white bars) and Foxp3DTR (black bars) mice were inoculated with1× 10 pfu of MCMV. 8 months post-MCMV infection, both groups were injected with Diphtheria toxin (DT) on⁶ day 0, 3, 6 and sacrificed on day 7. Splenocytes from infected mice were stained with M25 (CD4 T cell) and M45, IE3, m139, M38 (CD8 T cell) tetramers day 7 (+DT) and analyzed with flow cytometry. Bar graphs (A) show the total number of M25- specific CD4 T cells and M45-, IE3-, m139- and M38-specific CD8 T cells from
MCMV infected WT C57BL/6 (white bars, N=6-7) and Foxp3DTR (black bars, N= 6-8) mice (mean+SEM).
134
Data are pooled from two independent experiments. Spleen cells (2 x 106 cells/well) from MCMV infected
mice day 7 (+DT) were ex vivo stimulated with M38 peptide (5 h at 37°C) and treated with Brefeldin A and
were stained for CD8, and IFN-γ, TNF-α and analyzed by flow cytometry. Representative plots and bar graphs (B) show the average frequency and total number of CD8+ T lymphocytes producing IFN-γ and/or
TNF-α (mean+SEM). WT C57BL/6 (N=6), Foxp3DTR (N=6). Spleen cells from infected groups (+DT) were ex-vivo stimulated with MCMV-peptide (M38) for 5 hours during which CD107α antibodies were incubated with the cells. Representative plots and bar graphs (C) show the average frequency and total number of
CD8+T lymphocytes expressing CD107α (mean+SEM).WT C57BL/6 (N=11), Foxp3DTR(N=10). Statistical analysis, *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 (Student’s t test).
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Fig.3. Treg promote latent MCMV infection in the spleen. 5-6 week old WT control and Foxp3DTR mice were inoculated with 1× 10 pfu of MCMV (N=9/group). 8 months post-MCMV infection, both groups were injected with Diphtheria toxin⁶ (DT) on day 0, 3, 6 and sacrificed on day 7. Spleens were analyzed for reactivation from latency by spleen explant assay as described in methods. Supernatants were assayed for infectious virus by plaque assay weekly. Bar graph (A) indicates the percentage of mice positive for virus reactivation from the spleens in either WT control (white bars) and Foxp3DTR (black bars) mice, with the numbers of positive mice shown above the bars. Bar graph (B) represents average of virus titer of replicating virus in the supernatants of the spleen explant cultures of WT control (white bars) and
Foxp3DTR (black bars) MCMV infected mice day7 post Treg depletion (mean+SEM). Data include all of the mice in the experiment, including those mice with undetectable virus. A total of 1.5 ml of the explant cultures, representing ~37% of the total supernatant, was titered. Mice with undetectable amounts of virus were given a value of zero. Statistical analysis, (Student’s t test) *p ≤ 0.05.
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Fig.4. Treg are required to prevent MCMV reactivation and MCMV-specific CD4+ T cells in the SG.
5-6 week old WT C57BL/6 and Foxp3DTR mice were inoculated with1× 10 pfu of MCMV. 8 months post-
MCMV infection, both groups were injected with Diphtheria toxin (DT) on day⁶ 0, 3, 6 and sacrificed on day
7. Bar graph (A) shows the percentage of WT C57BL/6 uninfected control (UI, N=2 day 0, N=2 day 7, gray bar), MCMV infected (N=3 day 0, N=11 day7 white bars) and Foxp3DTR (N=6 day 0, N=10 day 7,
black bars) mice positive for virus replication in the SGs before Treg depletion (indicated as day0) and
day7 post Treg depletion with the numbers of positive mice in each group shown above the bars. (B) Bar graph shows the average viral titer of individual SGs of MCMV infected mice before Treg depletion
(indicated as day0) and day7 post Treg depletion as described above (mean+SEM). The presence of replicating virus was detected by plaque assay. Samples with no detectable virus were assigned a titer of
0.7 log pfu/ml, the limit of detection for the plaque assay as indicated by the dashed line. Data is representative of five independent experiments. Statistical analysis, *p ≤ 0.05, **p ≤ 0.01 (Student’s t tests) comparing infected mice. Single cell suspensions were generated from the SGs of MCMV infected mice and were stained first with MCMV-tetramers, M38 (CD8), M25 (CD4) and then cells were surface
137 stained for CD8, CD4, and analyzed with flow cytometry. Bar graph (C) shows the total number of M38- specific CD8 T cells in the SGs (mean+SEM). WT C57BL/6 (N=4). Foxp3DTR (N=5). Bar graph (D) shows the total number of M25-specific CD4 T cells in the SGs (mean+SEM). WT C57BL/6 (N=6). Foxp3DTR
(N=8). Statistical analysis, *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 (Student’s t test).
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Fig.5. Treg suppress CD4+ Foxp3- IL-10+ cells in the SG and IL-10 is critical for limiting viral load in the SG after Treg depletion. 5-6 weeks old WT C57BL/6 (white bars) and Foxp3DTR (black bars) mice
were inoculated with 1× 10 pfu of MCMV. 8 months post-MCMV infection, both groups were injected with
Diphtheria toxin (DT) on day⁶ 0, 3, 6 and sacrificed on day 7. Bar graph (A) shows the average normalized
IL-10 mRNA level (mean+SEM) in SG extracts of MCMV-infected mice (day7) post Treg depletion. WT
C57BL/6 (N=11). Foxp3DTR (N=10). Single cell suspensions were generated from the SGs of MCMV
infected mice (day7 post Treg depletion). Cells were stained for TCRß, CD4, CD8, Foxp3 and IL-10
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following stimulation with or without PMA and ionomycin for 5 hours, in the presence of brefeldinA. Bar
graph (B) shows the average of frequency of IL-10+ in Foxp3- CD4+ TCRβ+ and CD8+ TCRβ+ cells and
TCR-β− cells (mean+SEM) in the SG. WT C57BL/6 (N=5). Foxp3DTR (N=5). Statistical analysis, *p ≤ 0.05,
**p ≤ 0.01 (Student’s t test). (C) Schematic representation of diphtheria toxin and IL-10 neutralization treatment. Briefly, 5-6 week old WT C57BL/6 (N=24) and Foxp3DTR (N=26) mice were inoculated with 1×
10 plaque-forming unit (pfu, p.i.) of MCMV. 8-12 months post-MCMV infection, both groups were injected⁶ with DT on day 0, 3, 6, 8 and sacrificed on day 10. Groups were split and were treated with
500µg of either isotype control antibody or with anti–IL-10R neutralizing antibody on day 2, 5, 8 and were sacrificed on day10. Bar graph (D) indicates the percentage of mice positive for virus reactivation in the
SGs (day 10) post Treg depletion and IL-10R neutralization with the numbers of mice in each group shown above the bars. WT C57BL/6 uninfected (UI, N=6), WT C57BL/6 MCMV infected (N=24) and
Foxp3DTR (N=26). The presence of replicating virus was detected by plaque assay (data combined from two independent experiments). Statistics for percentage of mice with virus reactivation which was made using Fisher’s exact test (*P ≥ 0.05, ** P≥ 0.01, *** P≥ 0.001) comparing all groups.
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Supporting information:
Day(0) Naïve WT Foxp3-DTR MCMV MCMV Spleen 0/2 0/3 0/6
Lung 0/2 0/3 0/6 Liver 0/2 0/3 0/6
Pancreas 0/2 0/3 0/6 Salivary 0/2 0/3 0/6 Gland
S1 Table. Establishment of latent MCMV infection. Table shows the number of mice with positive
MCMV titers (replicating virus) in the spleen, lung, liver, pancreas and salivary gland within the three groups: Naïve, WT control and Foxp3-DTR, 8 months post MCMV infection. Titers were quantified via plaque assay before Treg depletion, indicated here as Day0. 0/number of mice in each group indicates absence of actively replicating virus and confirms the establishment of latency in all tissues.
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S2 Table. Absence of actively replicating virus in the spleen in controls and Foxp3-DTR mice 7 days post Treg depletion. Table shows the number of mice with positive MCMV titers (replicating virus) in the spleen within the two groups: WT control and Foxp3-DTR, 8 months post MCMV infection. Titers were quantified by plaque assay 7 days after Treg depletion, indicated here as Day7. 0/number of mice in each group indicates absence of actively replicating virus and confirms the establishment of latency.
(N=9/group).
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S1 Fig. Treg in the spleen during latent MCMV. Splenocytes were isolated from naïve (9.5months), or aged matched MCMV-latently infected mice (8months p.i.). Cells were stained for CD4 and Foxp3 and analyzed by flow cytometry. Graph shows the number of Foxp3+ cells in total CD4+ cells. Naïve (N=4),
WT MCMV infected (N=6).
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S2 Fig. Highly activated, proliferating MCMV-specific CD8+ T cells in the spleen post Treg depletion. 5-6 week old WT C57BL/6 and Foxp3DTR mice were inoculated with 1× 10 pfu of MCMV. 8 months post-MCMV infection, splenocytes were isolated from infected mice. Cells were⁶ stained for IE3, m139, M38 (CD8 T cell) tetramers day 0 (-DT) and analyzed with flow cytometry. A) Graph shows the total number of IE3-, m139- and M38-specific CD8 T from WT C57BL/6 (white bars, N=3) and Foxp3DTR
(black bars, N=6) mice. 5-6 week old C57BL/6 and Foxp3DTRmice were inoculated with1× 10 pfu of
MCMV. 8 months post-MCMV infection, both groups were injected with Diphtheria toxin (DT) on day⁶ 0, 3,
6 and sacrificed on day 7. Spleen cells were analyzed by flow cytometry. B) Bar graphs show the average
frequency and absolute number of CD4+ Foxp3+Treg in the spleen (mean+SEM). WT C57BL/6 (N=11).
Foxp3DTR (N=10). Spleen cells isolated from the two groups were stained with MCMV CD8-specific tetramers and then surface stained for expression of KLRG-1 and CD127 and intra-cellular expression of
Ki67. C) Bar graph shows the total numbers of effector subpopulations within gated m139-specific CD8 T cells (mean+SEM). WT C57BL/6 (N=6), Foxp3DTR (N=6). D) Bar graphs show the frequency and absolute number of Ki67+ cells within M45-, IE3-, m139- and M38-specific CD8 T (mean+SEM). WT C57BL/6
(N=6), Foxp3DTR(N=6). Statistical analysis, *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 (Student’st-test).
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S3 Fig. MCMV viral load in the spleen. Genomic DNA was isolated from the spleens of WT C57BL/6
and DTR mice at day 7 post treg depletion. MCMV E1 was detected by quantitative PCR, and data
expressed as genome copy number per 100 ng genomic DNA as described in Materials and Methods.
Results are pooled from 4 independent experiments (WT, N=20 and DTR, N=22) and show the
mean+SEM. The average reduction in viral load in DTR mice in 4 independent experiments was 62.7% ±
9.4 (one sample t-test p<0.007).
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S4 Fig. Early MCMV viral reactivation post Treg depletion in the SG. 5-6 week old WT C57BL/6
(white) and Foxp3DTR (black) mice were inoculated with 1× 10 pfu of MCMV (N=8/group). 9 months post-
MCMV infection, both groups were injected with Diphtheria toxin⁶ (DT) on day 0, 3 and sacrificed on day4.
A) Bar graph shows the percentage of mice positive for virus reactivation of naïve (UI) and WT MCMV infected mice in the SGs day4 post Treg depletion with the numbers of mice in each group shown above the bars. The presence of replicating virus was detected by plaque assay. B) Bar graph shows viral titers of individually homogenized SGs of naïve and MCMV infected mice Day4 post Treg depletion
(mean+SEM).
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S5 Fig. Efficiency of Treg depletion in the SG. 5-6 weeks old WT C57BL/6 and Foxp3DTR mice were
inoculated with1× 10 pfu of MCMV. 8 months post-MCMV infection, both groups were injected with
Diphtheria toxin (DT) ⁶on day 0, 3, 6 and sacrificed on day 7. A single cell suspension from the SGs of
naïve and MCMV infected mice was analyzed by flow cytometry. A) Bar graphs show the average
frequency and absolute number of CD4+ Foxp3+ Treg in the SG upon DT administration (mean+SEM).
WT C57BL/6 (N=7) Foxp3DTR (N=8). B) Bar graph shows the average frequency of tissue resident memory (CD103+CD69+) cells in the SG within the total M38-specific CD8 T cells (mean+SEM). C57BL/6
(N=4). Foxp3DTR (N=5). Statistical analysis, *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001 (Student’s t-test).
