characterization of a novel dendritic cell population · 2013-10-17 · 1 chapter 1 introduction...
TRANSCRIPT
Characterization of a Novel Dendritic Cell Population
by
Anastassia Mikhailova
A thesis submitted in conformity with the requirements for the degree of Master of Science
Institute of Medical Science University of Toronto
© Copyright by Anastassia Mikhailova 2012
ii
Characterization of a Novel Dendritic Cell Poluation
Anastassia Mikhailova
Master of Science
Institute of Medical Science University of Toronto
2012
Abstract
Conventional DC (cDC) arise from circulating immediate precursors (pre-cDC), and are
currently thought to be terminally differentiated. Here we show that cDC are capable of
generating progeny that lost all characteristic features of cDC and aquired regulatory properties.
Sorted bone marrow pre-cDCs were cultured on a stromal monolayer in the presence or absence
of granulocyte-macrophage colony stimulating factor (GM-CSF). In the absence of GM-CSF,
pre-cDC derived DCs gave rise to a homogeneous population of CD11clow MHClow cells (DC-
regs) on day 8-10 of culture. DC-regs failed to up-regulate major histocompatibility complex
class II (MHCII) and co-stimulatory molecules in response to DC maturation stimuli, were poor
stimulators in T cell proliferation assays and suppressed T cell proliferation in cultures
containing immuno-stimulatory DC. Co-transfer of DC-regs with DCs in vivo did not inhibit
proliferation of T cells. These findings reveal the potential of DCs to generate a regulatory DC
population with immunosuppressive properties.
iii
Acknowledgments
I would like to thank my supervisor, Dr. Mark Cattral for providing me with the opportunity to
learn, grow, and evolve under his mentorship. I am extremely appreciative of all his support,
guidance, and advice.
I would also like to thank my co-supervisor, Dr. Reginal Gorczynski for his support, wisdom,
constructive criticism and feedback on my work as well as inspiration. I am very lucky to have
such wise superiors as supervisors.
I would like to thank Jun Diao and Jun Zhao for training in logic, design of scientific
experiments and technical training. I would not achieve what I have without your support.
Also, I would like to thank all members of Gorczynski Lab for ongoing support, conversations
and humor.
Thank you also to my family for their support and understanding.
Thank you also to Heart and Stroke Foundation and Canadian Institute for Health and Research
for their financial contribution that allowed this research to take place.
iv
Table of Contents
Abstract .......................................................................................................................................... ii
Acknowledgments ........................................................................................................................ iii
List of Figures .............................................................................................................................. vii
CHAPTER 1 .................................................................................................................................. 1
1 Types of DCs .......................................................................................................................... 1
1.1 Conventional DCs ........................................................................................................... 1 1.2 Plasmacytoid DCs ........................................................................................................... 2 1.3 Lymphoid tissue resident DCs ........................................................................................ 3 1.4 Peripheral tissue DCs ...................................................................................................... 5 1.5 Intestinal DCs.................................................................................................................. 7 1.6 Thymic DCs .................................................................................................................... 7
2 Current model of DC ontogeny ........................................................................................... 8
2.1 Generation of DC from monocytes ................................................................................. 9 2.2 In vitro methods of dendritic cell generation ................................................................ 11
3 Dendritic cell activation of immunity ................................................................................ 11
4 Dendritic cell activation of tolerance ................................................................................ 12
4.1 Regulatory dendritic cells ............................................................................................. 12 DCs in central tolerance ....................................................................................................... 12
DCs in peripheral tolerance ................................................................................................. 13
5 Mechanisms of immunosuppression ................................................................................. 14
5.1 PD-L1/PD-L2 ................................................................................................................ 14 5.2 Arginase and nitric oxide synthase (NOS).................................................................... 15 5.3 Indoleamine 2,3-dioxygenase (IDO) ............................................................................ 17 5.4 IL-10 ............................................................................................................................. 18
6 Antigen presenting cells in cancer immunology ............................................................... 19
6.1 Tumor-derived dendritic cells ....................................................................................... 19 6.2 Myeloid derived suppressor cells (MDSC)................................................................... 20 6.3 Tumor associated macrophages (TAMs) ...................................................................... 21
CHAPTER 2 ................................................................................................................................ 23
v
1 Preliminary observations ................................................................................................... 23
2 Hypothesis ............................................................................................................................ 24
3 Objective .............................................................................................................................. 24
4 Specific aims ........................................................................................................................ 24
CHAPTER 3 Methodology ........................................................................................................ 25
1 Mice ...................................................................................................................................... 25
2 Primary skin stromal cell preparation .............................................................................. 25
3 Cell Isolation ........................................................................................................................ 25
4 Flow cytometry .................................................................................................................... 26
5 Mixed lymphocyte reactions .............................................................................................. 26
6 CFSE labelling ..................................................................................................................... 27
7 Reverse transcriptase PCR ................................................................................................ 27
8 Arginase and iNOS activity assays .................................................................................... 28
9 Adoptive transfer studies ................................................................................................... 28
10 Statistics ............................................................................................................................... 29
CHAPTER 4 Results .................................................................................................................. 30
1 CD11clow MHCIIlow cells exhibit potent immuno-suppressive properties in vitro ......... 30
2 Mechanisms of immuno-suppression mediated by CD11clow MHCIIlow DC-regs ......... 32
3 Immunosuppressive activity of CD11clow MHCIIlow cells in vivo .................................... 35
CHAPTER 5 Discussion ............................................................................................................. 37
vi
CHAPTER 6 ................................................................................................................................ 46
1 Conclusion ........................................................................................................................... 46
2 Future directions ................................................................................................................. 46
References .................................................................................................................................... 73
vii
List of Figures
Figure 1. Dendritic cell subtypes grouped based on physiological location .................................. 4
Figure 2. Schematic representation of DC ontogeny. .................................................................. 10
Figure 3. Differentiation and proliferation of pre-cDC on stroma .............................................. 50
Figure 4. CD11clow MHCII low arise from CD11c+ MHC II+cDC ................................................ 51
Figure 5. Immunophenotype of cDC-derived CD11clow MHCII low cells ..................................... 52
Figure 6. DC-regs fail to up-regulate co-stimulatory molecules in response to maturation stimuli........................................................................................................................................................ 53
Figure 7. CD11clow MHCII low cells are poor stimulators of allogeneic T cell lymphocytes ........ 54
Figure 8. CD11clow MHCII low DC-derived cells have increased phagocytic capacity. ............... 55
Figure 9. CD11clow MHCII low suppress T cell proliferation in allogeneic mixed lymphocyte reaction. ......................................................................................................................................... 56
Figure 10. DC-regs suppress effector function of allogeneic T cells in mixed lymphocyte cultures .......................................................................................................................................... 57
Figure 11. DC-regs suppress OT-II T cell proliferation in response to OVA-pulsed DCs .......... 58
Figure 12. DC-regs suppress OT-II T cell proliferation .............................................................. 59
Figure 13. DC-regs do not suppress OT-II T cell cytokine release ............................................. 60
Figure 14. Expression of CD25 and CD44 by T cells in allogeneic mixed lymphocyte reaction in the presence or absence of DC-regs .............................................................................................. 61
Figure 15. DC-regs do not induce T cell death in mixed lymphocyte reactions .......................... 62
Figure 16. DC-regs do not induce Foxp3+ Tregs in mixed lymphocyte cultures ........................ 63
Figure 17. Mechanism of DC-reg-mediated immuno-suppression involves both soluble and contact-dependent factors ............................................................................................................. 64
Figure 18. RT-PCR expression of candidate molecules responsible for observed in vitro immuno-suppression ..................................................................................................................... 65
Figure 19. DC-regs express high levels of arginase1 and iNOS (reflected by nitrite production) activity when stimulated with LPS ............................................................................................... 66
viii
Figure 20. Both DCs and DC-regs express high levels of PD-L1. 1×106 DC or DC-regs were pulsed with 2 µg/mL LPS overnight. ............................................................................................ 67
Figure 21. LPS-pulsed DC-regs express high levels of IL-10. Freshly isolated spleen DCs or DC-regs were pulsed with 2 µg/mL LPS overnight ..................................................................... 68
Figure 22. DC-regs suppress T cell proliferation through an iNOS-dependent mechanism ....... 69
Figure 23. DC-regs do not suppress T cell proliferation and activation in vivo .......................... 70
Figure 24. DC-regs fail to suppress T cell proliferation and activation in vivo ........................... 71
Figure 25. DC-regs induce OT-II cell activation in vivo ............................................................. 72
1
CHAPTER 1
Introduction
Dendritic cells comprise a heterogeneous population of cells that are specialized in antigen
uptake, processing, and presentation, and play a key role in linking innate and adapitive immune
responses. Dendritic cells were first discovered in mouse spleen by Steinman in 1973 1, and
were named dendritic cells because of their numerous motile cellular processes - dendrites.
Subsequent studies revealed that DCs were potent stimulators of allogeneic T cells in mixed
lymphocyte reaction 2 and their potency exceeded that of other “professional” antigen presenting
cells (i.e. macrophages and B cells).
1 Types of DCs
DCs are currently divided into two major categories: 1) interferon-producing plasmacytoid
(pDCs); and 2) conventional DCs (cDCs). DC can be further divided based on their location
(lymphoid, migratory), expression of cell-surface markers, and functional attributes. Lymphoid
resident DCs reside within lymphoid tissues throughout their life cycle, whereas migratory DCs
migrate from peripheral tissues to the lymph nodes. DC migration occurs continously during
steady-state conditions, and increases with inflammation. DCs can also be classified based on
their ability to polarize differentially T cell responses in tolerance and immunity.
1.1 Conventional DCs
Conventional DCs (cDCs) have a typical heterogeneous morphology with abundant cytoplasm,
multiple dendrites and irregular nucleus 5. These cells are widely distributed in lymphoid and
peripheral tissues (Figure 1). cDCs are superior to macrophages and B cells in Ag presentation
because of their higher capacity to capture and process Ag 6. Upon encounter of foreign Ag,
cDCs undergo a process of maturation where they up-regulate surface expression of MHCII and
co-stimulatory molecules (CD80, CD86, CD40) and activate naïve Ag-specific T cells. Three
major subtypes of cDCs exist: CD4+ CD8-, CD4- CD8+, CD4- CD8-, all of which are found
2
within multiple physiological compartments in the body and differ in their ability to induce
differential T cell responses. These subsets are described in detail below.
Physiological DC localization is driven by expression of chemokine receptors. Immature cDCs
express receptors for inflammatory chemokines including CXCR1, CCR1, CCR2 and CCR5 17,
which drive DC migration towards lymphoid organs. Chemokines produced by freshly isolated
splenic cDCs were identified as macrophage inflammatory protein 1 alpha (MIP-1α or CCL3),
MIP-1β or CCL4 and regulated upon activation, normal T cell expressed and secreted (RANTES
or CCL5). Moreover, it was observed that different subsets of splenic cDCs express these
chemokines in different proportions with all three (CCL3, CCL4 and CCL5) expressed highest
on CD4+ cDCs 18. During maturation, when cDCs encounter foreign Ag and inflammatory
stimuli, they upregulate MHCII, co-stimulatory molecules and their migratory capacity. Mature
cDCs migrate in response to chemokines to lymph nodes via afferent lymphatics and localize in
T cell areas of LN. DC migration to peripheral lymphatic vessels is guided by the
chemoattractant gradient of CCL19 and CCL21, which bind to chemokine receptor CCR7 found
on cDCs 19, 20. CCR7 is upregulated after DC encounter maturation stimuli 21. It was
demonstrated that CCR7 KO mice not only have deficient DC and T cell migration to LN, but
also fail to mount primary immune response 19. CXCR4 was also observed to be upregulated on
mature cDCs 20. Additionally, sensitivity of DCs to CCL3, CCL4 and CCL5 22 as well as
expression of CCR1 and CCR5 21 upon maturation is dramatically reduced.
1.2 Plasmacytoid DCs
Plasmacytoid DCs (pDCs) were first identified in humans and were later shown to exist in mice 45 as lin- CD11cint CD11b- Ly6C+ B220+ cells. pDCs were identified in lymphoid organs, bone
marrow, lung, liver, blood and skin 5, 46. Poor in vitro survival is observed when cells are
cultured in liquid medium alone. Survival is moderately enhanced when medium is
supplemented with GM-CSF alone or in combination with IL-3 47. Further survival and
maturation are induced with addition of IFN-α, influenza virus, CpG or CpG and GM-CSF 5, 45.
The in vivo life span of pDCs was determined to be about two weeks 48.
Immature pDCs have a round shape, smooth surface and eccentric nucleus and acquire dendritic
cell-like morphology upon activation with CD40L or CpG. Similarly to cDCs, pDCs upregulate
3
surface MHCII as well as CD40 and CD86 expression. In contrast to cDCs, freshly isolated
pDCs fail to stimulate T cell proliferation in allogeneic mixed lymphocyte reaction and only do
so upon maturation 45. When activated with viral stimuli pDCs produce large amounts of type I
IFNs (IFNα, IFNβ) and moderate levels of IL-12 5. They migrate to inflamed LN and cluster
around high endothelial venules 47. In contrast to pDCs, cDCs and monocyte migrate from non-
lymphoid tissues to T cell rich areas of lymph nodes through afferent lymphatics 48. pDCs
express TLR7, TLR8, which recognize imidazoquinolins and ssRNA; and TLR9, which
recognizes bacterial DNA 49. pDCs, however, lack TLR2, 3, 4, 5 and, therefore, do not respond
to microbial stimuli such as LPS or poly I:C 15, 46. Immature pDCs were shown to induce IL-10
production in CD4+ T cells 50. It was also suggested that immature pDCs are able to induce Treg
cells in vitro 51. Activation of pDCs leads to rapid activation of NK cells and CD8+ T cell, IFN-γ
production and Th1 differentiation leading to anti-viral responses in both humans 50 and mice 15,
52. Moreover, they promote differentiation and maturation of cDCs 52 and stimulate B cells 46.
1.3 Lymphoid tissue resident DCs
Spleen contains about 20% pDCs and 80% cDCs 6 (Figure 1). Three major populations of cDCs
can be subdivided based on CD4 and CD8 staining: CD4+ CD8α- (60% of total); CD4- CD8α+
(20%); CD4- CD8α- (20%) 7 8. Other cell surface markers segregate with CD4 and CD8. For
example, CD8α+ cDC are CD11b- DEC-205+, whereas CD8α- cDC are CD11b+ and DEC-205-.
Both CD4+ CD8α- and CD4- CD8α- DCs exist mainly in the marginal zone in the steady state
and move to T cell zones upon maturation. The marginal zone is located between the red pulp,
which filters the blood of damaged red blood cells and other debris, and the white pulp, which
mainly contains lymphocytes. By contrast, CD8α+ cDCs are located in T cell zones within the
white pulp in the steady state 6. LNs contain all DC subsets seen in spleen along with additional
migratory DC subtypes (dermal DCs and epidermal Langerhans cells). CD8aint and CD8αlow
DCs are also present in LN, but not in spleen 9. All splenic cDC subpopulations showed half-life
kinetics of about 1.5 days, with all cells replenished by day 3 10.
4
CD8a- CD4+
CD8a+ CD4-
CD8a- CD4-
pDC
cDC
Lymph nodes
pDC
cDC
CD8a- CD4+
CD8a+ CD4-
CD8a- CD4-
CD8alow
CD8ahi
Dermal DC
Langerin+ LC
Peripheral
tissues
Langerin+ LC
CD8+ Langerinlow DC
CD8- Langerinhi DC
Resident
Migratory
Intestine:
Peyer Patches
CD11b= CD103+ CX3CR1=
CD11b+ CD103- CX3CR1+
Intestine:
Lamina Propria
CD11b+ CD8-
CD11b- CD8+
CD11b- CD8-
cDC
Thymus
pDC
CD11b- CD8+ CD172- cDC
CD11b+ CD8- CD172+ cDC
cDC
LEGEND
DENDRITIC CELL SUBTYPES
Spleen
Figure 1 Dendritic cell subtypes grouped based on physiological location. pDC, plasmacytoid DC; cDC, conventional DC; LC, langerhan cell
5
Although all splenic cDC subtypes capture antigens effectively, each have specialized properties
in inducing Ag-specific responses 10. For example, CD8+ DCs possess a potent capacity for
cross-presentation of soluble and cell-associated Ag to CD8+ T cells 11. CD8a+ DCs also release
high levels of IL-12 and preferentially induce Th1 responses. By contrast, CD8- DCs show
superior priming of CD4+ T cell 12, and preferentially induce Th2 responses 13, 14. However, it is
now recognized that the microenvironment plays a large role in directing how DCs skew Th
responses. It appears that different subtypes of DCs are specialized to recognize a particular type
of microbial stimulus and release a defined array of cytokines, which directs T cell
differentiation from Th0 into Th1 or Th2. For example, microbial molecules such as soluble
tachyzoite Ag (STAg) and CpG trigger cDCs to pomote Th1 differentiation, whereas nematode
antigens or yeast toxin trigger cDCs to drive Th2 differentiation 13, 15.
Microbial structural units or pathogen associated molecular patterns (PAMPs) are recognized by
Toll-like receptors expressed by DC. TLR expression on cDCs appears to be fairly ubiquitous,
with some exceptions. TLR3, which recognizes viral double stranded RNA, is expressed highest
on CD8α+ cDC and lowest on CD4+ cDC and vice versa for TLR5, which recognizes bacterial
flagellin. Additionally, TLR7, which recognizes endosomal single stranded RNA, has low
expression on CD8α+ cDC but is expressed on the other cDC subtypes 16. Therefore, ligation of
different TLRs determines the subtype of DC activated, which subsequently drives appropriate T
cell response. It has also been suggested that Ag dose affects CD4+ T cell response directed by
cDCs. Both CD8a+ and CD8a- cDCs were observed to induce Th1 response at high Ag doses and
Th2 response at low Ag doses 15.
1.4 Peripheral tissue DCs
Langerhan cells (LCs) or epidermal DCs and dermal DCs reside in the periphery and sample Ag
from skin and mucosal body surfaces. LCs account for 3-5% of all nucleated cells in the
epidermis and form a cellular network that provides the first immunological barrier to
environmental insults 23, 24. LCs express langerin (CD207), which is also expressed on some
dermal DC subtypes 25, 26. LC are also distinguished by expression of CD45, CD11c, CD11b,
F4/80, DEC-205, high expression of MHCII, absence of CD103 and the presence of Birbeck
granules in the cytoplasm 27, 23. The formation of Birbeck granules, which is associated with Ag
6
capture, is a consequence of langerin expression 24. Upon Ag capture, langerin associates with
Birbeck granules and facilitates transport of captured ligands into a non-classical Ag-processing
pathway 25. Upon capture of Ag, LCs migrate via dermal lymphatics to skin draining LN 6. The
rate of migration increases during inflammation. Using a model of allergic contact dermatitis it
was demonstrated that LCs mediate immunity to cutaneous Ag 28. However, another study
showed that LCs are unable to prime CD8+ T cells in epidermal infection with Herpes Simplex
virus 218. The precise role of LCs in cutaneous immunity, therefore, appears to depend on the
type of Ag.
LC in subcutaneous LN can be distinguished from other DC subtypes by their larger size, higher
expression level of MHCII, low CD11b expression and intermediate CD8 expression 9. In the
steady state, LCs arise from a local pool of radioresistant hematopoietic precursors 25, 29. During
inflammatory processes, circulating monocytes appear to have a role in replenishing LCs31.
TGF-β is required for LC differentiation or maintenance 30 and mice lacking M-CSFR also lack
LCs 31. LCs turnover is slower than for other DC subtypes, as shown by only 50% of the cells
staining positive for BrdU at day 2132.
Another migratory DC subtype found in the skin, cutaneous and mesenteric lymph nodes
displays the phenotype of langerin+ CD11c+ MHCII+ CD11b+ CD205+ F4/80low CD103- 26
(Figure 1). Other langerin+ DC subtypes exist. For example, langerin positive dermal DCs 26, 33
divide into CD8+ langerinlow and CD8- langerinhi subsets 25. Langerin+ DCs also express high
levels of CD11c and MHCII 26. However, they can be distinguished from LCs by the presence
of CD103 marker 33, which is not expressed on LCs, and by the absence of F4/80 expression and
Birbeck granules 23. Additionally, LCs express higher levels of CD11b and the adhesion
molecule EpCAM 33. Dermal DCs were observed in the dermis when LCs were conditionally
ablated 26, 33. The life time of dermal langerin+ DCs is much shorter than that of epidermal LCs
and is marked by rapid repopulation of these cells with conditional ablation. Both dermal DC
populations repopulate dermis within 5 days (for CD8+ langerinlow DCs) and within 14 days (for
CD8- langerinhi subset) from bone marrow precursors migrating from the blood, long before LCs
repopulate epidermis 25, 26. BrdU labelling studies also demonstrated that langerin+ dermal DCs
proliferate at a higher rate than LCs 26. The kinetics of dermal DCs proliferation is similar to
7
those of spleen and LN DCs. Moreover, development of these cells seems to be independent of
TGFβ and M-CSFR.
1.5 Intestinal DCs
In the gut, DCs are found in the lamina propria (LP) of the small and large intestine (loose
connective tissue underlying gut epithelium) as well as in gut associated lymphoid organs such
as Peyer’s patches (PPs), mesenteric lymph nodes (MLN) and isolated lymphoid follicles. LP
DCs directly sample the luminal environment of the gut by penetrating epithelial tight junctions 37. Two main DC subtypes in the LP are defined by cell surface markers: CD11b- CD103+
CX3CR1- and CD11b+ CD103- CX3CR1+ 34, 35 (Figure 1). CD11b+ DCs originate from
monocytes 34, whereas the CD11b- subset originates from pre-cDC 34. CD11b+ DCs appear to
promote inflammaton because ablation of CD11b- DCs exacerbated colitis in a murine model.
Secretion of TNF-α 34 and induction of a Th17 response was associated with the development of
colitis 35, 36. At low APC:T cell ratios, all LP DC subsets can induce generation of FoxP3+ Tregs 35.
The main DC subsets in PP are CD11b+ CD8a-, CD11b- CD8a+ and CD11b- CD8a- 38 (Figure 1).
CD11b+ CD8a- DCs were found to be located in the subepithelial dome of PP where they pick up
Ag transported across intestinal epithelium by M cells. A CD11b- CD8a+ fraction was detected
exclusively in the interfollicular region where it likely activates naïve T cells 38. Here, CD11b+
CD8a- DCs were shown to induce differentiation of IL-10 and IL-4 producing Th2 cells 39 40.
CD11b- CD8a+ and CD11b- CD8a- DCs were shown to produce IL-12 and induce Th1 type
responses 39. CD11b- CD8a- DCs are located in both PP compartments. Gut DCs were shown to
be involved in tolerance induction towards oral Ag and commensal bacteria.
1.6 Thymic DCs
DCs in the thymus constitute only 0.5% of total cell number 41 and localize almost exclusively to
the thymic medulla 42. Three thymic DC subsets have been identified. Two are of the cDC
phenotype: CD11chi CD11b- CD8a+ CD172- and CD11chi CD11b+ CD8a- CD172+ 43 (Figure 1).
These cells displayed all markers typical of mature DCs and were similar to CD8a+ DCs in
spleen 7. About 35% of total thymic DCs are pDCs, which stain CD11cint MHCII low CD45RAhi
8
41. It has been demonstrated recently that only the CD11chi CD11b- CD8a+ subset is of
intrathymic origin and that CD11chi CD11b+ CD8a- and pDC subsets are emigrants into the
thymus 43. Migratory cDC and pDC take up circulating soluble and particulate Ag and transport
it to the thymus 43. Shortly after emigration into the thymus, migratory DCs mature and
upregulate both MHCII and co-stimulatory molecules. However, most thymic DCs are present in
an immature state with low expression of co-stimulatory molecules and moderate expression of
surface MHCII 44.
2 Current model of DC ontogeny
DCs develop from bone marrow derived lin- Sca+ c-kithi hematopoietic stem cells (HSC) 2
(Figure 2). Early studies suggested that DCs belong to myeloid lineage and arise from a common
myeloid progenitor (CMP) 53. More recent evidence suggests developmental flexibility exist in
the DC lineage. Early stages of differentiation can occur from both a common lymphoid
progenitor (CLP) and CMP. A CMP gives rise to the myeloid lineage of immune cells including
macrophages, monocytes, megakaryocytes and erythrocytes. CLP gives rise to lymphoid lineage
cells such as NK cells, T and B cells.
The first evidence for a possible lymphoid origin of DC came from the observation that early T
cell precursors in the thymus can generate thymic DC 54. Later it was observed that CD8a+ DCs
in both spleen and thymus differentiate from thymic T cell progenitors 55. For three years, CD8a+
DCs were considered to be of lymphoid origin and CD8a- of myeloid origin. In 2000, it was
reported that CD8a+ and CD8a- DCs were both capable of differentiating from CMP 56 and that
both DC types differentiated from the same CD4low lymphoid precursor population 57. Finally,
both CMP and CLP were observed to give rise to DC with similar efficiency 58 59. A CMP was
defined as lin- FcRyII/IIIslow CD34+ c-kit+ Sca-1- IL7Ra- and a CLP as lin- c-kitint Sca-1int IL7Ra+
Thy1.1- 59. Accordingly, the concept of myeloid vs lymphoid DC was abandoned.
The next steps in DC differentiation after CMP and CLP involve a sequential series of
precursors: common macrophage DC progenitors (MDP; lin- c-kit+ CX3CR1+ CD115+ Flt3+)60;
common DC precursors (CPDs) 61; and CD11c+ MHCII - B220- pre-cDCs and B220+ pre-pDCs,
9
which give rise exclusively to cDCs and pDCs respectively 62, 63. CD24- pre-cDC are committed
to CD8a- cDCs and CD24+ pre-cDCs are pre-committed to CD8a+ cDCs 64.
The distinguishing feature of early stage precursors that give rise to the DC lineage is the
expression of fms-like tyrosine kinase 3 (Flt3) 65. In particular, the majority of CLP and some
CMP express Flt3 65, 66. Flt3 is then progressively down-regulated in granulocyte macrophage
progenitor (GMP) downstream of CMP and is completely absent in cells committed to both T
and B lineages as well as to megakaryocyte/erythrocyte lineage 66. Flt3 expression is also absent
in mature cells of hematopoietic lineage but present on most subtypes of DCs. Moreover, mice
lacking Flt3L have deficient DC lineage haematopoiesis 67 whereas treatment with Flt3L
dramatically increases dendritic cell population in spleen, lymph nodes, blood and other organs 68. By contrast, administration of GM-CSF alone or GM-CSF and IL-4 – cytokines that are
commonly used to generate DC in vitro –have little effect on DC generation in vivo 68.
Furthermore, mice lacking GM-CSF have normal DC numbers in lymphoid tissues 69.
Initial experiments evaluating DC proliferation showed very low numbers of BrdU+ DCs after 2
hours of labelling. This finding led to the conclusion that peripheral DC were replenished solely
by migrating non-replicating precursor populations 10, 32. Kinetic studies revealed a short half-
life of about 1.5 days for splenic cDCs 32. Subsequent studies, however, challenged this view
when it was established that 4% of spleen DCs and 3.6% of bone marrow DCs were in the
S/G2/M phases of the cell cycle 63 70. It is now generally accepted that in situ DC proliferation
plays a key role in maintaining the peripheral DC pool. In addition, dividing DCs can pass on
Ag to their progeny, thereby prolonging and expanding the potential for antigen presentation.
2.1 Generation of DC from monocytes
Monocytes are heterogeneous cells of the mononuclear phagocyte system that constitute less
than 2% of peripheral blood cells. Their phenotype is CD11b+ CD115+ F4/80+. They arise in
bone marrow and are released into blood to give rise to macrophages and dendritic cells in
tissues. Two monocyte subtypes exist: inflammatory monocytes are CCR2+ CD62L+ CXCR3-
GR-1+ (Ly6C+) and resident monocytes are CCR2- CD62L- CXCR3+ GR1- (Ly6C-) 72. During
inflammation GR-1+ inflammatory monocytes migrate into inflamed tissue and differentiate into
macrophages and into DCs in draining lymph nodes 72, 73. GR-1- resident monocytes home to the
10
non-inflammed tissues in the steady state to give rise to macrophages and DCs 72, 74. Bone
marrow derived monocytes were shown to replenish DC populations in peripheral tissues but not
in spleen 74. However, the monocyte lineage and monocyte-derived DCs were shown to be
distinct from DC lineage 64. In the steady state, pre-cDCs have a far superior ability to generate
spleen DCs when compared to monocytes, which were only 2% as effective 64. Flt3 is expressed
only on myeloid and lymphoid progenitor derived DC but not on monocyte-derived DC 66.
Systemic administration of Flt3L but not GM-CSF, which is a monocyte growth factor
responsible for DC differentiation from monocytes, expands the DC pool in vivo 68.