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S6 Fig. Treg suppress CD4+ Foxp3- IL-10+ cells in the spleen. 5-6 weeks old WT C57BL/6 (white
bars) and Foxp3DTR (black bars) mice were inoculated with 1× 10 pfu of MCMV. 8 months post-MCMV
infection, both groups were injected with Diphtheria toxin (DT) on ⁶day 0, 3, 6 and sacrificed on day 7. A)
Single cell suspensions were generated from the spleen of MCMV infected WT C57BL/6 (N=11) and
Foxp3DTR (N=10) mice (day7) post Treg depletion. Cells were stained for TCRß, CD4, CD8, Foxp3 and IL-
10 following stimulation with or without PMA and ionomycin for 5 hours, in the presence of brefeldinA in
the final 4hrs. Bar graph shows the average of frequency of IL-10+ in Foxp3- CD4+ TCRβ+ and CD8+
TCRβ+ cells and TCR-β− cells (mean+SEM). B) Graph shows the average normalized IL-10 mRNA level in spleen of MCMV-infected WT C57BL/6 (N=10) and Foxp3DTR (N=9) mice (day7) post Treg depletion
(mean+SEM). Statistical analysis, *p ≤ 0.05, **p ≤ 0.01 (Student’s t test).
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S7 Fig. Treg promote MCMV replication in the spleen. 5-6 week old WT C57BL/6 (white) and
Foxp3DTR (black) mice were inoculated with 1× 10 pfu of MCMV (N=8/group). 5 months post-MCMV infection, both groups were injected with Diphtheria toxin⁶ (DT) on day 0, 3, 6, 9,12 and sacrificed on day
14. A) Bar graph shows the percentage of mice positive for virus replication in the spleen day14 quantified by plaque assay post Treg depletion with the numbers of mice in each group shown above the bars. Viral titers were 18.6 pfu/ml +/- 15.5 in WT C57BL/6 mice and 2.4 pfu/ml +/- 2.24 in FoxP3-DTR mice; p=0.31. B) Genomic DNA was isolated from the spleens of WT C57BL/6 and DTR mice at day 14 post Treg depletion. MCMV E1 was detected by quantitative PCR, and data expressed as genome copy number per 100 ng genomic DNA as described in Materials and Methods (mean+SEM); p=0.36.
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S8 Fig. IFN-ƴ production upon Treg depletion in the SG.
Single cell suspensions were generated from the SGs of MCMV infected mice (day7 post Treg depletion).
Cells were stained for CD4, Foxp3 and IFN-ƴ following stimulation with or without PMA and ionomycin for
5 hours, in the presence of brefeldinA. Bar graph shows the average of frequency of IFN-ƴ+ in Foxp3-
CD4+ (mean+SEM) in the SG. WT C57BL/6 (N=5). Foxp3DTR (N=5). Statistical analysis, *p ≤ 0.05
(Student’s t test).
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Chapter4
IL-10 PRODUCING TFH CELLS COUNTER INFLAMMAGING BUT SUPPRESS HUMORAL IMMUNE RESPONSES
Maha Almanan1, Jana Raynor1, Ireti Ogunsulire2, Shibabrata Mukherjee1,4, Anna Malyshkina7, Jennifer Ingram3, Ankur Saini1,4, Markus M. Xie6, Theresa Alenghat1,4, George S. Deepe, Jr. 5, Senad Divanovic1, Harinder Singh1,4, Emily Miraldi1,4, Allan Zajac3, Dent, A.L.6, Christoph Hölscher2, Claire Chougnet1,4, and David A. Hildeman1,4¶* 1 Division of Immunobiology, Cincinnati Children's Hospital, University of Cincinnati College of
Medicine, Department of Pediatrics, University of Cincinnati College of Medicine, Cincinnati, OH
45229 and 4 Center for Systems Immunology, Cincinnati Children's Hospital, Cincinnati, OH 45229
2 Infection Immunology Research, Research Center Borstel, Borstel, Germany
3 Department of Microbiology, University of Alabama at Birmingham, Birmingham, AL
5 Division of Infectious Diseases, University of Cincinnati College of Medicine, Cincinnati, OH 45267
6 Department of Microbiology and Immunology, Indiana University School of Medicine, Indianapolis, IN
7Institute of Virology, Essen University Hospital, Essen, Germany
*Corresponding authors Email: [email protected]
Running title:
IL-10-secreting T follicular helper cells Keywords: Treg, IL-10, Tfh, aging, CD4+
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Abstract
Immune responses deteriorate with age and result in the decline of vaccine responsiveness.
Chronic low-level inflammation termed inflammaging is associated with the impairment of adaptive immune responses; however, the underlying mechanisms remain unclear. Here, we show that aged mice exhibit increased systemic IL-10 that is primarily produced by CD4+
FoxP3- T cells. These cells manifest a T follicular helper (Tfh) profile but express low levels of Bcl6 thereby enabling robust IL-10 expression. Consistent with their designation as Tfh10 cells, IL-6 and IL-21 are required for their accumulation with age. IL-21 promotes Tfh10 survival and maintains a systemic balance between IL-6 and IL-10. Importantly, blockade of
IL-10R signaling significantly restores Tfh-dependent antibody responses in aged mice.
Notably, we also found that Tfh10 cells are significantly increased in elderly humans. We propose that Tfh10 cells counter-regulate inflammaging but, in so doing, lead to impaired humoral responses with age.
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Introduction Declining immune function is well described in the elderly, leading to increased risk and severity of infection, poorer control of cancer, and impaired responses to vaccination (Lord,
2013; Zhang et al., 2017). Despite such declining adaptive immune function, aging is also characterized by a persistent low-grade immune activation (so-called inflammaging). Such inflammaging is implicated in many deleterious processes in the elderly, including Alzheimer’s disease, cardiovascular diseases and general frailty (Giunta et al., 2008; North and Sinclair,
2012). Indeed, high levels of circulating, pro-inflammatory IL-6 are associated with increased morbidity and mortality among older individuals (Maggio et al., 2006).
Most inflammatory reactions incite potent anti-inflammatory feedback loops (Hanada and Yoshimura, 2002). Amongst anti-inflammatory mediators, IL-10 has broad-ranging and potent properties and can arise in response to or concurrently with inflammatory stimuli
(Couper et al., 2008). Recent work has shown a significant age-related increase in serum IL-
10 in a cross-sectional analysis of more than 465 subjects, ranging in age from 21 to 88 years
(Lustig et al., 2017). In addition, an IL-10 genotype (-21082GG) that is associated with high production of IL-10 in Caucasians was more prevalent in centenarians than in younger individuals (65-73 yrs) (Lio et al., 2002), and similarly more prevalent in middle-aged controls than in age-matched patients with myocardial infarction (Lio et al., 2004). Also, elderly men with the highest serum levels of inflammatory cytokines, or with the lowest levels of IL-10, had the highest risk of frailty-associated pathologies (Cauley et al., 2016). Thus, IL-10 levels, appears to play an important role in counter-regulation of inflammation to promote healthy aging.
In contrast to its beneficial roles in aging, IL-10 limits protective responses to pathogens. IL-10 plays a deleterious role in chronic infections, in mice and humans, limiting microbial clearance (Belkaid et al., 2001; Brooks et al., 2006). Indeed, we recently showed 153 that IL-10 fosters reactivation and replication of cytomegalovirus in the salivary glands of latently infected mice (Almanan et al., 2017). Further, IL-10 dampens responses to infections or vaccines (Dobber et al., 1995; McKinstry et al., 2009). In fact, in humans, the IFN-γ:IL-10 ratio appears to be a critical predictor to protective immunity in the elderly (McElhaney et al.,
2006). Despite these proposed roles for IL-10, little is known about its function, its cellular source, or its molecular control in aged animals or humans.
IL-10 can be produced by many cells, including those of the innate immune system
(notably multiple myeloid cell subsets), the adaptive immune system (T cells and B cells), and even non-immune cells (e.g., keratinocytes and hepatocytes) (Moore et al., 2001). At baseline in the spleen of young mice, the majority of IL-10 expression is localized to B cells and CD4+
T cells (CD25+ and CD25-) (Madan et al., 2009). In aging, a few studies have investigated the cellular sources of IL-10 and suggested a role for memory CD4 T cells. Indeed, early work showed increased IL-10 production by aged CD44hi CD4+ T cells (Hobbs et al., 1994). More importantly, the proportion of IL-10+ influenza-specific CD4+ T cells increases in aged mice
(Lefebvre et al., 2016b). In contrast, B cells capable of IL-10 production appear to be decreased in older subjects (van der Geest et al., 2016). Nonetheless, the major cellular source of IL-10 in aging remains unclear.
Here, we report that systemic levels of IL-10 are increased in aged mice and negatively impact vaccine responsiveness. Notably, we found that CD4+ FoxP3-, not classic FoxP3+, cells were required for increased systemic IL-10 levels in aging. Further, these IL-10- producing T cells bore markers of T follicular helper cells (Tfh), were present in both mice and humans, and they required IL-6 for their accumulation. Interestingly, IL-21, another promoter of Tfh homeostasis, was also required for the accrual of these cells, and, importantly, to regulate the systemic balance between IL-6 and IL-10. Mechanistically, we also found that the canonical Tfh transcription factor, BCL6, was downregulated with age in Tfh cells, permitting 154 their IL-10 production. Together, our data show that inflammation and anti-inflammation are linked via IL-21 production, which promotes accrual of IL-10-secreting Tfh (Tfh10) cells that function to dampen immune responses and IL-6-driven inflammaging.
Results: Aged mice have increased systemic levels of IL-10. While IL-10 levels have been shown to increase in aged humans (Lustig et al., 2017), it is unclear if IL-10 levels increase in aged mice. Using the sensitive in vivo cytokine capture assay (IVCCA) we found that steady-state levels of IL-10 in the serum were increased 2-3 fold in old compared to young mice (Figure 1A). To determine the potential sources of this enhanced IL-10, we examined various lymphoid and non-lymphoid tissues and found an increase in IL-10 mRNA in the epididymal white adipose tissue (WAT), lymph nodes, and spleen of aged, compared to young mice (Figure 1B). These data show that the systemic levels of IL-10 are increased with age and that secondary lymphoid organs appear to be major contributors of augmented IL-10 expression in aging.
CD4+ FoxP3- T cells are the major source of IL-10 in aged mice.
To identify cells with enhanced IL-10 production in aged mice, we took advantage of IL-
10-reporter (VertX) mice, which possess an IL-10-IRES-eGFP cassette in the endogenous IL-
10 locus (Madan et al., 2009). VertX mice allowed us to examine baseline IL-10-production directly ex vivo, in the absence of exogenous stimulation, as GFP levels in these mice directly correlate with IL-10-production (Madan et al., 2009). Flow cytometric analysis of spleen cells in aged versus young VertX mice revealed a significantly increased frequency of GFP+ (IL-
10+) cells in multiple cell types, but the largest increase was observed in CD4+ T cells (Figure
2A). As B cells are the predominant immune cell type in the spleen, the total numbers of
GFP+ B cells were increased with age. However, there was no significant difference in the 155 numbers of aged IL-10+ B cells compared to IL-10+ CD4+ T cells (Figure 2A). Instead, the largest increase in the frequency of IL-10-producing cells was in CD4+ T cells (Figure 2A). In addition, the level of IL-10 produced per cell was significantly higher in CD4+ T cells than in either CD8+ T cells, CD19- or CD19+ B cells (Figure 2A).
Because FoxP3+ regulatory T cells (Treg) are a well-known source of IL-10 in young mice, and their frequency is increased in old mice (Raynor et al., 2012), we next determined whether they were the major contributor to this increased IL-10 in aged mice. Staining for
FoxP3 in VertX mice while maintaining GFP expression is technically infeasible, so we sorted naive (CD4+CD44loCD62LhiFoxP3GFPneg), memory (CD4+CD44hiCD62LloFoxP3GFPneg) and regulatory (FoxP3GFPpos) T cells from young and aged FoxP3-DTR-GFP mice (Kim et al.,
2007), stimulated them with PMA and Ionomycin (P+I), and measured their production of IL-
10 by ELISA. As expected, naïve T cells produced little IL-10 whether they were from young or old mice (Figure 2B). IL-10 production from FoxP3+ Treg was slightly increased in aged mice (~2-fold), (Figure 2B). However, IL-10 production from aged FoxP3- memory T cells was increased >10-fold (Figure 2B). Similarly, flow cytometric analysis of spleen cells of WT mice showed that the frequency of IL-10-producing CD4+ FoxP3+ cells was increased slightly with age, while IL-10-producing FoxP3- CD4+ T cells were ~10-times more frequent with age
(Figure 2C). Again, Foxp3- CD4 T cells showed significantly higher expression of IL-10 per cell compared to their young counterparts and aged Foxp3+ cells (supplementary figure 1).
Together, these three independent approaches show that CD4+ FoxP3- cells have the highest capacity for IL-10-production in the spleens of aged mice. In addition, they are required for the increased systemic levels of IL-10, as depletion of >95% of CD4+ T cells in the spleens of old mice nearly returned the serum levels of IL-10 to levels observed in young mice (Figure 2D). In contrast, depletion of FoxP3+ T cells increased systemic IL-10 levels
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(Figure 2D) and the frequency of IL-10-producing CD4+ T cells (Figure 2E). Thus, FoxP3-, but not FoxP3+, CD4+ T cells are required for the increased systemic levels of IL-10.
Accrual of IL-10-producing CD4+ FoxP3-T cells occurs in germ-free animals.
Recent work has shown that the microbiome changes with age (Odamaki et al., 2016;
Thevaranjan et al., 2017). Further, alterations in the microbiome can affect IL-10 production from CD4+ FoxP3+ and FoxP3- T cells (Mazmanian et al., 2008; Round and Mazmanian,
2010). To test whether the microbiome affects the accumulation of IL-10-producing CD4+ T cells, we aged several cohorts of mice in a germ-free facility. Interestingly, the accumulation of IL-10+ CD4+ FoxP3- cells was similar between age-matched mice housed under specific pathogen free conditions and germ-free animals across a range of ages (supplementary figure 2). Interestingly, we also show that age-driven changes to the microbiome do not appear to alter the accrual of IL-10-producing CD4+ T cells as these cells accumulate in germ-free mice. Further, the age-driven accrual of IL-10-producing CD4+ T cells occurred at four different institutions including: Cincinnati Children’s Hospital, Indiana University/Purdue
University Indianapolis, the University of Alabama-Birmingham, and the Research Center
Borstel in Germany. It is unlikely that the microbiomes of mice are the same at these different institutions. Therefore, the microbiome is not required for the accumulation of IL-10-producing
CD4+ FoxP3- T cells.
IL-10-producing CD4+ FoxP3- T cells in aged mice are predominantly Tfh cells.
Several distinct subsets of CD4+ FoxP3- T cells have been reported to produce IL-10, predominantly Th2, type I regulatory (TR1) T cells, “exTh17” cells, and exTreg cells (Gagliani et al., 2015; Roncarolo et al., 2014; Wang et al., 2005). Although they expressed LAG3, it is unlikely that the majority of the IL-10-producing cells were TR1 cells as they lacked expression of CD49b (supplementary figure 3A), an important marker on TR1 cells (Gagliani 157 et al., 2013). Very few IL-10+ CD4+T cells were capable of IL-4 or IL-17A co-production, ruling out the possibility that these were Th2 or Th17 cells (supplementary figure 3B). Next, analysis of IL-17A fate tracking mice (Hirota et al., 2011) revealed that the frequency of
“exTh17” cells within IL-10+ FoxP3- CD4+ T cells from aged mice was ~1% (supplementary figure 3C). Analysis of exTreg cells using FoxP3-CreRosaloxstoplox dTomato mice (Zhou et al.,
2008) (Madisen et al., 2010) revealed that ~20% of the IL-10+CD4+ T cells were dTomato+
GFP- “exTregs” in both young and old mice (supplementary figure 3D).Thus, neither Th2,
TR1, exTh17, nor exTreg make up the bulk of the IL-10-producing CD4+ T cells that accumulate in aged mice.
In our investigation of cytokine co-production by IL-10-producing CD4+ T cells, we found that the frequency and total numbers of IL-10+ cells that co-produced IL-21 was significantly increased in aged, compared to young, mice (Figure 3A). As IL-21 is typically produced by T follicular helper (Tfh) cells, we next assessed the frequency of IL-10+ CD4+ FoxP3- T cells that expressed CXCR5 and PD1, two canonical surface markers of Tfh cells, in conjunction with the transcription factor BCL6 (Haynes et al., 2007; Johnston et al., 2009). Strikingly, we found that the majority of IL-10+ FoxP3- CD4+ T cells were CXCR5+ and PD1+ in old mice
(Figure 3B). Further, there was a progressive age-related accrual of CXCR5+ PD1+ Tfh cells, including those that produce IL-10 (supplementary figure 4). Thus, the majority of the IL-10- producing T cells that accumulate with age bore markers of Tfh cells so for clarity, we will refer to them as Tfh10 cells.
IL-6 is required for Tfh10 development and for systemically increased IL-10 in aged mice.
We next examined the role of IL-6 in the accrual of Tfh10 T cells because IL-6: (i) controls Tfh development (Eto et al., 2011); (ii) promotes IL-10-production from CD4+ T cells (Jin et al.,
158
2013); and (iii) is a key inflammatory cytokine that is increased with age (Volpato et al., 2001).
To determine whether IL-6 promotes the accrual of Tfh10 cells with age, we aged IL-6-/- mice
≥16 mo and examined the proportion of Tfh10 cells. While no difference in the Tfh cells
(including those that produce IL-10) was observed between young WT and IL-6-/- mice, aged
IL-6-/- mice exhibited a dramatic reduction in the frequency and total number of Tfh and Tfh10 cells compared to aged WT mice (Figures 4A, 4B). Consistent with Tfh10 T cells being a major source of IL-10 in vivo, we found that systemic levels of IL-10 were significantly decreased in aged IL-6-/- mice (Figure 4C). To determine whether IL-6 was required for the development or survival of IL-10-producing FoxP3- CD4+ T cells, we blocked IL-6 after Tfh cells were formed and found that neutralization of IL-6 did not reduce the frequency or numbers of IL-10-producing cells (Figure 4D). Thus, IL-6 is required for the accrual of Tfh10 cells, likely by promoting their initial development.
IL-21 promotes accumulation of Tfh10 cells and regulates systemic IL-6/IL-10 balance.
As IL-21 is a critical cytokine produced by Tfh cells (Nurieva et al., 2007), we next examined whether IL-6 promoted IL-21 production by CD4+ T cells. As expected, and consistent with elevated Tfh cells with age, the proportion and absolute number of IL-21+ CD4+ T cells was significantly increased in aged, compared to young, mice (Figure 5A). However, in the absence of IL-6, the frequency and total numbers of IL-21-producing CD4+ T cells was completely abrogated (Figure 5A). As IL-21 is also critical for the development and homeostasis of Tfh cells (Vogelzang et al., 2008), we reasoned that IL-21 could contribute to the accrual of Tfh10 T cells with age. Similar to aged IL-6-/- mice, the loss of IL-21 prevented age-driven accrual of Tfh cells (Figure 5B) including those that produce IL-10 (Figure 5C).
Again, consistent with the loss of Tfh10 T cells, levels of systemic IL-10 were reduced in aged
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IL-21-/- mice compared to aged wild type controls (Figure 5D). Strikingly, the levels of IL-6 were increased in IL-21-deficient aged mice (Figure 5E). Together, these data show that IL-
21 is critical to balance systemic inflammation (e.g IL-6/IL-10 levels), likely by promoting the accrual of Tfh10 cells. As IL-6 and IL-21 have been reported to increase ICOS which is critical for survival of Tfh cells (Akiba et al., 2005), we considered the possibility that increased levels of ICOS on aged Tfh cells could be contributing to their accumulation. Interestingly, we found a significant but marginal effect of ICOS-L neutralization on overall Tfh cell number and no effect on IL-10 producing cells (supplementary figure 5). These data show that IL-21 plays a key role in promoting accrual of Tfh10 cells with age, whose production of IL-10 likely feeds back to suppress IL-6.
IL-21 promotes repression of Bim in aged Tfh10 cells leading to their enhanced survival.
The accumulation of Tfh10 cells with age could be due to their increased proliferation and/or increased survival. The frequency of Tfh10 T cells that stained positive for the proliferation marker Ki-67+ actually decreased with age, ruling out the possibility that increased proliferation explains their accrual (Figure 6A). Given our and others previous data implicating the pro-apoptotic molecule Bim in T cell survival, (Chougnet et al., 2011;
Tsukamoto et al., 2010), we examined the role of Bim in the survival of IL-10-producing CD4+
T cells. First, Bim levels were reduced in IL-10-producing CD4+ T cells from aged compared to young mice (Figure 6B). Second, the frequency and total number of CD4+ FoxP3- T cells that were IL-10+ was significantly increased in Bim-deficient mice, as early as 6 months of age (Figure 6C). Given that IL-21 promotes accumulation of Tfh10 cells, we next determined whether IL-21 contributed to their reduced expression of Bim. Indeed, IL-21 was critical to suppress the levels of Bim within Tfh10 cells, which likely contributes to their increased
160 survival (Figure 6D). Together, these data suggest that IL-21-driven suppression of Bim contributes to the accumulation of IL-10-producing cells by enhancing their survival.
Tfh10 cells in aged mice manifest diminished levels of BCL6 thereby enabling IL-10 expression.
BCL6 is essential for Tfh differentiation and is induced by IL-6 and IL-21, so we examined
BCL6 levels in young versus aged Tfh cells. Interestingly, BCL6 levels were actually decreased in aged Tfh10 cells (Figure 7A). Further, decreased levels of BCL6 were associated with higher levels of IL-10 (Figure 7A). To determine whether BCL6 is required for the accrual of Tfh cells as well as their production of IL-10 in aged mice we utilized CD4Cre-
BCL6f/f mice that have a T cell-specific loss of BCL6 (Hollister et al., 2013). As expected, given that BCL6 is critical for promoting Tfh cells (Yu et al., 2009), CD4Cre-BCL6f/f mice had a significant loss of CXCR5 and PD1 expressing CD4+ Foxp3- T cells (Figure 7B). Strikingly, already by one year of age, the loss of BCL6 led to a significant increase in the frequency and total number of CD4+ FoxP3- cells that produced IL-10 (Figure 7C). As BCL6 has been reported to suppress expression of Blimp1 and Blimp1 has been shown to promote IL-10 expression from T cells (Neumann et al., 2014), we next examined Blimp1 levels in aged mice with and without BCL6. Interestingly, the levels of Blimp1 did not change in CD4+ FoxP3- cells that produced IL-10 from aged mice, whether or not BCL6 was present, making it unlikely that Blimp1 is promoting increased IL-10 expression in aged CD4+ FoxP3- T cells
(supplementary figure 6). Further, our prior data show that the additional loss of Blimp did not reduce IL10-producing CD4+ FoxP3- T cells in BCL6-deficient mice (Xie et al., 2017). Thus,
Bcl-6 suppresses IL-10 production, likely via a Blimp1-independent mechanism.
IL-10 limits Tfh-dependent vaccine responses in aged mice.
We next sought to determine the physiologic relevance of Tfh10 cells with age. As vaccine 161 responsiveness is a major problem in elderly humans and Tfh cells are critical regulators of vaccine responses (Crotty, 2011), we used a classic mouse model of a Tfh-dependent antibody response, immunization with NP-KLH. We reasoned that, if Tfh10 cells were important for regulating vaccine responses, then limiting IL-10 signaling should affect vaccine responsiveness. As reported before (Sage et al., 2015), we confirmed that old mice displayed a significantly lower level of anti-NP antibody production, as well as significantly lower frequency and total numbers of NP-specific B cells compared to young mice (Figure 8A).
Importantly, neutralization of IL-10R during NP-KLH immunization significantly restored anti-
NP antibody production as well as the frequency and numbers of anti-NP-specific B cells to levels close to those observed in young mice (Figure 8B). Thus, IL-10 limits Tfh-dependent B cell responses in aged mice.
Tfh10 cells accumulate during aging in humans.
Given the above data in mice implicating Tfh10 cells as regulators of vaccine responsiveness, and the fact that vaccine responses decline significantly in aged humans (Lord, 2013), we next determined whether Tfh10 cells accumulated in aged humans. The fact that Tfh cells are located, and function, in secondary lymphoid organs, required us to analyze their proportion in the spleens of young and old organ donors. Importantly, the frequency of Tfh cells (CXCR5+
PD-1+) was increased in aged humans (Figure 9A). Because flow cytometric analysis of cytokines is affected by cryopreservation, we FACS-sorted memory CD4+ T cells (CD45RO+) into Tfh (CD25-CD127+PD-1+CXCR5+), Treg (CD25+CD127-PD-1-CXCR5-) and other memory cells (CD25-CD127+PD-1-CXCR5-) and analyzed their production of IL-10 and IL-21 after in vitro re-stimulation with anti-CD3/CD28 beads. As expected, IL-21 production was largely limited to Tfh cells and was significantly increased with age (Figure 9B). Strikingly, the population with the highest production of IL-10 was the old Tfh cells (Figure 9C). Thus, similar
162 to mice, Tfh10 cells accumulate in aged humans and may explain the well-known age-related impairment in vaccine responsiveness.
DISCUSSION Our data show a novel linkage between two age-related phenomena, inflammaging and immune suppression. Notably, we found that IL-6 (a hallmark of inflammaging) is critical for the emergence of IL-10-expressing Tfh cells in aged mice. IL-10 is a critical as a feedback mechanism to dampen IL-6-driven inflammaging. However, the increased IL-10 also results in the impairment of vaccine responsiveness. Thus, further understanding of the mechanism(s) underlying the regulation and function of Tfh10 cells may provide insights and potential therapeutic approaches to harness the beneficial effects of IL-10 (minimizing systemic inflammation and organ dysfunction, (Couper et al., 2008) while reducing the harmful effects of IL-10, one of which is the well-known reduction in vaccine responses with age.
Although we showed that IL-10 limits GC B cell responses to an NP-KLH vaccine in aged mice, the targets of IL-10 remain unclear. The effects of IL-10 on B cells is controversial, some studies suggest that IL-10 promotes the survival of germinal center (GC) B cells (Levy and Brouet, 1994). On the other hand, others have shown that IL-10 limits GC B cell responses, consistent with our data (Cai et al., 2012). One potential explanation for these seemingly disparate observations may be due to the use of in vitro versus in vivo model systems. Indeed, much of the work showing positive role of IL-10 on GC B cells has been done in vitro; while work showing a negative role of IL-10 on GC B cells has been done in vivo. Nonetheless, more work is required to understand the cellular targets of IL-10 that are required to suppress antibody responses.