CMP/CLP
pDC
cDC
Lymphoid tissues
HSC
Pre-pDC
Pre-cDC
CDP
(pro-DC)
MDP
Bone Marrow
monocytes
Non-lymphoid tissues
Inflammatory DC
Steady state DC
\\
Figure 2 Schematic representation of DC ontogeny. HSC, hematopoietic stem cell (Lin- Sca+ c-kithi); CMP, common myeloid progenitor (Lin- FcRγII/IIIs low CD34+ c-kit+ Sca-1- IL7Ra- Flt3+); CLP, common lymphoid progenitor (Lin- c-kitint Sca-1int IL7Ra+ Thy1.1- Flt3+); MDP, monocyte dendritic cell progenitor (Lin- c-kit+ CX3CR1+ CD115+ Flt3+); CDP, common DC progenitor (c-kit low CD115+ Flt3+); pre-cDC, immediate precursor of cDC (CD11c+ MHCII - B220-); pre-pDC, immediate precursor of pDC (CD11c+ MHCII - B220+); cDC, conventional DC; pDC, plasmacytoid DC.
11
2.2 In vitro methods of dendritic cell generation
When generated from bone marrow or peripheral blood, DCs are non-adherent or loosely
adherent cells which can be characterized based on expression of CD11c and MHC class II and
lack of lineage (lin) markers (CD3, CD19, B220, CD49b) 3, 4.
Inaba et al 75 was the first to describe a method for generating DC from cultured blood cells.
Cells suspensions were depleted of red blood cells and cultured overnight in GM-CSF. Non-
adherent cells were then removed and cultured for 10 more days and supplemented with GM-
CSF for last 3-4 days. Dendritic cell aggregates attached to an adherent monolayer were
harvested and cultured for further 4-10 days in the presence of GM-CSF 75. It was then
demonstrated that DCs could be obtained by culturing bone marrow supplemented with GM-CSF
alone or in combination with IL-4 for 6-10 days. Loosely attached cells are then harvested and
matured for 1-2 days in the presence of GM-CSF and TNFα or LPS 3, 4. This method generates
monocyte-drived DC similar to that which arise from monocytes in vivo during inflammatory
conditions. Alternatively, DCs can be generated by culturing bone marrow in liquid with human
Flt3L (100 ng/mL) for 9 days. These cultures generate both pDC and cDC and appear to
recapitulate DC generation under steady-state conditions. Non-adherent and loosely adherent
cells are harvested and matured in the presence of GM-CSF as well as IFN-γ or LPS for 24
hours76. Another method of generating DCs first described in Cattral’s laboratory involves
culturing immediate DC precursor, pre-cDC, on a stromal monolayer in the presence of GM-CSF
for 12 days 62. Mimicking physiologic conditions, this method generates immature DCs, which
can be matured overnight with LPS or TNFα to produce mature DCs. Culture of DC precursors
on a stromal monolayer generates a highly pure, homogeneous DC population, which is not
possible with bone marrrow culture. Moreover, the stromal monolayer allows for greater
expansion and longer survival of DCs as compared to liquid culture systems 212.
3 Dendritic cell activation of immunity
In the immunogenic model of DC activation, DC maturation is triggered by ligation of pattern
recognition receptors (PRRs) like TLRs, NODs, RIG-I-like, and c-type lectin receptors 6 and by
other pro-inflammatory signals such as cytokines that indicate injury or inflammation. Upon
12
activation, DC migrate from peripheral tissues into draining lymph nodes where they present Ag
and subsequently activate naïve T cells. Inflammatory cytokines secreted by Th1 and Th2 cells,
as well as ligation of CD40 on DC surface by primed T cells also provides activating signals.
During maturation, DCs upregulate surface expression of MHCII and co-stimulatory molecules
CD80 and CD86, which bind CD28 on T cell surface. They also release pro-inflammatory
cytokines such as IL-12, which triggers T cell activation and proliferation.
4 Dendritic cell activation of tolerance
Tolerogenic DCs generally have a distinct phenotype from that of mature stimulatory DCs. They
express low levels of surface MHCII and co-stimulatory molecules CD80, CD86 and CD40, do
not mature in response to classical DC activation stimuli (such as LPS, TNFa) and have low
capacity to prime T cells 77.
In the steady state in lymphoid organs DCs are present in immature state. It has been shown that
these DCs are able to sample and present Ag in the context of MHCI and MHCII without
maturation. For example, DCs sample apoptotic debris or self Ag and present these to naïve T
cells. Injection of immature DCs loaded with Ag renders T cells non-responsive to Ag (anergy)
and triggers Ag-specific T cell deletion or development of Tregs. Several factors can render DCs
tolerogenic. Both innate and adaptive immune systems can create local tolerogenic environment
dominated by immuno-suppressive cytokines such as IL-10 or TGFβ. Apoptotic debris can also
provide tolerogenic signals to DCs. Treg subsets such as FoxP3+ CD4+ and Tr1 cells can induce
DC tolerance.
4.1 Regulatory dendritic cells
DCs in central tolerance
Involvement of DCs in central tolerance was first mentioned in 1985 78. Four years later
Matzinger showed that when fetal thymuses were incubated with splenic DCs, donor-specific
tolerance developed 79. The role of thymic dendritic cells in central tolerance was later
demonstrated by studies of targeted expression of MHCII on DCs. It was demonstrated that these
13
cells mediate negative selection but not positive selection 80, 81. In one such study 80, the MHC
class II I-E transgene was expressed in DCs under a CD11c promoter in a C57BL/6 mouse. I-E-
specific T cells were deleted in this mouse. The frequency of I-E-specific T cells was much
lower then that of the wild type animal but equivalent to that of animals expressing MHC class II
I-E in all tissues. These results indicate that I-E expressing DCs mediated negative selection of I-
E-specific thymocytes. It was then shown that thymic DCs are able to pick up tissue-specific Ag
from medulary thymic epithelial cells and delete autoreactive T cells via cross-presentation 82.
Thymic DCs were also directly shown to delete Ag-specific single positive thymocytes in vivo 83.
Recently it was demonstrated that thymic CD11b+ CD8a- CD172+ DCs are also capable of
inducing natural Tregs 84.
DCs in peripheral tolerance
DCs are strategically positioned in the periphery (eg. skin, airway, and intestine) to capture Ag
and present Ag to T cells in draining LN. In the steady state, these DCs mediate peripheral cell
tolerance to harmless environmental Ag (ingested 85 or inhaled 86) or self-Ag. When exogenous
soluble Ag are fed to mice or introduced by inhalation, Tregs or Tr1 cells are induced via IL-10
or TGFβ. High levels of MHCII expression bound to self-Ag was observed on LN DCs. These
DCs were able to induce apoptosis of Ag-reactive T cells 86. Peripheral tolerance induction has
also been noted in DCs that present non-self Ag in immature state 87, 88, 89. In this situation, TCR
stimulation is not accompanied by co-stimulatory signal and anergy or deletion of peripheral T
cell takes place 87, 88, 90. Induction of IL-10-producing Tr1 cells has also been reported 89. In
addition, naïve T cells may be converted into CD4+ FoxP3+ Tregs or IL-10-producing Tr1 cells 91. CD103+ migratory DC in the gut 92, 93 and CD103- migratory DC in the skin 94 have been
observed to transport Ag to mesenteric LN and induce naïve CD4+ T cells to become Tregs via
TGFβ and retinoic acid dependent pathway.
Splenic CD8a+ DCs have been observed in multiple studies to be capable of tolerance induction.
In an airway hypersensitivity model, CD8a+ splenic DCs were able to inhibit Th2 cytokine
response and reverse airway hyper-responsiveness in vivo 95. Another study demonstrated that
spleen CD8a+ CD205+ DCs can convert naïve CD4+ T cells into Tregs via secretion of TGFβ 91.
Moreover, targeting OVA to DEC-205 – a scavenging receptor expressed on DC surface – has
14
been shown to induce tolerance of T cells to OVA. In one such study, adoptively transferred OT-
I cells exhibited defective cytokine production and were deleted from the system 90. Similarly,
anergy induction of Ag-specific T cells occurred in mice previously targeted with DEC-205-Ag
conjugates 87. When DCs are pulsed with apoptotic debris containing OVA, tolerance usually
ensues to OVA 88. It is important to note that in all of these studies, T cell proliferation was
evident at day 3, where as at time points beyond 10 days these T cells were anergic or deleted.
Numerous attempts to manipulate DCs to acquire stable tolerogenic properties have also been
undertaken. The rational for this approach is to avoid the use of immature DCs as these have an
unstable phenotype and can be converted to stimulatory DCs easily by exposure to inflammatory
conditions 77. Several groups have been able to convert immature DC into tolerogenic DCs by
using variety of culture conditions including low levels of GM-CSF, IL-10 96, 97 , or the
combination of IL-10 and TGFβ 77. For example, CD45RB+ CD11clow IL-10-producing
regulatory DCs were generated from c-kit+ progenitors by culturing them on spleen stromal
monolayer 98 99. Moreover, these cells could induce conversion of CD4+ T cells into FoxP3+
Tregs 96, induce IL-10-producing Tr1 cells, and induce T cell anergy 77, 97, 98. In vitro generated
tolerogenic DC have been used in adoptive transfer therapy to induce peripheral tolerance in
vivo, prolong allograft survival, and prevent GVHD 77.
Certain pathogens have also been observed to induce DC reprogramming towards tolerogenic
type. In particular, fungal morphocyte haphae induces DCs to activate Treg cells. S. masoni
conditions DCs through TLR2 signalling to induce Tregs. Filamentous hemagglutinin from
bacteria Bordetella Petrussis induces DCs to secrete IL-10 and prime Tr1 cells 100.
5 Mechanisms of immunosuppression
5.1 PD-L1/PD-L2
The B7 family of ligands includes co-stimulatory molecules (CD80 (B7-1) and CD86 (B7-2))
and two in inhibitory molecules (programmed death-1 ligand (PD-L1) or B7-H1 and PD-L2 or
B7-H2). In humans, IFN-γ induces PD-L1 and PD-L2 expression on PBMCs. When human
monocytes are cultured in the presence of IFN-γ, both PD-L1 and PD-L2 expression are induced
15
101, 102. PD-L1 expression was also high on both mature and immature human monocyte-derived
DCs 101 and was further increased upon stimulation with IFN-γ and LPS 102. The expression
pattern was similar on murine DC counterparts 102. PD-L1 was also expressed on human CD4+
and CD8+ T cells that were activated with anti-CD3 Ab. Moreover, PD-L1 expression was
observed on cells of non-hematopoietic lineage, such as vascular endothelial cells 103.
Both PD-L1 and PD-L2 were shown to inhibit T cell proliferation and cytokine production
through engagement of a PD-1 receptor 102, 104. PD-1 is expressed on activated B, T and myeloid
cells 105, 106. Engagement of PD-1 results in T cell cycle arrest 104 by limiting the production of
IL-2 105. When T cells were stimulated with immature DCs in allogeneic MLR in the presence of
anti-PD-L2 Ab, both proliferation and IFN-γ production were increased. The same result was
observed when a combination of anti-PD-L1 and anti-PD-L2 Abs were used, but not anti-PD-L1
alone 101. The same pattern was observed in an Ag-specific system. When CD4+ DO11.10 cells
were stimulated with DCs pulsed with OVA in the presence of anti-PD-L1, anti-PD-L2 or both
Abs, cytokine secretion was significantly increased. Moreover, PD-L1/PD-L2-/- NOD mice had
rapid onset of autoimmune disease, significantly earlier than their NOD counterparts 103.
5.2 Arginase and nitric oxide synthase (NOS)
Arginase metabolizes L-arginine to produce urea and L-orthinine, whereas NOS metabolizes L-
arginine to produce nitric oxide (NO) and L-citrulline 107. When NO combines with oxygen it
produces anions (NO2-, NO3
-) and peroxynitrites (ONOO-), which damage cellular lipid, protein
and DNA 108.
Arginase 1 (ARG1) is constitutively expressed in the cytosol of hepatocytes and is also induced
in myeloid cells in response to various stimuli such as Th2 cytokines and TGF-β 109, 110.
Induction of ARG1 by Th2 cytokines was also observed in bone marrow-derived DCs 111.
Arginase2 is a mitochondrial enzyme with wide tissue distribution. It is expressed in kidney,
lactating mammory gland, prostate, brain and small intestine. Constitutive expression of ARG2
was also observed in bone marrow derived macrophages, but was not up-regulated by ARG1
inducers 111.
16
NOS has three isoforms, two of which are constitutively expressed (neuronal and endothelial
NOS) and one which is inducible (iNOS). Here, only iNOS is described. iNOS is expressed in
cells of the immune system (including macrophages and DCs) 112 upon induction with Th1
cytokines, IFN-β and TNF 109-111. Both ARG1 and iNOS are induced by LPS 109, 110. In addition
to reciprocal regulation of iNOS and ARG1 by Th1 and Th2 cytokines respectively, these two
enzymes also negatively cross-regulate each other 109, 110. NG-hydroxy-L-arginine (NOHA)
released as a by-product of iNOS enzymatic activity during L-arginine metabolism inhibits ARG
1 and stimulates surrounding immune cells to produce NO by iNOS. This differential regulation
and enzyme production has been used to differentiate classically and alternatively activated
macrophages. Classically activated macrophages release pro-inflammatory cytokines, such as IL-
1, IL-6 and TNF as well as reactive oxygen and nitrogen species as byproducts of iNOS activity,
which, in turn, leads to its anti-microbial action 110. On the other hand, alternatively activated
macrophages secrete IL-10 and up-regulate ARG1 activity. These cells are responsible for tissue
repair and fibrosis.
The effect of NO, which is produced by iNOS, is microbicidal 107. iNOS KO mice were shown to
be more susceptible to L. major bacterial infection than heterozygous and WT mice 113. iNOS
activity also has potent immuno-suppressive effects. Although iNOS consumes L-arginine, its
immunosuppressive properties have been shown to be unrelated to L-arginine starvation of T
cells123. Rather, it has been noted that by-products of iNOS activity such as reactive nitrogen
species suppress T cell proliferation 114, 115. T cells from iNOS KO mice displayed higher levels
of proliferation and IFN-γ production but less IL-4 production in response to Leishmania Ag or
concanavalin A 113. The mechanism of suppression involved T cell cycle arrest through
impairment of IL-2R signalling (inhibited phosphorylation of STAT5, JAK3, Erk1/2 and Akt),
although IL-2R chain expression remained normal 116, 117. T cell suppression was reversible only
during first 24 hours of culture but not at later time points. iNOS was observed to be induced by
IFN-γ. However, blocking IFN-γ reversed suppression by about half, suggesting that other
mechanisms are involved in iNOS upregulation. Suppression of T cell responses also required
cell contact 116, 117. Induction of cell death was not observed in co-cultures 116, 117. Moreover, it
was observed that alveolar macrophages were able to suppress the stimulatory capacity of DC
via NO production by iNOS 118. The effect was reversed by inhibiting iNOS.