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While cell intrinsic defects in adaptive immune cells clearly contribute to age-related immune suppression, our data show a substantial contribution of IL-10. These data suggest that cell intrinsic defects can be at least partially reversed by IL-10 neutralization or inhibition.
This is consistent with prior work showing that pro-inflammatory stimuli can restore vaccine responsiveness in aged mice (Maue et al., 2009). Similarly, in aged humans a strong pro- inflammatory adjuvant MF59 was able to significantly boost antibody responses to influenza
(Della Cioppa et al., 2012). In both studies, increases in IFN-γ producing CD4+ T cell responses was observed. All together, these data show that the aged immune system is amenable to at least partial restoration by manipulating the inflammatory environment.
Our work is also consistent with prior data showing an increase in Tfh cells in aged mice (Lefebvre et al., 2016a) (Lefebvre et al., 2016b). These prior studies suggested that subtle maturation defects in aged Tfh cells (slightly decreased levels of GL7, CXCR5 and
ICOS), accounted for their decreased ability to provide help to B cells. However, their homing to GCs was not significantly affected compared to young mice when the data were corrected for GC size (Lefebvre et al., 2016b). Further, although these authors observed an increase in
IL-10-producing CD4+ T cells after influenza infection in aged mice, this increase was not attributed to Tfh cells, but rather to Treg or T follicular regulatory cells (Lefebvre et al., 2016b).
However, in our aged cohorts of mice, neither Tfr nor Treg appear to be substantial contributors to IL-10 production in vivo. At baseline, without acute infection, we found that IL-
10-producing Tfh cells are required for elevated systemic levels of IL-10 in aged mice.
Notably, we show that IL-6 and IL-21 are individually required to drive the accumulation of Tfh10 cells, consistent with prior work suggesting a unique role for each cytokine in Tfh development and homeostasis (Choi et al., 2013). Our data also suggest a role for IL-6 in the development, but not the maintenance of Tfh10 cells. Indeed, we saw a precipitous loss of
Tfh10 cells in IL-6-deficient mice, but no change in Tfh10 cells when IL-6 was neutralized after 164 their formation, similar to a recent study in which IL-6 was neutralized during development of a Tfh response in aged mice (Tsukamoto et al., 2015). Part of this IL-6-driven developmental program involves induction of IL-21 production (Nurieva et al., 2008), which our and others data suggest may be required for the long-term maintenance of Tfh and Tfh10 cells
(Linterman et al., 2010). Our data also provide some molecular insight into the role of IL-21 on maintenance of Tfh10 cells in that we show it is required to suppress their expression of
Bim, which regulates their long-term survival. This concept of dual cytokines individually promoting Th cell development vs maintenance may be a common theme for Th cells as a similar phenomenon was recently observed in Th2 cells with TSLP and IL-4 (Rochman et al.,
2018).
Paradoxically, unlike the loss of IL-6 and IL-21, the loss of BCL6, a transcription factor essential for Tfh development, actually enhanced the accrual of IL-10-producing CD4+
FoxP3- T cells. Although we were unable to ascribe a Tfh status to these cells as the Tfh signature markers CXCR5 and PD1 were both substantially reduced in the absence of BCL6, our data clearly show that BCL6 is a major negative regulator of IL-10 production. This result likely reveals the dual nature of BCL6. On the one hand, BCL6 is critical for Tfh development and for expression of the canonical Tfh markers CXCR5 and PD1, while on the other hand,
BCL6 represses IL-10 expression. Indeed, we find little IL-10-production from Tfh cells in young mice, who maintain high expression of BCL6. However, with age, BCL6 levels decline and IL-10 production from Tfh cells increases.
Repression of IL-10 by BCL6 in Tfh10 cells could occur via two, non-mutually exclusive mechanisms. First, BCL6 could act indirectly, through repression of Prdm1 (Blimp1) expression (Johnston et al., 2009), as Blimp1 promotes IL-10 expression in both CD4+ and
CD8+ T cells (Neumann et al., 2014). However, we recently showed that the additional loss of Blimp1 did not reduce enhanced IL-10 production in the absence of BCL6 in young mice 165
(Xie et al., 2017). Second, BCL6 could act directly to repress IL-10 expression by binding to sites in the IL-10 locus (Hollister et al., 2013). Given the negative data implicated Blimp1, our data are more consistent with a scenario in which BCL6 represses IL-10 expression in young
Tfh cells, however, the loss of BCL6 with age de-represses IL-10 production from Tfh cells, although this remains to be rigorously tested.
Importantly, our data may provide an explanation for prior work in elderly humans showing that the antigen-stimulated IFN-γ:IL-10 ratio is directly correlated with cell-mediated immunity to influenza infection and inversely correlated with the severity of infection
(McElhaney et al., 2016) and also with seroconversion following influenza vaccination (Corsini et al., 2006). Our data shows that Tfh cells are a major source of T cell derived IL-10 in aged mice and possibly in elderly humans and that blockade of IL-10 signaling largely restored vaccine responses in mice. These data suggest that transient blockade of IL-10 could be a novel strategy to enhance vaccine responses in the elderly and, due to its transient nature, is unlikely to have untoward effects on autoimmunity, cardiovascular disease or frailty.
Materials and Methods: Mice: Young (6weeks-4months) C57BL/6 mice were purchased from Taconic Farms
(Germantown,NY). Old (≥16months) or Middle age (12-15months) C57BL/6 mice were from
National Institutes of Aging colony located at Charles River Laboratories (Wilmington, MA).
Foxp3-IRES-DTR-GFP knock-in C57BL/6 mice (Kim et al., 2007), were a generous gift from
Dr. A. Rudensky and were aged in house. Bim-deficient (Bim knockout) mice were originally a kind gift from Drs. P. Bouillet and A. Strasser and were bred in-house. IL-6–deficient (IL-6 KO) mice on the C57BL/6 background were aged in-house. IL-10-reporter (VertX) mice which possess an IL-10-IRES-eGFP cassette in the endogenous IL-10 locus on the C57BL/6
166 background (Madan et al., 2009), were aged in-house. Young, middle age and old Germ-free mice on the C57BL/6 background were maintained in isolator units in the CCHMC Gnotobiotic
Mouse Facility. Young and old FoxP3-fate mapping mice (Foxp3CreRosa26dTomato) on the
C57BL/6 background were kindly provided by Dr. Sing S. Way (CCHMC). IL-17A fate tracking mice IL-17CreRosa26eYFP (Hirota et al., 2011) on the C57BL/6 background were bred and aged under specific-pathogen free conditions in the animal facility of the Research Centre
Borstel, Germany. Young and old IL-21–deficient (IL-21KO) mice on the C57BL/6 background were bred, maintained and aged in fully accredited facilities at the University of
Alabama at Birmingham. Spleens (controls and IL-21KO) were shipped overnight on ice and analyzed in Cincinnati. CD4Cre BCL6f/f mice on the C57BL/6 background were bred, maintained and aged in fully accredited facilities at the University of Indiana. Spleens (CD4Cre
BCL6f/f and control) were shipped overnight on ice and analyzed in Cincinnati. All animal protocols were reviewed and approved by the Institutional Animal Care and Use Committee at the Cincinnati Children’s Hospital Research Foundation (IACUC 2016-0087).
Immunization, neutralization and depletion treatments:
For depletion of Foxp3 Treg cells in old FoxP3-DTR mice, 1.25µg DT/mouse was injected i.p. and were sacrificed two days later. For CD4 T cell depletion, mice were injected with a single dose of 600ug /mouse of anti-CD4 i.p. (Clone: YTS191 BioXcell) or isotype control (Clone:
LFT-2 BioXcell) and were sacrificed two days later. For T cell–dependent immunization, mice were immunized intraperitoneally with 100 µg NP-KLH (Biosearch Technologies) mixed with
50% (vol/vol) alum (Thermo Scientific) and sacrificed 20 days later. For IL-10R neutralization, mice were injected with antiIL-10R blocking antibody (Clone: 1B1.3A BioXcell) or ratIgG1 isotype control (Clone: HRPN BioXcell) at day -1 (1mg), day 1 (250ug), day 3 (500ug), day 6
(500ug), day 8 (250ug) and were sacrificed 10 days after immunization. For IL-6 neutralization 167 mice were injected i.p. with 300ug α-IL-6 (Clone: MP5-20F3, BioXcell) or 300ug isotype control (Clone: HRPN BioXcell) on days 0 and sacrificed on day 2. For ICOS-L neutralization old C57BL/6 mice were injected i.p. with 150µg anti-ICOSL (HK5.3, BioXcell) or with rat
IgG2A isotype control (2A3, BioXcell), on days 0, 3, 6, 9 and sacrificed on day 12.
In vivo cytokine capture assay and ELISAs: IL-6 and IL-10 in vivo cytokine capture assay was performed as previously described
(Finkelman et al., 2003) employing biotinylated capture antibodies (Invitrogen). In brief, young
(1.5–4 mo) and old (≥16 mo) C57BL/6 mice were injected i.v. with 10 ug biotinylated anti–IL-6
(MP5-32C11; Invitrogen) and anti–IL-10 (JES5-16E3: Invitrogen)) capture antibodies; mice were bled within 24 h and serum was collected. A luminescent ELISA was performed using anti–IL-6 (MP5-20F3; Invitrogen) or anti-IL-10 (JES5-2A5: BD Biosciences) as the coating antibody. For NP-specific antibody titers, 96-well plates were coated overnight at 4 °C with
NP30-BSA (Biosearch), followed by blockade of nonspecific biding by incubation for 1-2 h at
25 °C with 5% BSA. Serum samples were loaded into plates with eight serial dilutions
(starting from 1:100 or 1:1000), followed by incubation for 2 h at 25 °C or overnight at 4 °C.
After samples were washed, horseradish peroxidase (HRP)-conjugated goat antibody to mouse IgG1 (PA1-74421; Thermo) was added to plates, followed by incubation for 2 h at 25
°C. The reactions were developed by incubation for 15 min at 37 °C with 50 µl TMB substrate
(BioLegend) and were stopped by the addition of 25 µl 10% H3PO4. The plates were read at
450 nm and 570 nm (for correction) with an enzyme-linked immunosorbent assay reader.
RT-PCR:
Samples from different tissues were homogenized and total cellular RNA was extracted and quantified. DNase-treated RNA was then used to synthesize cDNA. The primer sequences used for detection of IL-10 were: 5′- GCTCTTACTGACTGGCATGAG -3′ and 5′-
168
CGCAGCTCTAGGAGCATGTG -3′. Expression levels were normalized to S14 as internal control gene. The primer sequences used for S14 detection were 5′- GAG GAG TCT GGA
GAC GAC GA-3′ and 5′- TGG CAG ACA CCA AAC ACA TT-3′. Quantitative real-time PCR was performed with Roche LightCycler 480 SYBRGreen 1 Master Mix using the Roche
LightCycler 480 II instrument (Roche Diagnostics). Each reaction was performed in triplicate.
Flow cytometry and cell sorting: Human studies Spleen cells from young (median: 18.8, range 18-26 yrs, 3 males, 5 females) and old
(median: 62, range 60-67 yrs, 4 males, 4 females) organ donors with no immunological condition were rested overnight in RPMI medium supplemented with 10% fetal calf serum
(FCS), 1% penicillin, streptomycin, glutamine and 0.5% HEPES, at 37o C and 5% CO2. The cells were then washed with PBS+2%FCS and stained for CD4, CXCR5, PD-1, CD45RO
(Biolegend), CD25 (BD Bioscience), CD3 (Invitrogen), and CD127 (Beckman Coulter) for 30 mins in 4o C, fixed with 4% para-formaldehyde for 20 mins in 4o C. Cells were stained for
Foxp3 (Invitrogen) using Invitrogen Foxp3 permeabilization buffer and acquired on a flow cytometer. For sorting, CD4+ T cells were bead-purified by negative selection from spleen cells, surface stained with Abs against CD45RO, CD127, CD25, PD-1, CXCR5 and the following populations were sorted by FACS after gating on memory CD4+ T cells (CD45RO+)
: Tfh (CD25-CD127+PD-1+CXCR5+), Treg (CD25+CD127-PD-1-CXCR5-) and non Tfh memory cells (CD25-CD127+PD-1-CXCR5-). 10,000 cells were stimulated in vitro with anti-
CD3/CD28 beads at a 1:1 cell: bead ratio, or unstimulated. After 16h, supernatants were collected and analyzed by Luminex.