17
The role of NO in tumour killing has been recognized 107, 108. It was first observed that primary
mouse macrophages could kill mouse tumour cell lines 119, an effect that was mimicked by
addition of NO and abolished with the addition of an iNOS inhibitor and in iNOS KO
macrophages 108 119. An anti-tumour effect of microbe-induced NO production was also
observed in vivo 120. Mice that received bacillus Calmette-Guerin (BCG) bacterium ip, had
reduced number of ovarian tumour cells transplanted ip and no evident ascites in the peritoneum
as compared to mice that did not receive the bacterium injection. The effect was completely
reversed with co-administration of iNOS inhibitor and anti-IFN-γ Ab 121.
High ARG1 activity has been observed in patients with various malignancies 122. ARG1 activity
in tumor-associated macrophages (TAMs), MDSC and some types of tolerogenic DCs was also
shown to be immuno-suppressive in multiple studies. Tumor-derived mature myeloid cells
(identified as macrophages) were observed to be the source of ARG1 in 3LL murine lung
carcinoma. These cells also produced IL-10, IL-1 and IL-6 122 and suppressed T cell proliferation
in vitro by down-regulating the CD3ζ chain of the T cell receptor complex 123, 124. Another study
demonstrated down-regulation of both CD3ε and CD3ζ chains and inhibition of ARG1 in TAMs
restored T cell proliferation 122. An alternative mechanism of suppression mediated by ARG1 is
thought to be associated with inhibition of cell cycle progression and absence of cyclin D3 and
cdk4 expression – enzymes responsible for cell cycle progression 125.
Paradoxically, arginase and iNOS seem to be co-upregulated in some cell types 107. Several
studies have shown that tumour-derived MDSC 126 and certain types of DC-regs suppress T cell
proliferation in vitro via both arginase and iNOS-mediated L-arginine depletion or reactive
oxygen species generated as a by-product of these two enzyme activities. The relevance of iNOS
and arginase activity in reactive oxygen species production can also be appreciated in the context
of resolution of the immune response. In this setting, contraction of T cell response takes place,
possibly due to the mechanisms mentioned above 107.
5.3 Indoleamine 2,3-dioxygenase (IDO)
IDO is an intracellular enzyme expressed in many tissues. A role of IDO has been described in
tumor progression 128, T cell tolerance to tumors, inhibition of T cell proliferation both in vitro
18
127, 133 and in vivo 133 and as negative regulator of immune disorders. It was also observed that
over-expression of IDO results in immuno-suppression and tolerance 130.
IDO is part of the innate immune defence against pathogens. IDO metabolizes tryptophan to
yield its degradation by-products known as kynurenines. Some microorganisms depend on
exogenous tryptophan as a source of this essential amino acid. Limiting trypthophan in the
microenvironment by induction of IDO, therefore, acts as a microbicidal strategy of the innate
immune system 130. Constitutive IDO expression occurs at maternal-fetal interface as well as in
mouse gut, lymph nodes, spleen, thymus and gut130. Moreover, IDO expression exists in multiple
primary human tumors 128. Myeloid-lineage cells such as monocytes, macrophages and DCs
express IDO after exposure to IFN-γ 129, LPS 130 and CD40L or to a combination of these
molecules 127. Only certain DC subsets seem to be able to express IDO. These subsets include
CD8a+ cDCs and B220+ pDCs 131 and are termed ‘IDO competent DCs’. IDO-expressing pDCs
were identified in tumor-draining LN 132. By contrast, pro-inflammatory signals that trigger DC
maturation also down-regulate IDO production, whereas certain tolerogenic signals (eg. ligation
of CD80/CD86 by inhibitory receptor CTLA-4 131) up-regulate IDO in DCs.
The immuno-regulatory role of IDO was demonstrated in vitro when it inhibited proliferation of
tumor cells by consumption of amino acid tryptophan 129. This effect was mediated through
kinase GCN2, which triggers cell cycle arrest 133. Furthermore, one study found that T cell
unresponsiveness could be induced in response to the metabolites L-kynurenine and picolinic
acid produced as a by-product of IDO activity 134.
5.4 IL-10
IL-10 is an immuno-regulatory cytokine that prevents auto-immune and other inflammatory
pathologies. IL-10 or IL-10R KO mice do not develop systemic auto-immune disease but
develop colitis in the presence of microorganisms 135. IL-10 is widely expressed by cells of the
immune system. Its expression has been demonstrated in multiple subsets of T cells (such as
Th1, Th2, Th17, Tregs) as well as DCs, macrophages, mast cells, eosinophils, NK cells and
neutrophils 135. In macrophages and DCs, IL-10 can be induced by TLR ligands including
TLR2, TLR4, and TLR9 136. DC-SIGN and Dectin-1, which ligate other PRR, can also stimulate
IL-10 release from DCs 137.
19
IL-10 inhibits IFN-γ production by Th1 cells and drives T cell responses toward a Th2
phenotype 138. In another report it was demonstrated that IL-10 inhibits both proliferation and
cytokine production by both Th1 and Th2 cells 139 140. Moreover, when CD4+ T cells are
activated in vitro, IL-10 causes them to convert into regulatory Tr1 phenotype 140. The inhibitory
effect of IL-10 on T cell proliferation is mediated partly though inhibition of APC function 139.
IL-10 was demonstrated to suppress APC function of monocytes by reducing expression of
MHC 141 and co-stimulatory molecules 142, as well as reduce the release of pro-inflammatory
cytokines such as IL-12 143, TNF, IL-1β, IL-6 and GM-CSF 144. Additionally, IL-10 enhances
the release of soluble TNF-αR and IL1-βR antagonist that act in an anti-inflammatory fashion 145.
IL-10 can inhibit monocyte differentiation into DC and promote their differentiation into
macrophages 146. The immunosuppressive function of IL-10 has been shown to be mediated, at
least partly, by STAT3 signalling downstream of IL-10R 147.
6 Antigen presenting cells in cancer immunology
In addition to their importance in the maintenance of tolerance in the steady state, DCs, together
with other types of APCs, contribute to poor immune responses in various pathological
conditions. In cancer, it is thought that an immunosuppressive tumor microenvironment drives T
cell hyporesponsiveness and tolerance towards tumor Ag. Below, I describe key tumor-
associated APCs and their relevance to tumor tolerance.
6.1 Tumor-derived dendritic cells
Immune recognition of tumor antigens is thought to be mediated by tumor DCs that have
migrated into secondary lymphoid tissues. There, tumor DCs prime T cells, which then return to
the tumor to kill tumor cells. 148. For most patients, however, this process appears to be
ineffective, in part because of DC dysfunction at various levels. The first evidence for the
impairment of DC function in cancer came from observations that patients with advanced tumors
have reduced number of DCs in their blood 149. Decreased numbers of mature DCs were also
observed in spleen, lymph nodes and tumor in tumor-bearing mice 148. Similarly, DC
recruitment to tumors was impaired in a wide variety of primary tumors 150, 151, which correlated
with poor patient prognosis 152. Tumor-infiltrating DC expressed low levels of MHCII and co-
20
stimulatory molecules 149 153 154, failed to respond to maturation stimuli 155, and primed T cells
poorly 149 153. Increased numbers of immature DCs have been detected in peripheral blood of
cancer patients, indicating that DC dysfunction can be systemic149. DC maturation was also
inhibited when DCs were cultured with tumor-conditioned medium in vitro 149 153 156,157.
Immature tumor DCs suppress T cell responses and lead to unresponsiveness toward tumor Ag 158. In other reports, an increased rate of DCs apoptosis was detected in tumors 159.
Collectively, these factors result in reduced activation or direct suppression of T cell responses.
Tumor DCs arise from pre-cDCs that migrate into the tumor via a CCL3-dependent mechanism 160. Once in the tumor, DC differentiation is influenced by the intra-tumoral inflammatory
milieu, which has been shown to alter DC differentiation 161. Studies in our laboratory describe a
high proportion of GR-1+ DCs both in the tumor itself (up to 35% of total tumor DCs), in
draining lymph nodes and in spleen of tumor-bearing mice 161. Moreover, the frequency of GR-
1+ DCs in lymphoid tissues correlated directly with tumor size 161. These cells were defective in
priming T cells in allogeneic MLRs and had a reduced expression of MHCII and CD86 after
maturation. IL-10 was implicated in the defective T cell stimulation ability.
A variety of molecules in the tumor microenvironment have been implicated in the impairment
of tumor DC including IL-10, IL-6, VEGF, M-CSF and prostanoids 146. IL-6 inhibits DC
differentiation both in vitro and in vivo 162. VEGF has been associated with reduction in DC
number and accumulation of immature myeloid cells in tumor bearing mice and patients156 163.
Anti-VEGF Ab treatment increased the number of DC in spleen and LN of tumor-bearing mice
and increased the ability of the DCs to prime T cells156 163. A recent study suggested that lipid
accumulation in DC may contribute to DC dysfuntion in cancer patients 164.
6.2 Myeloid derived suppressor cells (MDSC)
MDSC are a heterogenous population of neutrophils, monocytes, and primitive myeloid cells that
increase in frequency in bone marrow, blood, and lymphoid tissues of tumor-bearing mice and
patients149 165 169. In mice, they are typically defined by the expression of CD11b and GR-1 107;
however, these markers are non-specific and cell populations expressing CD11b and GR-1 occur
in normal mice albeit at much lower frequencies (less than 1% of total circulating cells; 2-4% of
spleen cells; and up to 50% of bone marrow cells) 148 166. Further, these cells differentiate into
21
mature functional myeloid cells in normal mice 167. Recent studies in tumor-bearing mice
suggest that MDSC can be subdivided into CD11b+ Ly6c+ Ly6Glow monocyte-like and CD11b+
Ly6Clow Ly6G+ granulocyte-like cells that suppress immune responses through different
mechanisms 170 169. Monocytic MDSC produce higher levels of NO and induced substantially
elevated levels of tyrosine nitrosylation than granulocytic MDSC, which produce increased
levels of ROS 169. In other studies, the expresson of CD115 (M-CSFR) and CD124 (IL-4-Ra)
has been used to define MDSC subsets 169.
Immature myeloid CD11b+ cells or MDSC isolated from tumors were shown to be potent
suppressors of Ag-specific T cell proliferation 168 and function 171 both in vitro and in vivo 172.
MDSC from peripheral organs suppress T cell responses in an Ag-specific manner, whereas
MDSC from tumor sites mediate Ag-non-specific suppression 126. However, MDSC isolated
from the spleen of tumor-bearing mice showed a mixed suppressive activity 173 and were not
suppressive in some studies 126. It was observed that MDSC did not engage TCR or activate T
cells 171 but caused dissociation of TCR complex with the CD3ζ chain and with CD8 in OTI
cells 171 176. MDSC have also been shown to inhibit IL-12 production by macrophages in an IL-
10-dependent fashion 174.
Most studies have reported that the suppressive effects of MDSCs are cell-contact dependent.
Activation of MDSC results in upregulation of ARG1 and iNOS enzymatic activity and
increased production of ROS and NO via STAT3175. High ARG1 and iNOS activity are
considered the main mechanisms by which MDSC suppress immune functions 126 177 116. In
particular, production of peroxynitrite, a byproduce of iNOS, results in nitration of the TCR and
CD8, which renders T cells unresponsive to Ag-specific stimulation 171. Some reports found
suppression to be IFN-γ-dependent 116, 170.
6.3 Tumor associated macrophages (TAMs)
Evidence for the role of macrophages in tumor progression has accumulated over decades.
TAMs are derived from blood circulating monocytes that are recruited to tumors by CCL2 as
well as CCL5, CCL7, CCL8, CXCL12, VEGF, PDGF and M-CSF produced within tumor
microenvironment 178 179 180. Multiple studies have reported that high numbers of TAMs
correlates with poor outcome in many human cancers 178. M-CFS is the main growth factor
22
responsible for survival, proliferation, differentiation and chemotaxis of cells of mononuclear
phagocyte system. High levels of M-CSF correlate with poor prognosis in patients with cancer 181.
In established tumors, TAMs have a characteristic M2 macrophage phenotype 182. Under normal
physiological conditions, M2 cells promote wound healing, tissue remodelling, angiogenesis, and
suppression of immune responses. In tumors, these cells produce low amount of IL-12 and high
amounts of IL-10, TGF-β and arginase, which act in immunosuppressive fashion. TAMs
preferentially recruit naïve T lymphocytes devoid of cytotoxic function as well as Th2 and Treg
cells to tumors via CCL18 183, CCL17 and CCL22 respectively 184 185. Direct T cell immuno-
suppressive function of TAMs was reported to involve PD-L1 ligand 186.
Analogous to the wound interior, tumors exhibit a highly hypoxic microenvironment. Hypoxia
up-regulates HIF-1α 187 and HIF-2α 188 expression in TAMS resulting in increased VEGF
production, which stmulates tumor angiogenesis 189. M-CFS has also been shown to stimulate
VEGF release from TAMs 190. TAMs produce other pro-angiogenic factors such as PDGF, TNF,
and CXCL8 178. TAMs can also be found in vascular areas inside the tumor 191, where they
enhance nutrient and oxygen supply for tumor growth 192. The production of metalloproteases
(MMP2, MMP9) by macrophages, a component of tissue remodelling, promotes cancer invasion
and metastasis 193.
23
CHAPTER 2
Statement of the Problem
1 Preliminary observations
Conventional DCs (cDCs) arise from pre-cDCs, an immediate precursor population originally
identified in bone marrow by Diao et al 62. When placed on a stromal monolayer in the presence
of GM-CSF, pre-cDCs differentiate into a homogenous population of proliferating cDCs that
continue to divide over 10-12 days (Figure 3A). In the absence of GM-CSF, pre-cDCs also
generate cDCs initially, but by day 9 the cells lose surface expression of both CD11c and
MHCII, the classic hallmarks of cDCs (Figure 3B). The morphology of the cells also changes
(Figure 3c): cDCs appear as loosely adherent clusters with cells displaying motile denditic
processes; cells generated in the absence of GM-CSF are round with few dendrites and appear
tightly adherent or embedded in the monolayer.