Mouse studies Spleens were harvested and crushed through100-mm filters (BD Falcon) to generate single- cell suspensions. A total of 2X106 cells plated incubated with Fc block and were surface 169 stained with a combination of the following Abs for surface staining: -CD4, -CD8α, -TCRβ, -
LAG3, -Fas (BD Biosciences), -CD19, -PD1, -CXCR5, -GL-7, CD49B (Invitrogen), B220,
IgG1(Biolegend). Cells were intracellularly stained with antibodies against Bim (Cell Signaling
Technology), Ki67, Foxp3 (Invitrogen), BCL6 (BD Biosciences). For cytokine staining, cells were stimulated with 25ng/ml PMA and 0.5μg/ml ionomycin for 5 hours, in the presence of brefeldinA for the final 4 h and fixed with 2%methanol-free formaldehyde for 1 hour followed by intracellular staining for IL-10, IFN-γ (Biolegend), IL-17, IL-4 (Invitrogen) using Invitrogen
Foxp3 permeabilization buffer. For IL-21 staining, cells were fixed, permeabilized with perm buffer from Invitrogen and incubated with IL-21R/Fc (R&D systems) chimera for 45mins-1hr at
4°C. Cells were then washed with perm buffer and stained with AF488 or AF-647-conjugated affinity-purified F(ab')2 fragment of goat anti–human Fcγ antibody (Jackson ImmunoResearch
Laboratories) for 45-1hr at 4°C. Data were acquired on an LSRII flow cytometer (BD
Biosciences) and analyzed using FACSDiva software (BD Biosciences) or FlowJo software
(FlowJo, Ashland, OR). For sorting, spleen cells from young (3 mo, n=3) and old (≥18 mo, n=4) Foxp3-IRES-DTR-GFP mice were enriched for CD4+ T cells using the negative selection
MACS CD4+ T cell isolation kit II (Miltenyi Biotec, San Diego, CA). Enriched cells were stained with anti-CD4, -CD44, -CD62L Abs, and the following populations were sorted by a
FACSAria (BD Biosciences): CD4+ Foxp3GFP+ (Treg), CD4+ Foxp3-GFP- CD44lo CD62Lhi
(naive CD4+), CD4+ Foxp3-GFP- CD44hi CD62Llo (memory CD4+). All three populations were stimulated with 50ng/ml PMA and 1μg/ml ionomycin and supernatants were collected after 15 hours.
Single cell RNA-seq:
Young and old CD4+GFP+ CD25-CD69- cells were sorted and post sort analysis showed that
<1% of the sorted cells were FoxP3+. Single cells were isolated on a FLUIDIGM C1 170 instrument, cDNA isolated, amplified, barcoded, and subjected to high throughput sequencing.
Following transcript pseudoalingment and normalization using Kallisto in AltAnalyze, predominant gene expression identified from individual cells by ICGS. ICGS provides the sensitivity to detect rare and common cell populations as well as possible transitional cell states. Four major cell populations were reported along with statistically enriched Gene
Ontology terms for gene clusters.
Acknowledgements:
This work was supported by funds from the National Institute of Health Grant AG033057 (to
D.A.H. and C.A.C.). We thank members of the Hildeman and Chougnet laboratories (Sarah
Jones, Cyd Castro Rojas, Rachel Walters, Sharmila Shanmuganad, Kun-Po Li, Courtney
Jackson) for their suggestions and help with mouse experiments. The authors would also like to thank Drs. Jeremy M. Kinder and Sing Sing Way for their help with the FoxP3-fate mapping mice and Calvin Chan for his help with adipose tissue IL-10 gene studies.
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Figures
Figure 1. Aged mice have increased systemic levels of IL-10. (A) Young (white bar, n=6) and old
(black bar, n=5) C57BL/6 mice were i.v. injected with biotinylated anti–IL-10 capturing Abs. Serum was collected 24 h later, and IL-10 levels were measured by ELISA. Graph shows the average serum IL-10
(mean±SEM). Data are representative of at least two independent experiments. (B) IL-10 mRNA gene expression was measured by real-time RT-PCR on cDNA isolated from the spleen, liver, gut, lymph nodes, (inguinal, epididymal) white and brown adipose tissue, from individual young (n=4-8) and old
(n=5-9) mice. Graph shows the mean fold change in IL-10 mRNA expression calculated by dividing the individual expression level in old mice by the average expression level of the young mice
(mean±SEM). Dashed line represents equal level of expression in young and old mice. Data pooled from two independent experiments. *p ≤0.05, **p ≤0.01, ***p ≤0.001, ****p ≤0.0001, Student’s t-test.
180 Figure 2. CD4+FoxP3- T cells are the major source of IL-10 in aged mice. (A) Splenocytes from young (n=3) and old (n=5) IL-10gfp (Vertex) mice were stained with Abs against CD4, CD8, TCRβ and
CD19 and analyzed by flow cytometry. The representative plots show the gating strategy and
181 frequencies of CD4+, CD8+, CD19+ and CD19- that are GFP+. Graphs show the total number (upper) and frequency (middle) of cells that are GFP+ in young (white bar) and old (black bar) mice
(mean±SEM). The lower graph shows the level of GFP expression in old CD4+, CD8+, CD19+ and
CD19- that are GFP+ (mean±SEM). **p ≤0.01, Student’s t-test. Data are representative of at least two independent experiments. (B) Splenocytes from young (3mo, n=4) and old (≥18mo, n=4) FoxP3-IRES-
DTR-GFP mice were enriched for CD4+ cells using CD4 T cell isolation kit (Miltenyi) and stained with
Abs against CD4, CD62L and CD44. CD4+FoxP3GFP+, CD4+FoxP3GFP− CD44hi CD62Llo (memory) and
CD4+FoxP3GFP− CD44lo CD62Lhi (naïve) cells were FACS sorted. Purified cells (3 × 105) were stimulated with PMA and Ionomycin (P+I) for 15hrs and IL-10 levels in culture supernatants quantified by ELISA. Graph shows the average IL-10 levels (mean±SEM). Data pooled from two independent experiments. (C) Splenocytes from young (n=4) and old (n=4) C57BL/6 mice were stimulated with
(P+I), stained with Abs against TCRβ, CD8, Foxp3 and IL-10, and assessed for cytokine production by flow cytometry. The representative plots show the gating strategy and frequencies of Foxp3+ or
Foxp3- that are IL-10+ from young or old mice. Graphs show the frequency and the total number of cells that are IL-10+ in young (white bar) and old (black bar) mice (mean±SEM). (D) Young (n=6) and old (n=14) C57BL/6 mice were treated with a single dose (600 μg) of anti-CD4 depleting Ab or isotype control (n=16) at d0. Old Foxp3-DTR C57BL/6 mice (n=6) were treated with a single dose (1.25μg) of
Diphtheria Toxin (DT). Mice were injected i.v. with biotinylated α-IL-10 capturing Ab at d1. Serum was collected at d2 to measure IL-10 levels by ELISA. Graph shows the average serum IL10 levels
(mean±SEM) and representative data pooled from two independent experiments. (E) Splenocytes from isotype control old mice (n=8) and old Foxp3DTR C57BL/6 mice treated with a single dose
(1.25μg) of Diphtheria Toxin (DT) (n=6) were stimulated as above (C), stained with Abs against TCRβ,
CD8, Foxp3 and IL-10 and analyzed by flow cytometry. Graph shows the frequency of Foxp3- cells that are IL-10+ (mean±SEM). *p ≤0.05, **p ≤0.01, ***p ≤0.001, Student’s t-test.
182 Figure 3. IL-10-producing FoxP3- CD4+ T cells in aged mice are predominantly Tfh cells. (A)
Splenocytes from young (n=6) and old (n=6) C57BL/6 mice were stimulated with (P+I), stained with
Abs against TCRβ, CD8, Foxp3, IL-10, IL-21 and analyzed by flow cytometry. The representative histograms and graphs show the frequencies and total numbers of IL-21+ cells within Foxp3- that are
IL-10+ (mean±SEM). *p ≤0.05, **p ≤ 0.01, Student’s t-test. Data are representative of at least two independent experiments. (B) Splenocytes from young (n=4) and old (n=4) C57BL/6 mice were stimulated as above and stained with Abs against TCRβ, CD8, Foxp3, CXCR5, PD-1 and IL-10, and analyzed by flow cytometry. The representative plots and graphs show the frequencies and total numbers of indicated subsets within Foxp3- that are IL-10+ (mean±SEM). *p ≤0.05, **p ≤ 0.01,
Student’s t-test. Data are representative of at least two independent experiments.
183 Figure 4. IL-6 is required for Tfh10 cells and for systemic levels of IL-10 in aged mice. (A, B)
Splenocytes from young and old C57BL/6 or IL-6-/- mice (n≥4/group) were stimulated with (P+I), stained with Ab against TCRß, CD8, CXCR5, PD1, Foxp3 and IL-10, and analyzed by flow cytometry.
184 The representative plots and bar graphs show the frequency and total number of Foxp3- that CXCR5+
PD1+ and their IL-10 production (mean±SEM). (C) Old C57BL/6 (n=8) and IL-6-/- (n=8) mice were i.v. injected with biotinylated anti–IL-10 Abs, serum was collected 24 hrs later, and IL-10 levels were measured by ELISA. Graph shows the average serum IL-10 (mean±SEM). *p ≤0.05, **p ≤0.01,
Student’s t-test. (D) Old C57BL/6 (19mo) mice were treated with isotype control (n=6) or α-IL-6 blocking antibody (n=8) on day0 and sacrificed on day2. Splenocytes were stimulated with (P+I), stained with Ab against TCRß, CD8, Foxp3 and IL-10 and analyzed with flow cytometry. The representative bar graph shows the frequency of Foxp3- that are IL-10+ (mean±SEM).
185 186 Figure 5. IL-21 contributes to accrual of Tfh10 cells and regulates the systemic IL-6/IL-10 balance. (A) Splenocytes from young and old C57BL/6 or IL-21-/- mice (n≥4/group) were stimulated with (P+I), stained with Ab against TCRß, CD8, Foxp3 and IL-21 and analyzed by flow cytometry. Bar graphs show the frequency and total number of Foxp3- that are IL-21+ (mean±SEM). (B, C)
Splenocytes from old C57BL/6 or IL-21-/- mice (n≥3/group) were stimulated as above, stained with Ab against TCRß, CD8, CXCR5, PD1, Foxp3 and IL-10, and analyzed with flow cytometry. The representative plots and bar graphs show the frequency and total number of Foxp3- CXCR5+ PD1+ and their IL-10 production (mean±SEM). Data are pooled from two independent experiments. (D, E)
Old C57BL/6 (n=4) and old IL-21-/- (n=3) mice were i.v. injected with biotinylated anti–IL-10 and anti-IL-
6 capturing Abs, serum was collected 24 h later, and IL-10 and IL-6 levels were measured by ELISA.
Graphs show the average serum IL-10 and IL-6 (mean±SEM)
187 Figure 6. IL-21 driven repression of Bim in aged Tfh10 cells results in their enhanced survival.
Splenocytes from young (n=4) and old (n=4) mice were stimulated with (P+I), stained with Abs against
TCRß, CD8, Foxp3, IL-10, Ki67, Bim and analyzed by flow cytometry. (A) Graph shows the frequency
188 of Foxp3- IL-10+ cells that are Ki67+( mean±SEM). (B) Graph shows the level of expression of Bim in
Foxp3- IL-10+ cells (mean±SEM). (C) Splenocytes from 6month old wild-type and Bim-/- mice
(n=6/group) were stimulated as above, stained with Ab against TCRß, CD4, Foxp3 and IL-10 and analyzed by flow cytometry. Plots and bar graphs show the frequency and total number of Foxp3- that are IL-10+ (mean±SEM). ). (D) Splenocytes from C57BL/6 or IL-21-/- mice (n≥3/group) were stimulated as above, stained with Ab against TCRß, CD8, Foxp3, IL-10 and Bim and analyzed with flow cytometry. Graph shows the level of expression of Bim in Foxp3- that are IL-10+ cells (mean±SEM). *p
≤0.05, **p ≤0.01, ***p ≤0.001, Student’s t-test. *p ≤0.05, **p ≤ 0.01, Student’s t-test.
189 Figure 7. Tfh10 cells in aged mice manifest diminished levels of BCL6 thereby enabling IL-10 expression. (A) Splenocytes from young and old C57BL/6 mice (n=4/group) were stimulated with
(P+I), stained with Ab against, CD8, CXCR5, PD1, Foxp3 and IL-10, and analyzed with flow cytometry.
The representative bar graphs show the level of expression of BCL6 and IL-10 in Foxp3- CXCR5+ PD-
1+ that are IL-10+ (mean±SEM). (B) Splenocytes from middle age wild-type or CD4Cre BCL6f/f mice
(n≥3/group) were stained with Ab against CXCR5, PD1and Foxp3. The representative plot and bar graph show the frequency of Foxp3- cells that are CXCR5+ PD1+ (mean±SEM). (C) Splenocytes from middle age wild-type or CD4Cre BCL6f/f mice (n≥3/group) were stimulated as above (A), stained with
Abs against TCRβ, CD8, Foxp3 and IL-10 and analyzed with flow cytometry. The representative histograms and bar graphs show the frequency and total number of Foxp3- cells that are IL-10+
(mean±SEM). *p ≤0.05, **p ≤0.01, Student’s t-test.