The development of CD11c- MHC II- cells from CD11c+ MHCII - cDC was unexpected as it
is generally believed that cDCs are terminally differentiated. To further confirm that CD11c-
MHCII - cells arose from proliferating cDCs, CD11c-Cre+ Rosa26-EGFP transgenic mice and
their CD11c-Cre- littermate controls were used to trace the life history of CD11c- cells back to
CD11c+ progenitors. Cre recombinase, driven by the CD11c promoter, deletes the stop codon
for ROSA-GFP. Cells that express CD11c are permanently tagged by GFP irrespective of
subsequent CD11c expression levels in the progeny. Before culture, GFP could be detected in
about 10-20% of pre-cDC from CD11c-Cre+/- Rosa26-EGFP transgenic mice (Figure 4). Cre+/-
pre-cDCs differentiated into MHCII+ cDC at day 3 concurrent with the up-regulation of GFP
expression. At day 10, when the Cre+/- cells had lost CD11c and MHCII expression, GFP
expression persisted.
Phenotypic characterization of CD11clow MHCII low cells revealed absence of most
phenotypic markers characteristic of DC subsets (Figure 5). The cells stain negative for CD11c,
MHCII, CD4, CD8, CD103 and GR-1. However, the cells express high levels of CD11b and
CD172a. Moreover, contrary to cDCs, these cells are resistant to maturation stimuli such as LPS
24
and fail to up-regulate MHCII and co-stimulatory molecules (Figure 6). Additionally, these cells
were observed to be poor stimulators of T cell proliferation in allogeneic mixed lymphocyte
reaction (Figure 7). Although these cells are poor T cell stimulators, they demonstrated an Ag
uptake capacity that was comparable to that of DCs (Figure 8).
2 Hypothesis
Previous studies suggest that DC with low expession levels of MHCII and co-stimulatory
molecules are immunosuppressive and promote immunologic tolerance77. Based on preliminary
investigations of pre-cDC-derived CD11clow MHCII low cells, we hypothesize that they have
immuno-suppressive properties in vitro and in vivo.
3 Objective
To characterize the functional properties of DC-derived CD11clow MHCII low cells
4 Specific aims
Aim 1: To investigate the functional and immunosuppresive properties of CD11clow
MHCII low cells in vitro.
Aim 2: To define the mechanisms by which CD11clow MHCII low cells promote unresponsiveness.
Aim 3: To evaluate the in vivo effects of CD11clow MHCII low.
25
CHAPTER 3
Methodology
1 Mice
C57BL/6, Balb/c, C57BL/6.SJL congenic and OT-II OVA (323-339)-specific TCR transgenic
mice (B6.Cg-Tg(TcraTcrb)425Cbn/J) were purchased from the Jackson Laboratory. Mice were
maintained in pathogen-free conditions in accordance with institutional guidelines, and used at 6-
8 weeks of age.
2 Primary skin stromal cell preparation
New born skin stroma was prepared from newborn C57BL/6 mice. The skin of new born mice
was minced and cultured at 37ºC in 10 cm plates containing DMEM medium supplemented with
10% FBS, penicillin (50 U/mL) and streptomycin (50 ug/mL). After 1 week, when the cells had
formed a confluent monolayer, the cells were treated with 0.25% trypsin/1mM EDTA, split, and
passaged three times. Immediately prior to co-culture with pre-cDCs, the confluent monolayer
was irradiated (25 Gy).
3 Cell Isolation
DCs were isolated from spleens of C57BL/6 mice, which were inoculated previously with a Flt3
ligand-producing B16 melanoma cell line (B16-Flt3L). Spleens were minced and digested with
collagenase D and DNase for 0.5 hours at 37ºC. Cells were passed through a 0.42 µm nylon
mesh and subjected to density gradient centrifugation using Nycodenz 194. Nycodenz was
prepared by using 7:7:16 v/v/v Nycoprep (Cederlane Laboratories), tricine and PBS with 5%
FBS and 2mM EDTA (binding buffer). Dendritic cells were further enriched for CD11c+ cells
by positive selection using MACS (Milteniy Biotech) CD11c+ immuno-magnetic beads. Cells
were washed with binding buffer during all steps. Cells were subsequently cultured overnight
before use at a density of 1×106 cells/mL in RPMI supplemented with 10% FBS, 50 uM 2-ME,
26
1mM sodium pyruvate, 10 mM non-essential amino acids, 50 U/mL penicillin and 50 ug/mL
streptomycin (complete medium) in the presence of GM-CSF (4 ng/mL; BD Pharmingen).
Pre-cDCs were isolated from bone marrow of B16-Ftl3L melanoma treated C57BL/6 mice.
Femurs and tibia were flushed with binding buffer and subjected to Lympholyte-M (Cederlane
Laboratories) density gradient centrifugation. Cell suspensions were layered on top of
Lympholyte-M at ratio of 2:1 v/v. CD11c+ bone marrow precursors were further enriched by
MACS (Milteniy Biotech) CD11c+ immuno-magnetic beads. Cells retained in the column were
eluted and labelled with anti-I-Ab-PE, anti-CD11c-APC and anti-lineage makers (anti-CD3-,
anti-CD19-, anti-B220-, anti-CD49b-FITC) mAbs. Lin- CD11c+ MHCII - pre-cDC were isolated
using MoFlo High Speed Cell Sorter using Summit acquisition and analysis software
(DakoCytomation). The purity of the pre-cDCs population used was routinely ≥99% based on
reanalyzed samples.
4 Flow cytometry
Flow cytometry was performed on Beckman FC500 using CXP Analysis software (Beckman
Coulter). Prior to staining, cell suspensions were pre-incubated with anti-CD16/32 in binding
buffer to block FcRs for 25 min at 4ºC. Cells were then washed with binding buffer and stained
with mAb conjugates for 25 min at 4ºC in a final volume of 100 ul of binding buffer with cell
density of not more than 5×106/100 ul. Appropriate isotype controls were included.
For intracellular staining, cells were pre-incubated with anti-CD16/32 in binding buffer for 25
min at 4ºC, surface stained as above, fixed and permeabilized using BD Cytofix/Cytoperm kit
(BD Biosciences) according to manufacturer’s instructions. Anti-IL-2-, anti-IL-4-, anti-IFNγ-PE
mAbs were used for intracellular staining.
5 Mixed lymphocyte reactions
CD4+ OT-II T cells or CD8+ OT-I T cells were isolated from the spleen and lymph nodes of OT-
II or OT-I transgenic mice, respectively. Tissues were minced, passed through a 42 µm nylon
mesh and subjected to density gradient centrifugation with Lympholyte-M. CD4+ or CD8+
27
lymphocytes were enriched using MACS CD4+ or CD8+ immuno-magnetic beads. Graded
numbers of stimulator cells (splenic DCs) cultured overnight with GM-CSF and/or suppressor
cells (CD11clow MHCII low cells derived from pre-cDC) were seeded in triplicate in 96-well U-
bottom plates (BD Biosciences). Responder spleen cells (1×105/well) from BALB/c mice, CD4+
OT-II or CD8+ OT-I cells were added to the wells in total volume of 200 µl of RPMI 1640
complete medium. 10 µg/ml IL-10R Ab, 500 µM N ω-hydroxy-nor-Arginine (Nor-NOHA) and
200 µM N6-(1-iminoethyl)-L-lysine, dihydrochloride (L-NIL) were used to block IL-10R,
arginase I and iNOS, respectively. Cells were cultured in humidified atmosphere of 5% CO2 in
air at 37 ºC. Cultures were pulsed with 1 µCi of [H3]-thymidine (Amersham) 16 hours before
harvest and collected into glass fiber filters (Millipore). [H3]-thymidine incorporation was
quantified using Beckman scintillation counter. Results are expressed as mean cpm of triplicate
cultures.
6 CFSE labelling
Isolated cells were washed twice with PBS and stained with 1uM CFSE (Molecular Probes) for
15 min at 37 ºC. Cells were then washed twice with PBS.
7 Reverse transcriptase PCR
Total RNA was extracted from DCs and DC-regs with TRIzol (Invitrogen Life Technologies) as
per the manufacturer’s instructions. RNA was resuspended with RNase free water and treated
with DNase I (Invitrogen Life Technologies) to remove contaminating genomic DNA. RNA was
then reverse transcribed using M-MLV Reverse Transcriptase (Invitrogen Life Technologies)
and amplified by PCR using the following primers: murine PD-L1, sense: 5’-
GTGAAACCCTGAGTCTTATCC-3’, anti-sense: 5’-GACCATTCTGAGACAATTCC-3’; IDO,
sense: 5’-GTACATCACCATGGCGTATG-3’, anti-sense: 5’-
GCTTTCGTCAAGTCTTCATTG-3’; arginase1, sense: 5’-
CAGAGTATGACGTGAGAGACCAC-3’, anti-sense: 5’-
CAGCTTGTCTACTTCAGTCATGGAG-3’; iNOS, sense: 5’-
AGCTTCTGGCACTGAGTAAAGATAA-3’, anti-sense: 5’-TTCTCTGCTCTCAGCTCCAAG-
28
3’; FasL, sense: 5’-AACCCCAGTACACCCTCTGAAA-3’, anti-sense: 5’-
GGTTCCATATGTGTCTTCCCATTC-3’; TGFβ2, sense: 5’-
TGGCCGCCTGGAGCAAGAAA-3’, anti-sense: 5’-AAGCGGCTGGGGGATGAC-3’; IL-10,
sense: 5’-GGATCTTAGCTAACGGAAACAACT-3’, anti-sense: 5’-
AAGCGGCTGGGGGATGAC-3’; ICOS-L, sense: 5’-CTTGGTCTGTTCTTGCTGCTG-3’,
anti-sense: 5’- GGCTATTGTCCGTTGTGTTG-3’.
8 Arginase and iNOS activity assays
To determine nitrite production by DC-regs and DCs, 1×106 cells were pulsed with LPS (2
µg/mL) overnight at 37 ºC in RPMI 1640 complete medium and supernatants harvested. Nitrites
were quantified in supernatants using Griess Reagent Kit for Nitrite Determination (Invitrogen),
according to manufacturer’s instructions.
Arginase activity was measured in cell lysates of cells previously pulsed with LPS. After
overnight culture with LPS, cells were digested with 0.25% trypsin/2mM EDTA and washed
twice with PBS. Cells were lysed in 100 uL 0.1% Triton X-100 126. To 100 uL of protein lysate,
100 uL of 25 mM Tris-HCl and 10 uL of 10 mM MnCl2 were added and enzyme was activated
by heating for 10 min at 56ºC. Arginine hydrolysis was conducted by incubating the lysate with
200 uL of 0.5 mM L-arginine, pH 9.7, at 37ºC for 60 min. The reaction was stopped with 900 uL
of H2SO4 (96%)/H3PO4 (85%)/H2O (1/3/7, v/v/v). 40 uL of 9% beta- isonitrosopropiophenone
dissolved in 100% ethanol was then added and the reaction mixture was incubated at 95ºC for 25
min. Urea concentration was determined by measuring absorbance at 562 nm. One unit of
enzyme activity is defined as the amount of enzyme that catalyzes the formation of 1 µmol urea
per minute.
9 Adoptive transfer studies
OVA-pulsed CFSE-labelled cDCs or DC-regs (1×106) were injected i.p. into mice that had
received 1×106 CFSE-labelled OT-II CD4+ cells 24 hours earlier. Spleen and lymph nodes were
collected 3 and 5 days later. T cell proliferation was assessed by CFSE dilution. T cell function
29
was assessed by pulsing spleen and lymph node cells with OVA peptide in vitro and assessing
cytokine production 12 hours later.
10 Statistics
Continuous variables are expressed as mean±SE and were analyzed by two-tailed Student t test.
A P value below 0.05 was considered statistically significant.
30
CHAPTER 4
Results
1 CD11clow MHCIIlow cells exhibit potent immuno-suppressive properties in vitro
It was demonstrated previously that pre-cDCs generate proliferating cDC 62. In this study, pre-
cDCs were placed on a stromal monolayer and their proliferation was observed. On stroma
supplemented with DC growth factor GM-CSF, pre-cDCs generated a highly pure population of
cDCs by day 3. In the absence of GM-CSF, pre-cDCs also generated cDCs by day 3 (Figure
3A). However, when cDCs were cultured further in the absence of GM-CSF (up to day 14), loss
of both CD11c and MHCII expression, which normally characterize DCs, was observed.
Moreover, the cells acquired distinct morphology. In contrast to DCs, which appeared as small
irregular-shape cell clusters loosely attached to the monolayer, these cells were evenly
distributed through the stroma. The new cells appeared much larger than DC and had smooth
round shape without dendrites (Figure 3B).
Incubation of CD11clow MHCII low cells with LPS failed to upregulate co-stimulatory molecules
and MHCII (Figure 6), indicating that these cells were resistant to maturation. CD11chi MHCIIhi
cDC cells generated in the presence of GM-CSF efficiently stimulated T cell proliferation in
allogeneic MLR. In contrast, CD11clow MHCII low displayed very poor T cell stimulatory
capacity even when pulsed with TNF-α, another maturation stimulus (Figure 7).
DC with low expression of co-stimulatory molecules and poor T cell priming capacity in
allogeneic MLR were previously shown by Sato et al to have T cell regulatory activity. 77.
Svensson et al. showed that CD11clow CD45RB+ DCregs cells generated from bone marrow
progenitors on stroma in the absence of GM-CSF also had poor T cell priming capacity and had
potent immuno-suppressive properties98. It was, therefore, hypothesized that
CD11clow MHCII low cells generated from pre-cDC have the ability to actively suppress T cell
responses.
31
To test whether CD11clow MHCII low cells have suppressive properties, various numbers (0.25-
2x104) of CD11clow MHCII low cells were added to allogeneic mixed lymphocyte cultures
containing 1×105 BALB/c spleen cells and 1×104 CD57BL6 spleen cDCs. Addition of
CD11clow MHCII low cells suppressed T cell proliferation in a dose-dependent fashion (Figure 9).
When added at a ratio of 1:1 with stimulatory DCs, CD11clow MHCII low cells were able to
suppress T cell response by 75%. T cell proliferation was decreased to baseline when
CD11clow MHCII low cells were added at higher ratios. Because of this potent suppressive
property, it was decided to refer to CD11clow MHCII low as DC-regs.
Although DC-regs suppressed T cell proliferation, it was still possible that T cells were activated
and capable of mounting an effector response212. Therefore, the ability of T cells previously
primed with allogeneic DC-regs to produce cytokines after secondary re-stimulation was tested.
T cells were recovered from primary cultures with DC-regs and re-stimulated with anti-CD3 and
anti-CD28 Abs. Intracellular cytokine production was measured by flow cytometry. T cells that
were pre-incubated with DC-regs with or without DCs did not produce IL-2, IFN-γ or IL-4
(Figure 10). These results demonstrate that DC-regs are able to inhibit both allogeneic T cell
proliferation and activation, which was not reversed despite the absence of DC-regs in the
secondary MLR.
To determine if DC-regs suppressed Ag-specific T cell responses, TCR-transgenic CD4+ T cells
purified from spleens of OT-II mice, which are specific for OVA 323-339 peptide, were used as
responders. Splenic DCs pulsed with OVA protein and matured with GM-CSF overnight were
mixed at 1:1 ratio with DC-regs that were or were not pulsed with OVA overnight. OT-II cells
were then incubated with DC-OVA ± DC-reg / DC-reg-OVA or DC-reg / DC-reg-OVA alone.
The number of OT-II cells was measured on day 3 by flow cytometry (Figure 11). As expected,
DC-OVA stimulated proliferation and expansion of OT-II T cells. DC-regs alone did not induce
T cell proliferation regardless of whether they were pulsed with OVA or not. When OT-II cells
were pulsed with a combination of DC-OVA and DC-regs or DC-regs-OVA, T cell proliferation
was suppressed almost to baseline.
32
It was possible that the low T cell number observed was due to excessive proliferation and cell
death rather than suppressed proliferation. To exclude this possibility, CFSE-labelled OT-II cells
were analyzed after stimulation with various combinations of OVA-pulsed and control-pulsed
DC and DC-regs. These studies confirmed that DC-reg inhibited OT-II cell proliferation (Figure
12). Stronger suppression of proliferation occurred when DC-regs were not pulsed with OVA.
In contrast to the allogeneic system, DC-regs failed to suppress cytokine production in OT-II
primed with DC-OVA (Figure 13). In these studies, T cells were cultured for 7 days99 before re-
stimulation with anti-CD3 and anti-CD28 mAbs.
2 Mechanisms of immuno-suppression mediated by CD11clow
MHCIIlow DC-regs
There are several possible mechanisms that could account for suppression of T cell responses by
DC-regs:
1) DC-regs may induce production of Tregs. Previous studies have shown that peripheral
tolerance induction involves DC-regs or immature DCs and their ability to promote
generation of Tr1 and FoxP3+ Tregs 36, 92, 94, 99.
2) DC-regs may inhibit T cell cyling through the release of various metabolites. Inhibition
of T cell responses has been observed previously in conditions of L-arginine starvation
induced by arginase1-expressing cells including tumor-derived DCs, TAMs and MDSC.
High expression of arginase1 results in down-regulation of one of the TCR chains
(CD3ε122 or CD3ζ123, 124) which can be reversed with arginase1 inibition or addition of
excess L-arginine to the culture medium. Moreover, arginase1 was also reported to
inhibit cell cycle progression via suppression of cyclin D3 and cdk4 expression125.
Tryptophan starvation as a result of high IDO expression by certain subsets of DCs has
also been reported to inhibit T cell proliferation131, 133. Moreover, high levels of reactive
nitrogen species produced as a by-product of iNOS activity can induce cell cycle arrest
via inhibition of IL-2R signalling 116, 117. Suppression of T cell proliferation mediated by
iNOS was reported to be independent of its L-arginine consumption123.
33
3) DC-regs may induce T cell anergy by ligating T cell inhibitory receptors including
ICOS196 or PD-1/PD-2195. Ligation of the ICOS receptor on T cells also leads to IL-10
production, a well-recognized immunosuppressive cytokine196.
4) DC-regs may induce T cell death 90.
To establish the mechanism of immuno-suppression in our system, the activation status of T cells
in allogeneic MLR was assessed first by measuring expression levels of CD25 and CD44.
BALB/c T cells incubated with C57BL/6 DCs showed high expression of both markers at day 3
(Figure 14). By contrast, co-culture with C57BL/6 DC-regs blocked CD25 and CD44
expression. Similarly, no T cell activation was detected in cultures of T cells incubated with DC-
regs alone. These findings indicated that DC-regs suppress T cell activation.
It was next assessed whether DC-regs induced T cell apoptosis. OT-II cells were recovered from
cultures and stained with propidium iodide (PI). In necrotic or late apoptotic cells, in which cell
membrane integrity is compromised, PI can permeate into the cell and intercalate between DNA
bases; this results in increased fluorescence (20-30 fold). PI is effectively excluded from viable
cells. In the absence of stimulation, most T cells stained positive for PI (Figure 15). Similarly, T
cells incubated with DC-regs alone stained positive for PI indicating that DC-regs do not provide
sufficient T cell survival signals, which may be related to their low MHCII expression (Figure 5)
and, therefore, the inability to ligate the TCR complex on OT-II cells. Few dead cells were
detected when T cells were incubated with both DCs and DC-regs as DCs provide TCR
stimulation vital for T cell survival. These results indicate that DC-regs in co-cultures with cDC
do not actively induce T cell death or prevent survival signals from reaching T cells.
It was hypothesized that CD4+ CD25+ FoxP3+ Tregs might be induced by DC-regs. To test this,
OT-II cells were incubated with DC-OVA +/- DC-reg-OVA or DC-reg. After 7 days, OT-II cells
were recovered, stained for CD4, CD25 and FoxP3, and analyzed by flow cytometry. We did not
detect CD4+ CD25+ FoxP3+ Tregs present under any of the culture conditions (Figure 16).
Collectively, these results suggested that suppression of T cell proliferation and activation by
DC-regs does not involve FoxP3+ CD25+ Tregs, or regulation of T cell survival or death. Instead,
it seemed likely that DC-regs produce factors that regulate T cell activation and cell cycle
progression.
34
Trans-well experiments were conducted to determine whether suppression mediated by DC-regs
was cell contact dependent. OT-II cells (0.5×106) and DC-OVA cells (1×105) were placed in the
bottom chamber of the trans-well and DC-reg-OVA (1×105) were placed in the top chamber. The
chambers were separated by a membrane whose 8 µm pore size excluded DC-regs from the
bottom chamber. DC-reg suppression was reduced by about half in the transwells, suggesting
that both soluble and membrane bound mediators were involved (Figure 17). Passage of cell
processes through membrane pores was possible but unlikely, as DC-regs did not have dendritic
processes (Figure 3B).
Qualitative RT-PCR was used next to determine whether DC-regs express previously recognized
immunosuppressive molecules including IL-10 148, IDO 127, PD-L1 195, ICOS-L 196, iNOS 126,
arginase1 126 and FasL 197. Erythrocyte-depleted spleen cells were used as positive control for all
assessed molecules. FasL expression was not observed in DC-regs (Figure 18). This was in
agreement with the previous finding that DC-regs did not induce T cell death (Figure 15). IDO
expression was not detected in DC-regs, indicating that tryptophan starvation is likely not
involved in DC-reg mediated T cell suppression127. ICOS-L, PD-L1 and IL-10 were
constitutively expressed by DC-regs, whereas arginase1 and iNOS were induced upon LPS
stimulation (Figure 18).
The activity of ARGI and iNOS enzymes in DC-regs was determined with biochemical assays.
ARG1 activity was determined by incubating cell lysate with L-arginine and measuring urea
production. iNOS activity was determined by measuring nitrite production in the supernatant,
which is a by-product of iNOS activity. Consistent with the RT-PCR data, DC-regs displayed
high levels of both enzymes after overnight stimulation with LPS (Figure 19). By contrast,
freshly isolated spleen DCs did not display any enzymatic activity with or without LPS
stimulation.
Expression of PD-L1 was also assessed by flow cytometry (Figure 20). Both spleen DCs and
DC-regs showed high levels of PD-L1 expression whether or not they were incubated with LPS.
The high PD-L1 expression level in both cell populations suggests that PD-L1 is not involved in
DC-reg-mediated suppression.
35
IL-10 production was measured by ELISA (Figure 21). High levels of IL-10 were detected in
DC-regs pulsed with LPS, but not by DC. In contrast to RT-PCR data, IL-10 was undetectable
in DC-regs without LPS stimulation. This may indicate storage of intracellular IL-10 and its
release upon stimulation.
Collectively, these results demonstrate that suppression of T cell proliferation and activation is
most likely mediated by iNOS, arginase, and/or IL-10. PD-L1 and ICOS-L that were expressed
by DC-regs were also observed in DCs, which makes them unlikely candidates for DC-reg-
specific immuno-suppressive function.
To clarify the functional relevance of PD-L1, IL-10, ARG1 and iNOS, transgenic CD8+ OT-I T
cells were incubated with OVA-peptide pulsed spleen DCs (DC-OVA) in the presence or
absence of DC-regs and corresponding blocking reagents. Anti-IL-10R and anti-PD-L1 were
used to block binding of IL-10 to its receptor and PD-1, respectively. Nω-hydroxy-nor-Arginine
(Nor-NOHA) and L-NG-monomethyl arginine citrate (L-NMMA) were used to block arginase1
and iNOS enzymatic activity, respectively. Similar to our studies of OT-II cells, OT-I T cells
incubated with DC-OVA proliferated vigorously whereas the addition of DC-regs blocked
proliferation (Figure 22). Addition of anti-IL-10R Ab, anti-PD-L1, and and nor-NOHA had little
effect on immunosuppression. However addition of iNOS inhibitor L-NMMA restored T cell
proliferation by 60-70%. Similar results were obtained using the OT-II cells (data not shown).
These results indicate that DC-regs inhibit T cell proliferation through an iNOS-dependent
mechanism.
3 Immunosuppressive activity of CD11clow MHCIIlow cells in vivo
It has been reported that in vitro generated regulatory DC can suppress T cell proliferation in
vivo 77, 99. Immuno-suppressive properties of DC-regs were, therefore, tested in an adoptive T
cell transfer model. B6.SJL mice expressing the CD45.1 congenic marker were i.v. injected with
CFSE-labelled CD45.2+ CD4+ OT-II cells. Twenty-four hours later, OVA-pulsed DCs with or
without OVA-pulsed DC-regs were injected i.p. Proliferation of spleen OT-II T cells was
assessed at 3 and 5 days later by flow cytometery (Figure 23). OT-II T cells from mice injected
with DC and DC-regs showed more proliferation than mice injected with DCs alone. They also
36
produced more IFN-γ after re-stimulation with OVA peptide in vitro (Figure 23). By contrast,
OT-II T cells recovered on day 5 from mice receving DC-OVA + DC-regs showed lower levels
of proliferation and IFN-γ production than mice that received DC-OVA alone.
The ability of DC-regs pulsed with OVA to stimulate OT-II T cell proliferation on day 3 in vivo
was unexpected. It was speculated that DC-regs pulsed with OVA might have resulted in
transfer of OVA to immunostimlatory recipient APCs. To investigate this possibility, DC-regs
that were not pulsed with OVA were used in the adoptive transfer (Figure 24). However, DC-
regs were still unable to suppresss OT-II proliferation or IFN-γ production induced by OVA-
pulsed DC.
Because of these unexpected results, it was decided to assess the ability of DC-regs to prime T
cells in vivo. Here, mice were injected with either DC-OVA or DC-reg-OVA and proliferation
was assessed 3 days later. T cell activation markers were also assessed. T cells from both
groups displayed the same levels of proliferation. Moreover, both groups expressed similar
levels of CD44 and CD69 (Figure 25) indicating that DC-regs are able to prime T cells in vivo.
In summary, these results demonstrate that under the experimental conditions employed, DC-
regs fail to suppress OT-II T cell proliferation and activation.
37
CHAPTER 5
Discussion
This study demonstrates that cDCs retain functional and phenotypic plasticity. This novel
finding challenges the prevalent view that cDCs are terminally differentiated. cDC co-cultured
on a supportive stromal monolayer lost cell surface expression of CD11c and MHCII and
differentiated into a cell population with potent immuno-suppressive properties, which were
termed DC-regs. DC-regs have lost the usual constellation of morphologic and functional
features that are used to define cDC including morphology, responsiveness to maturaton stimuli,
and the capacity to stimulte lymphocytes in vitro. DC-regs also acquired the capacity to produce
high levels of iNOS, IL-10, and arginase. It was further shown that iNOS contributes to their
immunsuppressive properties.
Failure of previous studies to demonstrate this novel pathway of cDC development is multi-
dimensional. First, the culture conditions used by other groups to generate cDC in vitro did not
recapitulate the natural pathway of cDC development. DCs or their precursors are commonly
cultured in liquid medium, which is drastically different from the in vivo developmental
conditions. The natural environment for DC differentiation first takes place in the bone marrow,
where progenitors and precursors are embedded in the bone marrow stroma. DC precursors then
migrate to lymphoid tissues to finish their differentiation in tight association with lymphoid
stroma. Therefore, the use of a stromal monolayer for in vitro modeling of DC development is
imperative. Zhang et al212 generated DCs in liquid culture and placed fully differentiated DCs on
a stromal monolayer for further developmental studies. This approach may not be optimal
because unnatural DC development in liquid system may have altered DC differentiation
program and skewed further development towards abnormal immuno-regulatory populations
such as diffDCs described in the study. Our laboratory was the first to study DC differentiation
by culturing an immediate DC precursor (pre-cDC) on a stromal monolayer. This system allows
for tracing the natural progression of DC differentiation in the ‘steady state-like’ environment.
Pre-cDC placed on stroma proliferated vigorously to give rise to a pure population of cDCs
within 3 days. This population resembled in vivo isolated DCs – it was moderately potent in
stimulating allogeneic T cells proliferation and the stimulatory capacity was dramatically
38
increased when the cells were matured with LPS or TNFα. Furthermore, these cDCs responded
well to GM-CSF and, similarly to the in vivo inflammatory state, expanded dramatically in
number. When cDCs were allowed to propagate in the absence of GM-CSF, however, they did
not undergo apoptosis. Instead, these cells were observed to lose their DC identity (presence of
dendrites as part of classical DC morphology, expression of CD11c and MHCII, T cell
stimulatory capacity, response to classical DC maturation stimuli) and become potent immuno-
regulatory cells, which suppressed allogeneic and Ag-specific T cell proliferation.
Secondly, cytokine cocktail used in previous studies may not be optimal for identification of DC-
reg population arising from DCs. GM-CSF is usually added to the liquid culture to mature and
expand DCs generated from bone marrow progenitors. Some studies generating regulatory DCs
in vitro have used low levels of GM-CSF and/or IL1077, 96, 97. In the present study, however, GM-
CSF was shown to block the pathway leading to further DC differentiation into DC-regs.
Interestingly, GM-CSF was not identified as a key factor playing a role in generation of
regulatory populations in other studies98. The dependence on GM-CSF to generate DC-regs may
be explored further in some therapeutic applications like cancer immuno-therapy, where further
transformation of DCs into DC-regs is undesirable.