190 Figure 8. IL-10 limits Tfh-dependent vaccine responses in aged mice. (A) Young (n=6) and old
(n=5) mice were immunized with NP-KLH in Alum and sacrificed 20 days later. Splenocytes were stained with Abs against CD19, B220, GL7 and Fas and analyzed by flow cytometry. Representative
191 plots identifying GC B cells NP-specific as Fashi GL7hi that are IgG1+ NP tetramer+. Graphs show the frequency and the total number of splenic B cells that are IgG1+ NP+ (mean±SEM), as well as serum levels of immunoglobulin specific for NP (IgG1) of young vs old mice obtained 20 days after immunization (mean±SEM). (B) Mice were immunized as above d0 and then treated with isotype (n=7) or anti–IL-10R neutralizing Ab (n=8) on day -1, 1, 3, 6, 9 and sacrificed on day10. Representative plots display the frequency of GC B cells (NP-specific). Graphs show the frequency and the total number of
B cells that are IgG1+ NP+ (mean±SEM), as well as serum levels of immunoglobulin specific for NP
(IgG1) in mice with or without IL-10R neutralization obtained 10 days after immunization (mean±SEM).
*p ≤0.05, **p ≤ 0.01, Student’s t-test.
192 Figure 9. Tfh10 cells accumulate during aging in humans. Human spleen cells from young (n=8) and old (n=8) individuals were surface stained with Abs against CD3, CD4, CD45RO, CXCR5, PD-1 and Foxp3 and analyzed with flow cytometry. (A) Graph shows the frequency of CD3+ CD4+
CD45RO+ Foxp3- that are CXCR5+ PD-1+ (mean±SEM). (B, C) CD4+ T cells were bead-purified by negative selection, FACS-sorted memory CD4+ T cells (CD45RO+) into Tfh (CD25-CD127+PD-
1+CXCR5+), Treg (CD25+CD127-PD-1-CXCR5-) and other memory cells (CD25-CD127+PD-1-CXCR5-).
10,000 cells were stimulated in vitro with anti-CD3/CD28 beads at a 1:1 cell: bead ratio, or unstimulated. After 16h, supernatants were collected and analyzed by Luminex. Cytokines were undetectable in unstimulated cultures. Each individual is represented by a symbol, Y (young), O (old)
(mean±SEM). ***p ≤0.005, Student’s t-test.
193 *** 8000 Young *** O ld 6000
4000 IL-10 MFI IL-10 2000
0 FOXP3+ FOXP3-
Supplementary Fig.1. Splenocytes from young (n=4) and old (n=4) C57BL/6 mice were stimulated with (P+I), stained with Abs against TCRβ, CD8, Foxp3 and IL-10 and analyzed by flow cytometry.
Graph shows the mean level of IL-10 in Foxp3+ IL-10+ and Foxp3- IL-10+ cells (mean±SEM). ***p ≤
0.001, Student’s t-test.
Supplementary Fig.2. Splenocytes from microbiota-replete conventionally-housed (Con) (n=2-
4/group) or germ free (GF) (n=2-5/group) mice were stimulated with (P+I), stained with Abs against
TCRβ, CD8, Foxp3 and IL-10 and analyzed by flow cytometry. Graph shows frequency of Foxp3- cells that are IL-10+ (mean±SEM). Data pooled from 3 independent experiments. 194 Supplementary Fig.3. (A) Splenocytes from young (n=8) and old (n=8) mice were stimulated with
(P+I), stained with Abs against TCRβ, CD8, CD49B, LAG3 and IL-10, and analyzed by flow cytometry.
The plots and graph show the frequency of indicated subsets in Foxp3- IL-10+ cells (mean±SEM). (B)
Splenocytes from young (n =4) and old (n =4) mice were stimulated as above, stained with Abs 195 against TCRβ, CD8, IL-10, IL-17, IL-4 and analyzed by flow cytometry. The plots and graph show the frequency of indicated subsets in Foxp3- cells (mean±SEM). (C) Spleen cells from young (2mo, n=5) and middle-age (12mo, n=5) IL-17Acre Rosa26 YFP (R26YFP) mice were stimulated as above, stained with Abs against TCRβ, CD8, IL-10, IL-17 and analyzed with flow cytometry. Bar graph shows the
(mean±SEM), within the total FOXP3- IL-10+ cells, the frequency of cells producing IL-17 (IL-17+YFP+, gray), exTh17 (IL-17-YFP+, white) or those that never produced IL- 17 (IL-17-YFP-, black). (D) Spleen cells from middle-age (12mo, n=4) and young (6weeks, n=3) Foxp3CreRosa26dTomato mice were stimulated with (P+I), stained with Abs against TCRβ, CD8, IL-10 and Foxp3, and analyzed with flow cytometry. Plots and bar graph show, within the total Foxp3- IL-10+ cells, the frequency of exTreg cells
(Foxp3- IL-10+ dTomato+). Data pooled from two independent experiments (mean±SEM).
196 Supplementary Fig.4. Splenocytes from young, middle age and old C57BL/6 mice (n≥4/group) were stimulated with (P+I) and stained with Ab against TCRß, CD8, CXCR5, PD1, Foxp3 and IL-10. The representative bar graphs show the frequency of Foxp3- that CXCR5+ PD1+ and those that produce
IL-10 (mean±SEM). *p ≤0.05, **p ≤0.01, Student’s t-test.
197 Supplementary Fig.5. Old C57BL/6 mice were treated with isotype control (n=6) or α-ICOSL blocking antibody (n=5) on day (0, 3, 6, 9) and sacrificed on day 12. Splenocytes were stimulated with (P+I), stained with Ab against TCRß, CD4, CD8, CXCR5, PD1, Foxp3 and IL-10, and analyzed by flow cytometry. The representative plots and bar graphs show the frequency of Foxp3- that CXCR5+ PD1+ and the frequency of Foxp3- that IL-10+ (mean±SEM). 198 1200
900
600
300
IL-10+ BLIMP-1 MFIIL-10+ 0 WT BCL6 cKO
Supplementary Fig.6. Splenocytes from middle age wild-type or CD4Cre Bcl-6f/f mice (n≥3/group) were stimulated with P+I and stained with Abs against TCRβ, CD8, Foxp3 and IL-10 and Blimp-1 and analyzed by flow cytometry. Graph shows the mean fluorescence intensity (MFI) for Blimp-1 in IL-10+
Foxp3- cells (mean±SEM).
199 Chapter 5
Summary
The world is experiencing an extraordinary increase in elderly population. The average life expectancy will reach 84 years old by 2050 compared to 68 years old in 1950 (1).
Such an increase in longevity will have a huge impact on health care system. A significant part of this burden on health care is the age-driven loss of immune competence (2-6). Indeed, aging is associated with quantitative and qualitative defects in T and B cells resulting in adverse health outcomes and an increase in the susceptibility to acquire infections and a failure to respond to vaccines (7-10).
Another feature of the dysregulated immune responses with age is the emergence of chronic low-level inflammation termed inflammaging (11-13). Inflammaging is thought to be a major driver for serious age-related diseases, such as atherosclerosis, cardiovascular diseases, Alzheimer's disease, rheumatoid arthritis, cancer, and frailty
(11, 12). One major focus in the field of aging is to address possible compensatory regulatory mechanisms that can combat this inflammation with age favoring reduction in age related pathologies and improving the quality of life for elderly.
Human CMV (HCMV) is a ubiquitous infection, infecting 40–90% of world’s population
(14). HCMV is never cleared and persists in human hosts for life (15). Such persistence is thought to accelerate age-related immune dysregulation and inflammaging with a detrimental effect on host immunity (16). Understanding the interplay between aging, latent CMV and the mechanisms regulating its persistence and reactivation might
200 provide novel interventions to control the adverse effects of age-related CMV infection.
In chapter 3 we aimed at dissecting the regulatory mechanisms that shaped the immune
responses towards latent CMV. Surprisingly, we found that the relationship between
CD4+ Foxp3- IL-10 producing T cells and regulatory Foxp3+ T (Treg) cells was more
complex and intertwined than previously appreciated. Additionally, the role of Treg and
Foxp3- IL-10 producing cells was highly integrated resulting in differential control of
CMV reactivation/replication in the spleen and salivary glands. A brief summary of our
findings from chapter 3 and potential future directions for this work will be addressed in
the next section.
Inflammaging is characterized by an increase in plasma levels of pro-inflammatory
cytokines such as Interleukin-6 (IL-6) (17), which is a major predictor of mortality and
morbidity with age (18). Given our findings with IL-10 and control of latent MCMV
reactivation/replication, we reasoned that the production of anti-inflammatory cytokines could be a potential mechanism to counterbalance such inflammaging. Indeed, we observed that increased production of IL-6 was counterbalanced by a concomitant
production of large quantities of IL-10 with age. To further understand the role of IL-10
with age, we investigated its cellular sources, the molecular mechanisms driving its
production and its role in regulating immune responses with age. A summary of our
findings from chapter 4 and future directions will be addressed in the next section.
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Major findings and future directions
Chapter 3
1- Foxp3+ Treg cells play differential roles in the spleen and salivary glands during latent MCMV infection.
Following primary infection with MCMV, the adaptive immune response clears most replicating virus, however the virus establishes latency within multiple tissues (19). CD8
T cells are critical for controlling both acute and latent MCMV infection (20). Indeed,
persistent MCMV infection in the spleen is known to induce the formation of activated, highly differentiated memory CD8 T cells (21). However, the regulatory mechanisms controlling CD8 T cells memory responses to MCMV and their potential contribution to controlling the latent viral pool are not clear. Our data provided evidence that, CD8 T cells are negatively regulated by Foxp3+ Treg cells. Depletion of Treg cells resulted in enhanced effector CD8 T cell responses in the spleen and a reduction in latent viral pool. The data clearly highlight an unappreciated in vivo role for regulatory T cells during latent MCMV.
Interestingly, while Treg depletion resulted in reduced latent viral pool in the spleen, it enhanced viral reactivation/replication in the salivary gland (SG). The SG represents a major site of cytomegalovirus replication and transmission (22). Furthermore, the production of IL-10 from Foxp3- CD4+ T cells attenuated antiviral responses and promoted viral replication in the SG (23). Interestingly, the reactivation of MCMV in the
SG was accompanied by the emergence of IL-10+ Foxp3- T cells. The data implicates
IL-10 from Foxp3- cells in viral replication/reactivation.
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Notably, TNF-α, is an important cytokine in the control of MCMV reactivation/replication
(24, 25). Indeed, we observed significant increases in the expression of TNF-α in the
SG in the absence of Treg cells. Thus, our data is consistent with a scenario in which
IL-10+ Foxp3- cells and their production of IL-10 might enhance viral replication following TNF-α mediated MCMV reactivation.
While we saw an increase in viral reactivation/replication in the SG we observed reduced viral load in the spleen. One question that arose from our results is why the viral load in the spleen was reduced despite the elevated levels of IL-10-producing
CD4+ T cells in that organ? A possible explanation for this scenario is that upon Treg depletion expansion of MCMV-specific cytotoxic CD8 T cells could detect and kill cells harboring this reactivating virus. In contrast to the spleen, CD8 T cells are incapable of reducing virus in the SG due to loss of expression of MHC class I molecules in virally- infected cells in that organ.
These data also raise the question regarding the mechanism by which regulatory T cells are controlling CD8 T cells in the spleen. One possibility is that Treg might utilize TGF-ß,
IL-35 or CTLA-4 to suppress CD8 T cells (26). One way to test this possibility is through the individual neutralization of potential regulatory factors (Table1) in groups of MCMV latently infected mice. One group of control mice would undergo Treg depletion, while another control group would be injected with an isotype control antibody. The spleen
PCR for viral load and an explant assay would be utilized to examine the reactivation of the virus and flow cytometry will be utilized to investigate the levels of antigen- specific
CD8 T cells and their effector responses (cytokine production and cytotoxicity).
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Table 1
Regulatory molecule Neutralizing Ab
IL-35 anti-IL-35
TGF-ß anti-TGF-ß
CTLA-4 CTLA-4Ig
IL-35/CTLA-4/TGF-ß anti-IL-35/ anti-TGF-ß/ CTLA-4Ig
It is possible that we would not see an effect of the neutralization of the regulatory factors separately relative to Treg depleted mice. For example, Treg depleted mice would experience enhanced CD8 T cell responses and reduction in viral load in the spleen. However, infected mice with neutralization of (e.g. IL-35) would show no effect on CD8 T cells and viral load. This result would suggest possible redundancy in the regulatory mechanisms utilized by Treg cells in controlling CD8+ T cell responses. Such possibility would require testing by combined neutralization of all three potential mechanisms (Table 1) which might be the best way to recapitulate the in vivo effect that we see with total depletion of Treg cells. One caveat that might arise from the experiment in the spleen is the maintenance of the neutralization effect ex vivo in the spleen explants. However, we expect that if single neutralization of one regulatory marker or all of them was efficient to enhance the CD8 T cells responses in vivo then spleen PCR would show significant reduction in viral load between neutralized mice compared to the control group which would reflect reduction in latent viral pool and hence delayed viral reactivation in spleen explants as well.
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2- IL-10 is critical for MCMV replication in the salivary gland (SG).
Neutralization of IL-10R resulted in reduction in reactivation/replication of virus in the
SG after depletion of Treg. However, we did not distinguish whether the effects of IL-
10R blockade were to limit outright viral reactivation or to limit viral replication.
Nonetheless, our data are consistent with IL-10 promoting viral replication more than outright viral reactivation as three key pieces of data suggest that IL-10 is not involved in direct viral reactivation. First, IL-10 has been shown to regulate viral replication during primary infection rather than reactivation (27). Second, the neutralization of IL-10R was done two days after Treg depletion. Thus, it is possible that reactivation of the virus has already been established before the treatment. Lastly, we attempted to induce viral reactivation in our in vitro explant assay and found that the addition of IL-10 to the assay was not sufficient to exacerbate viral reactivation. While we cannot conclusively rule out the role of IL-10 in viral reactivation, the data suggests the presence of other stimuli driving viral reactivation in the absence of Treg cells.
Interestingly, we found an increase in TNF-α after Treg depletion in the SG. Notably, prior in vitro studies have shown a pivotal role for inflammatory cytokines, specifically of
TNF-α in the reactivation of latent MCMV via the activation of MIE enhancer (24, 28).
Thus, one possibility is that TNF-α can induce the first stage of reactivation followed by
IL-10 enhancing the replication. We hypothesize that TNF-α is controlling viral reactivation in the SG upon Treg depletion. Interestingly, our preliminary data neutralizing TNF-α showed a significant reduction in SG viral titers (Figure1), consistent with the concept that TNF-α is controlling viral reactivation. The results from TNF-α neutralization and IL-10R neutralization experiments support a scenario where Treg
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depletion would lead to the production TNF-α, the activation of MIE enhancer and
subsequently viral reactivation. This would constitute the first checkpoint, while
simultaneous IL-10 production upon Treg depletion provides a second checkpoint
resulting in further enhancement of viral replication. Notably, the neutralization of TNF-α
in latent MCMV infected mice was done simultaneously with Treg depletion, while IL-
10R neutralization was done after two days of Treg depletion. This might also support
our proposed scenario that TNF-α controlled the reactivation of the virus which is happening very early once Treg are depleted while IL-10 controlled the replication after two days of reactivation.
We propose to investigate the role of TNF-α in MCMV reactivation in the SG through a series of experiments, some of which involve the tissue explant reactivation assay.
Notably, the explant assay is a functional assay to assess reactivation from latency. If the virus is typically replicating in the tissue at the time of isolation, it will be detected by seven days post explants. However, if virus is latent in the tissue at the time of isolation, it will take longer and usually will reactivate after day7. We anticipate that SG tissue
cultured with TNF-α will exacerbate the reactivation and therefore result in increased
viral production relative to culture in the absence of TNF-α. Such results would provide
strong evidence that TNF-α is controlling reactivation rather than replication of the virus.
However, if we see no evidence for differences in the reactivation between the cultures
with and without TNF-α, this would suggest that either TNF-α is not involved in
reactivation per se and is either redundant with other factors controlling reactivation or
TNF-α contributes to viral replication in the SG upon Treg depletion.
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Major findings and future directions
Chapter4
1- A novel population of Tfh cells accumulates with age and is the dominant
source of IL-10 in aged mice.
In chapter 4 we describe a novel population of T follicular helper cells that accumulated
in aged mice and produced IL-10 (Tfh10). In agreement with previous reports
supporting the role of IL-21 and IL-6 in Tfh differentiation (29, 30), we found that both IL-
6 and IL-21 were required for the accumulation of Tfh10 cells with age. Notably, we did
not observe any change in Tfh10 cells following IL-6 neutralization which suggested that
IL-21 is required for their maintenance rather than IL-6. Indeed, previous reports have
shown that IL-6 can be the first inducer for IL-21 production from Tfh cells and IL-21 is
supporting their maintenance (31) (32). Our data also suggested that IL-21 regulated
the survival of Tfh10 cells via down regulation of Bim. However, the detailed molecular
mechanism(s) underlying the survival effect under IL-21/Bim axis were still not clear.
Mechanistically, IL-21 signaling may promote Tfh survival through activation of the
PI3K/Akt/Foxo3a axis which plays a major role in negatively regulating Bim expression
(33, 34).
Indeed, a previous study showed that the direct binding of IL-21 to its receptor on T cells leads to the activation of the PI3K signaling pathway and increased cell survival
(35). However, the authors attributed the IL-21 driven survival to the up-regulation of the survival molecule BCL-2 and the effect of IL-21/PI3K axis on Bim levels was not
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addressed. To investigate the role of IL-21/Bim axis in the maintenance and survival of
Tfh10 cells we would propose several experiments.
To assess whether IL-21 is critical for the maintenance/survival of Tfh10 cells, we would
block IL-21R signaling in vivo using an IL-21R neutralizing antibody. We expect to see
lower levels of Tfh10 cells in aged IL-21R neutralized compared to isotype mice. We
would also expect to see higher levels of Bim in the remaining Tfh10 cells upon IL-21R
neutralization. The data would suggest that IL-21 is required to control levels of Bim in
Tfh10 cells and their homeostasis. Second, we would breed Bim-/- to IL-21-/- mice and assess their phenotype. We expect to see restoration in Tfh10 cells compared to their loss in Bim+/+ IL-21-/- mice. The results would further support the role of Bim in IL-21-
driven Tfh10 survival.
Third, to test the role of IL-21/PI3K/Bim in the survival of Tfh10 cells we would sort old
Tfh10 cells (CXCR5+ PD1+ Foxp3 RFP- IL-10 GFP+) from Foxp3-RFP IL-10-GFP reporter mice and culture the cells with IL-21. Expression of Bim with and without stimulation would be analyzed with flow cytometry. We expect to see lower levels of Bim after culture with IL-21. To further test the role of PI3K on IL-21 driven suppression of
Bim, we would use PI3K inhibitor. In this experiment we expect that PI3K inhibition would abrogate the down-regulatory effect of IL-21 on Bim levels, which would implicate
IL-21-driven PI3K on Bim expression and potentially in the survival of Tfh10 cells with age. However, no change in Bim levels under IL-21 stimulatory conditions might suggest an indirect effect of IL-21 on Bim in Tfh10 cells. In other words, IL-21 might need to indirectly signal to other cells that control the survival of Tfh10 cells and their
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Bim levels. This scenario would require targeted deletion of IL-21R on different cells
such as B cells, T cells and dendritic cells (DCs).
2- IL-10 is increased with age and counterbalances inflammaging.
Inflammaging is characterized by a large increase in the production of IL-6 which is a
trigger for serious inflammatory diseases and frailty with age (17). Several critical questions have arisen in the field of aging regarding the compensatory mechanisms that would enable the human body to cope with and regulate the persistent production of IL-
6 with age. Indeed, experimental evidence through utilizing IL-10-/- mice (IL-10 deficient mice in pathogen free environment) indicated that IL-10 is able to counteract the appearance of major physical and biological characteristics of human frailty related to
IL-6 such as sarcopenia, muscular weakness, weight loss as well as age-related
increase of serum IL-6 (36). Thus, IL-10 might represent a key to counterbalance IL-6
inflammation with age. However, the role of IL-10 in regulating inflammaging and its
cellular sources are still unclear.
In chapter 4, we reported that IL-10 expression from different tissues implicated
secondary lymphoid organs (spleen, peripheral lymph nodes) as major site for
increased IL-10 production with age. One scenario that we considered for this enhanced
IL-10 production initially was the accrual of aged Treg cells which are known to produce
IL-10. Although IL-10 production from Treg cells was increased with age, the production
of IL-10 was far greater in Foxp3- CD4+ T cells in aged mice. More importantly,
depletion studies provided strong evidence that Foxp3- cells were the predominant
source of systemic IL-10 in aged mice.
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Interestingly, similar to our data with IL-10, we found an increase in IL-6 expression in
the spleen and lymph nodes of old mice. More importantly, when IL-10 was decreased
in the absence of IL-21, IL-6 levels were increased with age. The findings suggest that
the increase in IL-10 production from Tfh10 cells with age is a counter-regulatory
mechanism for the increased IL-6 production with age. However, this needs to be
tested.
We propose to test the role of IL-10 in counterbalancing IL-6 with age through series of
experiments. First, we will use IL-21R neutralizing antibody in aged mice as mentioned
earlier and we will measure serum IL-10 and IL-6 levels via IVCCA. We expect an
increase in systemic IL-6 and decrease in IL-10 levels upon IL-21R neutralization similar
to what we see in aged IL-21-/- mice. We will follow these experiments with additional
transfer experiments. For example, we will transfer old Tfh10 cells into a group of aged
IL-21 deficient mice. We expect that the transfer of aged Tfh10 cells would restore the homeostatic IL-6/IL-10 balance. Such findings would support the role of IL-10 in counterbalancing IL-6 with age. However, absence of expected results would suggest that IL-21 is directly controlling IL-6/IL-10 balance with age.
3- BCL6 limits IL-10 production with age.
Our study highlights a major role for BCL6 in controlling IL-10 production with age.
Indeed, we found that BCL6 is naturally downregulated in aged Tfh10 cells and the
specific loss of BCL6 in T cells resulted in a significant increase in IL-10 production from
Foxp3- CD4+ T cells, with the caveat that these IL-10+ cells no longer expressed
markers of Tfh cells. Nonetheless, these data strongly implicate BCL6 as a physiologic
210 suppressor of IL-10. We envision a few mechanisms for how BCL6 may promote IL-10 expression in Tfh10 cells. 1- Through the lack of direct repression on IL-10 due to reduction of BCL6 expression. 2- Loss of repression due to other transcription factors that promote IL-10 expression.
To examine the first scenario that BCL6 expression levels are regulating IL-10 expression in aged Tfh cells we will restore expression of BCL6 in aged Tfh cells and assess its effects on IL-10 production. Aged Tfh10 cells from Foxp3-RFP IL-10-GFP double reporter mice will be transducer with a BCL6 retrovirus or an empty vector as a control. The cells will be stimulated subsequently to examine the production of IL-10 under PMA and Ionomycin. We anticipate that over expression of BCL6 in aged Tfh10 cells would result in decreased production of IL-10 compared to control cells transduced with empty vector. Such results would provide evidence that the lower expression of
BCL6 in aged Tfh10 cells is regulating their IL-10 production.
These results would be followed with more experiments to determine the mechanisms that could be regulating the reduction in BCL6 levels in Tfh10 cells with age. Notably, both IL-6 and IL-21 are inducers of STAT3 activation, while IL-21 also induces STAT5.
More importantly, a previous study, provided evidence for STAT5 outcompeting STAT3 for binding at the BCL6 locus and directly repressing BCL6 expression in the absence of
STAT3 (37, 38). Thus, we propose that dysregulated signaling of STAT5/STAT3 in
Tfh10 cells under the effect of IL-21 and IL-6 with age would result in reduced BCL6 levels and enhanced IL-10 production. Mechanistically, expression of SOCS molecules
(e.g. SOCS3, SOCS1) following STAT3 activation in aged Tfh10 cells can result in a
211 negative feedback loop and reduction in STAT3 phosphorylation via interfering with
JAK/STAT signaling pathway in the presence of gp130 chain of IL-6R (39).
We will examine the levels of p-STAT3 and p-STAT5 in a group of young (3 month) and old (17 month) mice (Foxp3-RFP IL-10-GFP double reporter mice) to sort on CXCR5+
PD1+ RFP- GFP+ (Tfh10) cells. Sorted cells will be cultured under IL-21/IL-6 stimulatory conditions and the level of p-STAT3, p-STAT5 will be analyzed with flow cytometry. We expect that in response to IL-21/IL-6, aged Tfh10 cells would have reduced levels of p-STAT3 and higher levels of p-STAT5 compared to young counterparts. Further, we expect that this stimulation with IL-21/IL-6 will lead to reduced expression of BCL6. Together, these data would suggest that a shift in p-STAT5/p-
STAT3 under IL-21/IL-6 is regulating the reduced levels of BCL6 in Tfh10 cells with age.
This could be further tested using STAT5 inhibitor which would result in restoration of levels of BCL6 in aged Tfh10 relative to young Tfh10 cells under IL-21/IL-6 conditions.
It is possible that we would not observe a change in p-STAT3/p-STAT5 under stimulation but reduction in BCL6 levels. This result might implicate dyregulation in p-
STAT1 in aged Tfh10 cells. Notably, STAT1 can also regulate BCL6 induction downstream of IL-6R (29). Therefore, a reduction in p-STAT1 levels under IL-6 stimulatory conditions can result in the reduced expression of BCL6 in Tfh10 cells with age. Further, multiple signaling pathways contribute to BCL6 induction in Tfh cells. For example, the transcription factor BATF has been shown to positively regulate the expression of BCL6 in Tfh cells (40), while FOXO1, which is regulated by PI3K pathway, may negatively regulate BCL6 expression (41). Should we fail to observe involvement of
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altered STAT5 signaling on BCL6 expression, we would investigate the roles of these
other pathways that may contribute to age-related loss of BCL6 expression.