The third drawback of previous studies is the use of crude bone marrow extracts to generate
DCs, which makes difficult the study of developmental relationships in the DC lineage. The use
of bone marrow progenitors to generate immuno-suppressive populations generated in some
studies puts their DC origin under question. Moreover, it becomes impossible to dissect the
developmental origin of these immune-suppressive populations. For example, Svensson et al98
placed lineage- c-kit+ progenitors on stroma and generated CD11clow CD45RB+ CD11b+ cells. It
is, however, without reason that Svensson calls these cells DCs as merely excluding lineage cells
does not prove that they are DCs. In our study we definitively demonstrate that DC-regs do
indeed belong to the DC lineage. A highly pure FACS sorted pre-cDCs were cultured on stroma.
These cells were previously demonstrated by our group to give rise to DCs62. In the present
study, pre-cDCs gave rise to cDCs by day 3 and gradual loss of MHCII was observed by day 6.
By day 9 all cells lost CD11c expression as well. These results were confirmed by clonal assay
where single pre-cDC clones were cultured and generation of CD11c+ MHCII+ DCs and
subsequently CD11clow MHCII low DC-regs was seen (data not shown).
39
In vivo detection of regulatory dendritic cell populations has proven to be particularly
challenging. Most studies attribute regulatory properties to immature DCs occurring in lymphoid
organs and the periphery85-90. For example, CD103+ DCs have been described as capable of
inducing peripheral tolerance towards Ag in the gut and splenic CD8+ DCs have been shown to
induce T cell tolerance in spleen90, 91, 95. In addition, immature DCs with reduced T cell priming
capacity have been described in tumors149, 153, 154. However, regulatory populations that lack
typical DC features (such as expression of CD11c and MHCII) described in most studies have
not been detected in vivo. Wakkach et al has recently described a population of CD11clow
CD45RB+ population that was isolated from IL-10 transgenic as well as C56Bl/6 and BALB/c
mice99. These cells were first generated in vitro and then detected in vivo. However, their DC
origin remains to be determined as they were generated from crude bone marrow extract and in
the presence of low levels of GM-CSF.
Detection of DC-regs described in this study is also difficult using previous approaches. Past
studies used adoptive transfer model to demonstrate developmental relationships between
precursors and their progeny. For example, Naik et al61 isolated potential DC precursors from
spleen based on their density and expression of CD11c, CD45RA, SIRPα and CD43 and injected
them back into uninfected non-irradiated mice. The occurrence of donor-derived spleen cDCs
was then quantified by flow cytometry. This approach could not be utilized for the current study
as DC-regs described here could not be detected in the steady state conditions. Moreover,
previous detection of DC-regs was difficult due to lack of specific markers that could be used to
identify this population in vivo. Expression of CD11b, F4/80 and SIRPα that was detected on
DC-reg surface is non-specific and has also been described in other populations such as DCs,
macrophages and monocytes. Therefore, the use of CD11c-Cre+/- Rosa26-EGFP transgenic mice
in this study proved to be particularly useful. The expression of GFP driven by CD11c promoter
allowed not only to detect CD11clow MHCII low DC-regs in tumors but also to trace their
developmental history from CD11c+ MHCII+ cDCs. Using this model, this population was
detected in tumors (data not shown).
Under steady state conditions, lymphoid and non-lymphoid tissues contain stable numbers of
DCs, which is achieved through dynamic interactions between influx of new recruits, emigration,
and death. Bone marrow-derived pre-cDC, the immediate precursor of cDC, migrate via blood
40
and enter peripheral lymphoid and non-lymphoid tissues, where they differentiate into cDC that
proliferate for several generations. Non-lymphoid tissue cDC emigrate continuously to
lymphoid tissues via lymphatics into draining lymph nodes. Like the resident populations of
lymphoid DCs, most migrant tissue cDCs die in lymphoid tissue and do not re-enter the blood
circulation. The estimated life span of lymphoid tissue cDC is 2-3 days198. It remains unclear
whether cDC terminate through apoptosis exclusively. In the study performed by Chen et al. 199,
DC apoptosis was inhibited by expressing p35 under CD11c promoter. However, an increase in
total DC number became evident only in in mice > 3 months of age. Of note, inhibition of DC
apoptosis did not affect DC proliferation or DC generation from precursors. Moreover, no
autoimmune Ab generation was observed up to 9 months of age. These results suggest the
existence of additional mechanisms of systemic DC removal and, thereby, maintenance of
peripheral tolerance. This study raises the possibility that DC-reg differentiation is an alternative
fate for cDC.
The notion of functional and phenotypic plasticity in cDC is not unique. Recent evidence points
to the essential role of microenvironment in shaping the immunophenotype and function of other
cells of the innate immune system. Tumor associated macrophages have been shown to retain
their functional plasticity within the tumor. Upon treatment with IL-12, these cells down-
regulated pro-tumorigenic factors IL-10, TGF-β and MCP-1, and upregulated anti-tumorigenic
factors TNFα, IL-15 and IL-18 204. Moreover, interaction of M2 macrophages with Th1 cells has
been shown to skew macrophage phenotype towards M1 205. Recent evidence points to plasticity
in neutrophils as well. It has been argued that granulocytic CD11b+ Ly6C- Ly6G+ MDSCs seen
in tumors arise from neutrophils that have been modified by tumor microenvironment219. Besides
having a constellation of cell surface markers indicative of neutrophils, granulocytic MDSC also
express functional molecules commonly seen in neutrophils. For example, these MDSCs produce
high amounts of L-arginine and iNOS as well as have high oxidative capacity via high
expression of NADPH oxidase.
Cell plasticity is also being increasingly recognized in the adaptive immune system. For
example, propagation of committed Th17 precursors in the presence of IL-23 but in the absence
of TGF-β resulted in progressive loss of IL17F secretion and appearance of Th1-like IFNγ-
producing cells. Moreover, stimulation of Th17 cells with IL-12 led to rapid loss of Th17-
41
associated transcription factors and emergence of a Th1 gene signature 200. Several studies have
also shown the ability of Tregs to reprogram into inflammatory Th17 type cells in tumor draining
lymph nodes in the absence of IDO and presence of IL-6 201. In mice, Tregs have been shown to
directly differentiate into Th17 in the presence of IL-6 and in the absence of exogenous TGFβ 202. In humans, Tregs are also able to reprogram into Th17 cells when stimulated by allogeneic
monocytes in the presence of IL-2 and IL-15 203.
Hematopoietic cell lineage plasticity urges us to reconsider the classical view of the immune
system in the context of health and disease. The possibility of manipulating the
microenvironment to alter the functionality of a desired cell population has been explored for
some time. For example, administration of cytokines and growth factors for cancer
immunotherapy has been used in clinical trails with relative success220. IFNα has been
demonstrated in a number of trials to be successful at inducing anti-cancer response and
prolonging the survival of chronic myelogenous leukemia (CML) patients221. This therapy has
been shown to increase a number of CML-specific CD8+ T cells. Moreover, administration of
IL-2 was shown to be effective in a subset of patients with renal cell carcinoma222. This has been
shown to enhance both NK and CD8+ T cell function as well as increase vascular permeability.
TNFα, IL-12, GM-CSF and other cytokines have also been tested in clinical trials220. These
cytokines were shown to increase tumor cell apoptosis; enhance T cell cytotoxicity and inhibit
tumor angiogenesis; as well as increase tumor Ag presentation for TNFα, IL-12 and GM-CSF
respectively220. Unfortunately, systemic administration of cytokines for cancer immunotherapy is
limited by severe toxicities associated with these treatments. Moreover, this approach needs to be
optimized to achieve full therapeutic benefit for human use.
DC-regs generated from cDC potently inhibited proliferation of allogeneic and antigen-specific
OT-II T cells in vitro. For allogeneic T cells, decreased proliferation was accompanied by
reduced expression of effector cytokines. By contrast, intracellular cytokine expression in OT-II
T cells was unaffected by DC-regs. This difference may be due to the sensitivity of clonal OTI-II
T cells to re-stimulation. This result was similar to observation made by Zhang et al where
diffDCs suppressed proliferation of OVA-specific transgenic DO11.10 CD4+ T cells but not
cytokine production212.
42
In this study, DC-regs express high levels of arginase1, iNOS and IL-10 activity after stimulation
with LPS. They also constitutively expressed PD-L1 and ICOS-L. Moreover, iNOS was
determined to be responsible, at least in part, for the immunosuppressive properties of DC-regs.
An immunosuppressive effect of iNOS was reported to be due to generation of reactive nitrogen
species, which suppress T cell proliferation via impairment of IL-2R signalling 114, 115, 116, 117. NO
produced by iNOS is short lived and acts locally in a paracrine manner. In this study, the
suppression was reduced by about half when DC-regs were separated by a semi-permeable
membrane. It is not surprising considering that NO acts in a paracrine manner and may have,
therefore, been degraded before it could reach T cells in the lower compartment of a transwell
system. Similar to this study, Lukacs-Kornek et al223 recently reported inhibition of T cell
proliferation by stromal cells via iNOS-dependent mechanism. In this report T cell proliferation
was restored when a transwell membrane was inserted between T cells and stromal cells.
Moreover, nitrite production was decreased to baseline. Both these and our results indicate that
direct contact or close cell proximity between T cells and regulatory cells may be necessary for
iNOS induction and subsequent immunosuppression.
In addition to cell contact requirement, Lukacs-Kornek reports the involvement of IFNγ
produced by activated T cells in the induction of immunosuppression. We have tested induction
of potential immune-suppressive molecules by LPS. Although iNOS was produced in this
system, LPS is not the stimulus for iNOS induction in T cell proliferation assays. Here, T cells
were activated by allogeneic or OVA-pulsed DCs. This may have triggered IFNγ production,
which in turn could activate or induce iNOS. DC-regs were also observed to suppress
proliferation of effector memory T cells (data not shown), which secrete high amount of IFNγ.
The role of IFNγ in inhibition of T cell proliferation remains to be directly tested in our system.
Although DC-regs produced high levels of IL-10 upon stimulation with LPS, it was not directly
involved in inhibition of T cell proliferation. In this respect, DCregs in this study were similar to
diffDCs described by Zhang et al 212. DC-regs suppression of T cell proliferation was stronger
when they were not pulsed with OVA. This may be due to transfer of OVA Ag between DC-reg-
OVA and DCs or minor stimulation of TCR receptor on T cells through MHCII on DC-regs.
DC-regs suppressed cytokine release in T cells upon restimulation only in allogeneic but not in
the OT-II system. The disparity in DC-reg suppression between the allogeneic and clonal system
43
may be attributed to difference in sensitivity to Ag stimulation of both systems. In addition to
direct suppression of T cell proliferation, we evaluated the ability of DC-regs to induce CD4+
FoxP3+ Treg population. However, development of Tr1 cells in co-cultures was also possible.
The presence of this population was not assessed directly. Induction of other Treg subtypes (γδ
T cells, NKT cells, CD8+ Tregs) in co-cultures of OT-II cells with DC-regs is also unlikely as
highly purified CD4+ T cells were used in the initial cultures, which excludes other T cell
populations.
iNOS and ARG1 have been shown to be the major players in MDSC and TAM-mediated
immuno-suppression 166. These cells occur mostly in cancer and promote tumor progression 166.
In this study, DC-regs did not appear to be MDSCs, which comprise a heterogenous population
of Gr-1+ monocytes, neutrophils and primitive myeloid cells. DC-regs do not express Gr-1 and
are a homogeneous population derived from fully differentiated DC. High levels of iNOS and
ARG1 expression has also been previously described in macrophages 111. DC-regs resemble in
many respects alternatively activated macrophages and TAMs: they all express F4/80 (data not
shown) and high levels of CD11b, and have immuno-suppressive properties. A recent study
reported the existence of FoxP3+ macrophages in spleen, bone marrow, lymph nodes, thymus,
liver and other tissues of naïve mice; these cells inhibit T cell proliferation mainly via
prostaglandin E2 (PGE2) 209. Since blocking iNOS alone did not restore T cell proliferation
completely in our studies, it possible that other factors such as PGE2 expresson may be involved
in DCreg suppression. Future studies will address this possibility.
The immunosuppressive potency of DC-regs is similar to that reported in other CD11clow DC
populations. CD11clow diffDC generated by culturing mature DCs on embryonic spleen stroma
suppressed mature DC-induced DO11.10 T cell proliferation by ~70% but did not suppress
cytokine release 212, which was similar to our results in the OT-II system. Moreover, regulatory
capacity of our DC-regs cells in allogeneic system was also similar to those of
CD11clow CD45RB+ described by Svensson et al 98. These cells were generated on spleen stroma
from lin- c-kit+ progenitors. When compared to MDSC, DC-regs had similar potency with these
cells described in some studies 214 but were more than twice as potent as MDSC described in
other studies. It should be noted, however, that suppression of T cell response by MDSC was
44
assessed when T cells were stimulated non-specifically with anti-CD3 and anti-CD28 Abs126 or
with peptides without stimulator APCs present in culture 175, 215.
The role of DC-regs in immuno-suppression in vivo was also investigated in this study. The
results demonstrate that DC-regs do not suppress proliferation or effector function of adoptively
transferred T cells when injected i.p. Moreover, it was found that DC-regs pulsed with OVA are
able to prime naïve OT-II cells in vivo, induce up-regulation of T cell activation markers and
proliferation. The discrepancy between the in vivo and in vitro finding is difficult to explain. It is
possible that the read-out used in this study was not appropriate for detecting in vivo suppression.
Previous studies have shown that induction of T cell proliferation at day 3 does not always
correlate with the development of subsequent T cell tolerance. For example, mice injected with
a DEC-205:OVA conjugate, which delivers Ag to endogenous DC without causing DC
maturation, showed marked reduction in OT-II T cell numbers at day 12 as compared to mice
injected with DEC-205:OVA and anti-CD40 Ab, which causes DC maturaton, even though both
treatment regimens induced OT-II T cell proliferation initially 90. Similarly, another study found
marked T cell proliferation at day 2 in mice injected with a tolerogenic DEC-205:HEL
conjugate; however, T cell proliferation returned to baseline by day 7. By contrast, T cell
proliferation was sustained beyond 7 days in mice injected with immunostimulatory HEL in
CFA 87. It may, therefore, be worthwhile to evaluate T cell responses at later time point in our
model.
It is also possible that the inability of DC-regs to suppress in vivo responses represents their
failure to migrate to sites of DC-T cell interaction in lymphoid tissues. Since DC-regs resemble
macrophages, which are tissue-resident with limited migratory capacity 211, it seems reasonable
to speculate that i.p. injected DC-regs remained in the peritoneal cavity. In future studies, the
migratory capacity and chemokine receptor expression of DC-regs will be evaluated.