It is also possible that the decrease in BCL6 may not contribute significantly to the
increased production of IL-10 in aged Tfh cells. In this case, another possible scenario
regulating IL-10 expression in aged Tfh10 cells is an increase in other transcription
factors that positively regulate IL-10 expression. Indeed, we do observe an increase in
c-Maf in Tfh10 cells with age (Figure 2, A). c-Maf is involved in the differentiation of Tfh
cells as well as IL-10 production from CD4 T cells (42, 43). Interestingly, we observed a
significant reduction in c-Maf expression in aged Tfh10 cells in the absence of IL-
21(Figure2, B). Hence, c-Maf might play a role in the production of IL-10 from Tfh cells
with age. To assess the role of c-Maf in Tfh10 with age, we propose to examine the
levels of Tfh10 cells and their production of IL-10 in old mice through inducible deletion
of c-Maf in CD4 T cells using CD4ERCre+BCL6 f /f mice. Group of old CD4ERCre+BCL6 f /f and CD4ERCre-BCL6 f /f control mice will be treated with tamoxifen to induce c-Maf deletion. Expression of c-Maf in CD4 T cells will be first assessed by flow cytometry to assess efficiency of deletion. We will also measure the level of Tfh10 cells and their IL-
10 expression via flow cytometry. We anticipate that the loss of c-Maf in CD4 T cells to
result in reduction in Tfh cells with age and there production of IL-10 compared to
control group. To further investigate the intrinsic role of c-Maf in the expression of IL-10
in Tfh10 cells, we propose to knock down c-Maf in aged Tfh10 and determine the level
of expression of IL-10 in Tfh10 cells. We anticipate that c-Maf knock down would result
in the loss of IL-10 expression from aged Tfh cells compared to control group. The data
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would strongly support a positive intrinsic regulatory role for c-Maf in IL-10 expression in
Tfh10 cells with age.
4- IL-10 limits Tfh-dependent vaccine responses in aged mice.
Age-related increases in IL-10 can result in unwanted side effects, such as reducing
vaccine responses. Indeed, previous studies in humans have shown that lower
antibody titer to influenza A virus correlated with increased IL-10 production (44), and
levels of IFN-γ relative to IL-10 correlated with vaccine protection with age (45). A
previous study by Lefebvre et al, reported an increase in antigen specific IL-10
producing CD4 T cells which correlated with loss of vaccine responses in aged mice
(46). Although the authors attributed this increase in IL-10 production to the
accumulation of Foxp3+ Tfr cells, it was also possible that IL-10 from Tfh10 cells might
be hampering the vaccine responses in aged mice. Hence, the increase in systemic IL-
10 and the accrual of Tfh10 might hamper vaccination responses with age.
Based on these data, we hypothesized that inhibiting IL-10 would serve to enhance humoral and germinal center B cell responses with age. Our data with IL-10R
neutralization provided evidence for a negative regulation imposed by IL-10 on antigen
specific B cell responses with age. Our findings have important implications for the
development of therapeutic strategies to augment vaccination responses with age via
targeting IL-10 or Tfh10 cells. Indeed, although there are T and B cell intrinsic defects
with age, our data with IL-10R neutralization suggest that these defects are not
permanent and are amenable to manipulation.
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Notably, our findings can be further bolstered by additional experiments that would
enable us to identify the antigen specific Tfh and Tfh10 cells and their localization during
infection relative to B cells in the germinal center. Moreover, the effect of the
neutralization of IL-10 was very promising, yet it is unclear if such effect is sufficient to
provide future protection from infection. Thus, we propose to use a Tfh dependent
vaccination model using A/PR/8/34 influenza nucleoprotein (NP) which would enable us
to track antigen specific Tfh and B cells similar to a previous study (47). Mice will be
vaccinated with recombinant NP in alum and then challenged. We will utilize four groups
of young and old Foxp-3 RFP IL-10 GFP reporter mice (Table2), which would allow us
to specifically identify via flow cytometry three populations of interest that are antigen
specific, Tfh10 (CXCR5+ PD1+ RFP- GFP+), Tfh (CXCR5+ PD1+ RFP- GFP-) and Tfr
(CXCR5+ PD1+ RFP+ GFP-). NP-specific Tfh, Tfh10 and Tfr will be detected with I-Ab –
NP311-325 Tetramer and NP-specific GC B cells will be detected by staining with PE
labeled B cell tetramer in lungs and spleen.
Table 2 Treatment NP vaccine Influenza challenge PROTECTION Young iso + Young α-IL-10R + Old iso - Old α-IL-10R + iso+ α-IL-10R d -1, 3, 6, 8, 10, Vaccine d 0, Influenza infection d21 All mice will be vaccinated with recombinant NP in alum d0 and then infected with PR8 influenza d21 post vaccination. Similar to our previous model, young and old mice will be injected with anti-IL-10R neutralizing antibody at Day-1, 3, 6, 8 and 10 or isotype control. A group of young and old mice will be used as control unvaccinated mice.
Vaccinated mice will be infected intranasally with 400 EID50 of PR8 influenza and
215 sacrificed at day 5 post infection. Second, we will monitor the weight and survival of mice throughout the experiment. Third, we will determine the level of antibody titers via
ELISA. Finally, we will utilize flow cytometry as mentioned above.
Based on our previous results, we expect that the vaccinated young mice would have the highest protection (increased survival, weight and antibody titers) compared to vaccinated aged mice. We expect the vaccinated aged mice that received anti-IL-10R neutralizing antibody to have almost similar level of protection compared to young mice and higher levels compared to aged isotype control mice. If the vaccinated aged mice are not protected that might be due to the quality of neutralizing antibody. In other words, aged mice might have acquired similar levels of antigen specific antibodies compared to the young mice with anti-IL-10R, but they are still not potent enough to provide protection.
Notably, although an NP vaccine induces strong CD8 T cell protective immunity, the protein is weak at inducing neutralizing antibodies. Additionally, previous work has shown differential expression of antibody isotypes upon IL-10R neutralization (48).
Therefore, it is possible that the effect for blocking of IL-10 signaling is selective, increasing specific isotypes which might be involved in protection. Indeed, a previous study has shown that anti-IL-10 treatment of young and aged mice caused a significant increase in influenza-specific IgM and IgG1 titers and a decrease in IgG2b and IgG3.
The study did not examine the role of these isotypes in the protection against infection which would have been very informative. Although the differential effect of IL-10R neutralization on antibody isotypes in our experiment might depend on model of vaccination this possibility should be taken into consideration.
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Targeting IL-10 with age: Clinical implications
Several approaches have been used to improve vaccine efficacy for the elderly. For
example, TLR4 agonist as a vaccine adjuvant improved the cell-mediated immune
response with age through the stimulation of inflammatory cytokines (IL-6 and TNFα) in
myeloid DCs which was associated with reduction in IL-10 levels (49). Moreover,
another TLR4 agonist (GLA-SE), in combination with the influenza split-virus vaccine
increased significantly the ratio of IFN-γ:IL-10 produced by the aged PBMCs to levels similar to those obtained in young cells (50). Thus, our results in mice strongly suggest that interfering with IL-10 could have an adjuvant-like effect on bolstering immune responses in aged people. However, given that IL-10 deficiency in mice and humans results in significant autoimmunity, there is a concern that targeting IL-10 might results in unwanted side effects, especially in the intestine or a flare of an autoimmune disease which is a major concern. Therefore, safety of IL-10 inhibition or IL-10R blockade is important.
Fortunately, a recent study, demonstrated that blocking of IL-10 signaling at the time of immunization does not induce unwanted side effects in the intestine or result in increased risk of the development of autoimmune disease in mice (51). Indeed, results using human papillomavirus (HPV) immunization model with neutralization of IL-10 receptor during a period of two weeks indicated that the neutralization of IL-10R does
not increase the development of autoimmune disease in NOD mice or the infiltration of inflammatory cells into the intestine using inducible model of colitis, suggesting that blocking of IL-10R is relatively safe and well tolerated.
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Notably, the effect of IL-10 neutralization on vaccination responses might depend on the different roles of antibody isotypes in antiviral immunity. Indeed, previous studies provided evidence for distinct contributions of vaccine-induced IgG1 and IgG2a antibodies to protective immunity against influenza. Stimulation of IgG2a antibodies has been associated with increased efficacy of influenza vaccination and IgG2a antibodies are more efficient at clearing the virus compared to IgG1 isotype (52). Indeed, Fc portion of IgG2a antibodies are more efficient in interacting with complement components and Fc receptors than IgG1. This interaction strongly activates Fc receptor- mediated effector functions such as antibody-dependent cell-mediated cytotoxicity and opsonophagocytosis by macrophages (53, 54). Thus, utilization of IL-10R neutralization strategy in vaccination with age might depend on its effect on the different isotypes involved in control of the virus and protection. However, the effect of IL-10 or IL-10R neutralization might largely depend on the nature of the immune response. Hence, knowledge concerning the isotypes that are most efficient in the clearance of the virus is imperative.
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Integrated model for the regulators with age
In this dissertation we focused on understanding the molecular mechanisms that
underlie the accumulation of Tfh10 cells in aged mice. We propose a model (Figure 3)
for the homeostasis of Tfh10 with age where IL-6 is required for the initial development
of Tfh10 cells and the subsequent production of IL-21 is maintaining and regulating the
survival of Tfh10 cells by repressing Bim expression with age. More importantly, our
findings summarize the recent advance in our understanding of the crosstalk between
pro and anti-inflammatory factors with age and propose novel therapeutic strategies that
can be designed to specifically bolster immune responses with age. Indeed, work from
this dissertation provided compelling evidence for a unique interaction between aging,
CMV infection, Treg and Tfh10 cells with age. Our findings support the presence of a
strong interconnected network between the immune regulators with age. Disrupting this network can result in unforeseen consequences. Thus, therapeutic intervention on the basis of Treg and IL-10 manipulation with age should be approached with extreme caution. A considerable risk resulting from Treg manipulation in CMV infected individual is viral reactivation in the salivary glands and an increase in Tfh10 cells and production of IL-10 in the spleen thus hampering vaccination responses (Figure 4).
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1.5 * * 11/13 1.0 2/12 3/14 3/15 0/4
0.5
0.0
Virus Titer (Log10 pfu/ml) (Log10 Titer Virus UI
D T+ ISO D T+ ISO
DT+ Anti-TNF-a DT+ Anti-TNF-a WT DTR
Figure1. 5–6 week old WT C57BL/6 and Foxp3-DTR mice were inoculated with 1× 106 plaque-forming unit (pfu, p.i.) of MCMV. 8-9 months post-MCMV infection, both groups were injected with DT on day 0, 3, 6 and sacrificed on day 7. Groups were split and were treated with 500μg of either isotype control antibody or with anti–TNF-α neutralizing antibody on day 0, 2, 5 and were sacrificed on day7. Graph shows the average viral titer of individual salivary glands (SGs) of MCMV infected mice day7 post
Treg depletion and numbers on graph indicates the number of mice positive for virus in the SG. The presence of replicating virus was detected by plaque assay. Samples with no detectable virus were assigned a titer of 0.7 log pfu/ml, the limit of detection for the plaque assay as indicated by the dashed line. Statistical analysis, *p ≤ 0.05, **p ≤ 0.01
(Mann-Whitney test)
220
Figure 2. Splenocytes from young (n=4) and old (n=4) mice were stimulated with PMA and Ionomycin. Cells were stained for TCRß, CD8, CXCR5, PD1, Foxp3, IL-10 and c-
Maf, and analyzed by flow cytometry. (A) Graph shows the level of expression of c-Maf in CXCR5+ PD1+ that are IL-10+ cells (mean±SEM). (B) Splenocytes from old WT and
IL-21-/- were stimulated and stained as above. Graph shows the level of expression of c-
Maf in CXCR5+ PD1+ that are IL-10+ cells (mean±SEM). *p ≤0.05, ***p ≤ 0.001,
Student’s t-test.
221 Figure 3. Molecular events leading to the generation and maintenance of Tfh10 cells with age. In vivo production of IL-6 induces the differentiation of Tfh10 cells and their production of IL-21 and IL-10. Subsequent expression of IL-21 by differentiated Tfh10 cells enables them to maintain their phenotype in the absence of IL-6 in a self- sustainable manner. IL-21 produced by aged Tfh10 cells is required to regulate their survival by negatively regulating the expression of pro-apoptotic molecule Bim independent of conversion and proliferation mechanisms.
222
Figure 4. Diagram describing implications for targeting Treg Foxp3+ T cells in elderly
CMV-seropositive individual. Targeting Treg in an elderly individual can result in reactivation/replication of the virus due to the effect of IL-10 production from Foxp3- IL-
10 producing cells. In the spleen, targeting Treg can result in increased levels of Tfh10 cells. Enhanced IL-10 production from Tfh10 cells in the spleen would have negative impact on B and Tfh cells in the germinal centers resulting in reduced efficacy to vaccination.
223
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