The role of the microenvironment in shaping cDC fate has become increasingly apparent. This
study suggests that the relative concentration of GM-CSF in tissues has critical role in
determining the fate of proliferating cDC. In the presence of high GM-CSF levels, the
proliferating progeny of cDC maintained a cDC phenotype, whereas in the absence of exogenous
GM-CSF they differentiated into DC-regs. Although DC-regs were generated in vitro in the
45
present study, recent evidence from our laboratory indicates that a similar cell population
develops in tumors. This study suggests that manipulating GM-CSF levels in tissues or its
downstream target maybe a useful strategy in the treatment of cancer and inflammatory diseases.
46
CHAPTER 6
Conclusion and Future Directions
1 Conclusion
In summary, these studies demonstrate two previously unknown attributes of cDC life cycle.
First, proliferating cDCs are not terminally differentiated but retain differentiation potential and
plasticity. When cDCs are co-cultured on a stromal monolayer in the absence of exognenous
GM-CSF, a novel immunosuppressive regulatory population of DC-regs is generated. These
cells suppress allogeneic T cell response and Ag-specific T cell proliferation in vitro through an
iNOS-dependent mechanism. Second, when cultured on stroma in the absence of GM-CSF, cDC
life does not finish with apoptosis. Instead, in the absence of GM-CSF, these cells are capable of
avoiding death by changing the course of their differentiation. The importance of this finding in
vivo is unclear. In the steady state and inflammation, where we have not been able to detect the
occurance of DC-regs, cDCs are likely to encounter endogenous or foreign Ag and, after a few
rounds of cell division, die by apoptosis in the lymphoid tissues. However, in the pathological
conditions such as cancer, natural cDC life cycle may be altered by microenvironment to
produce a new cell type with regulatory properties. These cells may be an important cause of
diseases where DC dysfunction is known to play a key role. The relevance of DC-reg occurance
in such pathologies remains to be investigated.
2 Future directions
The key role of GM-CSF in the generation of CD11clow MHCII low DC-regs from DCs was
demonstrated by adding the cytokine on days 0-12 of culture. We observed that DC-reg failed to
emerge when GM-CSF was added up to day 6 of culture. After that time point, addition of GM-
CSF did not prevent the development of DC-regs from DCs. However, GM-CSF may not be the
only factor involved in this transformation. Contrary to stromal co-culture, DCs cultured in
liquid medium in the presence of GM-CSF die within 2 days. Soluble or membrane bound
molecules produced by stromal cells, therefore, appear to be critical in the maintenance of DC
longevity and generation of DC-regs. Incubation of DCs with stromal supernatant can determine
47
if soluble factors are involved in differentiation of DCs into DC-regs. Some of the cytokines that
might be involved include: TGFβ, IL-10, IL-6, VEGF, hepatocyte growth factor (HGF), PGE2
and others. Alternatively, fixation of stromal cells will determine if cell contact is important in
the transformation212. In the study by Zhang et al it was reported that fibronectin plays a critical
role in generation of diffDCs from maDCs 212. Other factors like members of the fibroblast
growth factor family, laminin and apoptotic debris from stroma may also be important.
It would also be imperative to investigate changes in the genetic program that occurs during
transformation of cDCs into DC-regs. To do this, genome wide microarray analysis would be
done first to identify genetic signature of DC-regs. This would also help determine if DC-regs
resemble or belong to any of the presently known cell lineage (like macrophages) or if they
constitute a totally new cell type. It would then be possible to pinpoint specific genes involved in
the transformation and identify key transcription factors that act as signatures of DC-reg lineage.
Identification of these molecules would allow for the design of DC-reg-specific inhibitors or
inducers which could then be used therapeutically for the treatment of pathologies requiring DC
activation such as cancer or suppression such as transplantation. Epigenetic changes involved in
the transformation would also be interesting to look at.
DC-regs were generated from DCs in vitro in the studies presented. It will be important to
determine whether the same process occurs in vivo. Using the CD11c-Cre+ Rosa26-EGFP
transgenic mouse model, it has not been possible to detect DC-derived GFP+ CD11c- DC-regs in
the steady state. However, it appears that DC regs do develop in subcutaneous tumors derived
from B16 melanoma and lewis lung carcinoma (data not shown). The immunosuppressive
microenvironment within the tumor likely plays a pivotal role in the development of DCregs,
similar to the accumulation of MDSC. Once factors involved in the promotion of DC-regs are
identified in the in vitro system, it may be possible to inhibit DC reg develpment in vivo by
infusing blocking monoclonal Abs i.v. or into the tumor. So far, in vitro evidence points to the
importance of GM-CSF in the maintenance of DC phenotype. To determine whether increasing
the concentration of GM-CSF in the tumors could influence cDC differentiation, tumor cells
could be induced with a retroviral vector encoding GM-CSF. In fact, several Phase I clinical
trials investigating the role of GM-CSF in tumor microenvironment have done just that. In these
studies, surgically excised tumors (metastatic melanoma or non-small-cell lung carcinoma) were
48
transduced with replication defective GM-CSF expressing retrovirus and re-injected back into
patients224-226. Another study injected allogeneic GM-CSF-secreting HER2-expressing tumor cell
line for vaccination against breast tumor227. In this study, one out of six patients developed anti-
HER2 Abs when injected with vaccine alone and 7 out of 22 patients when injected with vaccine
in combination with chemotherapy. Vaccination sites showed substantial infiltration of DCs,
macrophages and granulocytes224-226. Moreover, metastatic lesions that were excised after but not
before vaccination showed considerable infiltration of CD4+ and CD8+ T lymphocytes and B
cells as well as necrosis, fibrosis and oedema. It was also demonstrated in a pre-clinical mouse
model that tumors engineered to secrete GM-CSF induce massive infiltration and activation of
DCs into tumor tissue, which then prime Ag-specific CD4+ and CD8+ DCs228.
The occurrence of CD11clow DC-regs in other immune conditions is also possible. The
resolution phase of acute inflammation is a likely situation given that this process is
characterized by massive leukocyte apoptosis which occurs in association with increased
frequency of various regulatory cells. Key molecules involved in inflammation resolution
include prostaglandin E2, lipotoxin, resolvins and protectins 208. Involvement of lipids in DC
dysfunction has been reported before 216. It would be interesting to determine whether DCregs
develop during the resolution phase of inflammation and whether the immunosuppressive
molecules linked to this process promote DCreg development. Chronic inflammatory processes
may also favour the development of DCregs and should be investigated in the future.
In experimental conditions used, DC-regs did not show any immunosuppression in vivo. There
are several explanations for the observed phenomenon. One possibility is that T cell proliferation
measured at 3 days has no relationship with the establishment of tolerance mediated by DC-regs.
T cell proliferation or other read-outs for tolerance could be measured at later time points (eg.
after day 10). Moreover, alternative routes of T cell priming may be utilized. It was
demonstrated in this study that DC-regs upregulate various regulatory molecules upon exposure
to a TLR agonist LPS. In our in vivo model, we did not inject LPS into the system and OT- II
cells were primed by injection of OVA-pulsed DCs i.p. We could try challenging the mice with
LPS and testing if DC-regs could suppress systemic inflammatory response. OT-II cells could
also be primed with OVA in Complete Freud’s Adjuvant (CFA) to induce systemic T cell
priming and inflammation. Because CFA contains bacterial components, it is likely to act as
49
TLR agonist, activating regulatory molecule production similarly to LPS. This approach has
been taken by Kusmartsev et al229 where MDSC were observed to suppress OT-I T cell
proliferation in response to OVA in Incomplete Freud’s Adjuvant (IFA) challenge. Moreover, to
avoid DC-reg migration issue, DC-regs could be injected localy into the footpad to increase
interaction between T cells and DC-regs, which would be immunized with OVA in CFA. Since
DC-regs express high levels of IL-10 and iNOS, it would also be interesting to test whether DC-
regs could use therapeutically to down-regulate acute inflammatory responses.
50
Figures
MHC II
Lin-
+/- GM
No GM-CSFGM-CSF
MH
C II
CD
11c
CD11c
Pre-cDC
Day 3
Day 9
A
CD11clow
MHCIIlow CD11c+ MHCII+B
-T
NFα
+ T
NFα
C
Day 3 Day 10Day 6
CD11c
MH
C II
Day 0
Figure 3. Differentiation and proliferation of pre-cDC on stroma. A) FACS purified CD11c+ MHCII - lin- (CD3, CD19, B220, CD49b) pre-cDC were placed on a primary fibroblast stromal monolayer in the presence or absence of GM-CSF (4ng/mL). Pre-cDC generated proliferating cDC at day 3 under both conditions. In the absence of GM-CSF, the cDC progeny at 10 days did not express CD11c and MHCII markers and turned into CD11clow MHCII low cell type. B) Time course of pre-cDC development into CD11clow MHCII low cell through CD11c+ MHCII+ DC intermediate. C) Morphology of CD11clow MHCII low and CD11c+ MHCII+ cells following exposure to TNFα. Cells were Giemsa stained and viewed under the light microscope (40× magnification).
51
CD11c MHCII GFP
Day 0
Day 3
Day 10
CD11c-Cre
ROSA-stopFlox-GFP
CD11c-Cre-ROSA-GFP
X
A
B
Figure 4. CD11clow MHCIIlow arise from CD11c+ MHC II+cDC. A) CD11c-Cre mice were crossed with ROSA-stopFlox-GFP mouse to create CD11c-Cre-ROSA-GFP mouse. B) Pre-cDC from Cre+/- mice (white histogram) and Cre-/- littermate controls (grey histogram) were placed on stroma in the absence of GM-CSF. The histograms show expression of CD11c, MHC II, and GFP by pre-cDC at day 0, and their progeny at day 3 and day 10.
52
CD11c CD11bMHCII CD103 CD4 CD8 CD172a
Spleen cDC
CD11clow
MHCIIlow DC
Figure 5. Immunophenotype of cDC-derived CD11clow MHCIIlow cells. Cells were stained with the indicated markers and analyzed by flow cytometry.
53
DC DC+LPS
CD40
CD80
CD86
MHCII
DC-reg DC-reg+LPS
CD11c
Figure 6. DC-regs fail to up-regulate co-stimulatory molecules in response to maturation stimuli. Spleen DCs or cultured DC-regs were stimulated with LPS (2µg/mL) for 18 hours. Cell surface expression of co-stimulatory molecules (CD40, CD80, CD86) were assessed by flow cytometry. Data is representative of 3 independent experiments.
54
Figure 7. CD11clow MHCIIlow cells are poor stimulators of allogeneic T cell lymphocytes. BALB/c responder T cells were pulsed with C57BL/6 DCs or DC-regs at various ratios. T cell proliferation was determined on day 3 by 3H-thymidine incorporation.
55
Figure 8. CD11clow MHCIIlow DC-derived cells have increased phagocytic capacity. A) CD11clow MHCII low DC-derived cells and CD11c+ MHCII+ DCs were pulsed with OVA-FITC at 4º C and 37º C and mean fluorescence intensity was measured by flow cytometry. B) Cells were incubated with fluorescent dextran beads and viewed under confocal microscope (40× magnification). Results are representative of 3 independent experiments.
4oc 37 oc4oc 37oc
OVA
CD11clow MHCIIlow DC CD11c+ MHCII + DC
CD11clow
MHCIIlow DC CD11c +
MHCII+ DC
A
B
MFI=30 MFI=15
56
Figure 9. CD11clow MHCIIlow suppress T cell proliferation in allogeneic mixed lymphocyte reaction. Balb/c responder T cells were pulsed with DCs +/- DC-regs at various ratios. T cell proliferation was determined on day 3 by 3H-thymidine incorporation.
3
0
2
4
6
8
10
12
14
4:1 2:1 1:1 0.5:1 0:4 0:2 0:1 noAPC
3 H t
hym
idin
e in
corp
ora
tio
n (
cpm
) x
103
3
DC -reg: DC ratio
57
0
5
10
15
20
25
IFNy
% I
FN
y +
ve c
ells
0
2
4
6
8
10
12
IL4
% I
L-4
+ve
cel
ls
0
5
10
15
20
25
30
IL2
% I
L-2
+ve
ce
lls
DC DC+DC-reg DC-reg
DC DC+DC-reg DC-reg
DC DC+DC-reg DC-reg
*
* *
Figure 10. DC-regs suppress effector function of allogeneic T cells in mixed lymphocyte cultures. Balb/c spleen cells stimulated with DC+/- DC-regs for 7 days were harvested, washed, and re-stimulatedwith anti-CD3 and anti-CD28 monoclonal antibodies. Intracellular expression of IFN-γ, IL-2 and IL-4 was measured by flow cytometry. * p<0.01 by Student`s t-test. Data are representative of >3 independent experiments.
58
Figure 11. DC-regs suppress OT-II T cell proliferation in response to OVA-pulsed DCs. 1×105 OT-II T cells were stimulated with 1×104 DC-OVA +/- 1×104 DC-reg. T cells were harvested on day 4 and cell number was assessed by flow cytometry. * p<0.01 Results are pooled from 3 independent experiments.
0 2 4 6 8 10 12
treatment
cell number (*105)
No APC DC -OVA
DC-OVA + DC-reg-OVADC-OVA + DC-reg
DC-reg-OVA
DC- reg
*
*
*
*
59
CFSE
DC-OVADC-reg-OVA
DC-reg
+--
++-
+-+
-+-
--+
* White = no APC
Figure 12. DC-regs suppress OT-II T cell proliferation. 1×105 CFSE-labelled OT-II T cells were stimulated with 1×104 DC-OVA+/- 1×104 DC-regs. Cells were harvested on day 3 and T cell proliferation (CFSE dilution) was assessed by flow cytometry. Results are representative of 3 independent experiments.
60
0
5
10
15
20
25
30
35
40
IL2
% I
L2
+ve
cel
ls
DC-OVA
DC-reg
DC-reg-OVA
+ + + - -
- + - + -
- - + - +
0
2
4
6
8
10
12
14
16
IFNy
% I
FN
y +
ve c
ells
Figure 13. DC-regs do not suppress OT-II T cell cytokine release. 1×105 OT-II T cells stimulated with 1×104 DC-OVA +/- 1×104 DC-regs for 7 days. Cells were then harvested, washed and re-stimulated with anti-CD3 and anti-CD28 Abs for 12 hours. Intracellular cytokine production was assessed by flow cytometry.
61
DC DC + DC-reg DC-reg
Day 1
Day 3
CD25
Day 1
Day 3
CD44
Figure 14. Expression of CD25 and CD44 by T cells in allogeneic mixed lymphocyte reaction in the presence or absence of DC-regs. 1×105 BALB/c T cells were isolated from spleen and stimulated with 1×104 C57BL/6 DC +/- 1×104 C57BL DC-regs for 3 days. Expression of CD25 and CD44 was the assessed by flow cytometry. White histograms indicate un-stimulated T cells. Results are representative of 3 independent experiments.
62
DC-ovaDC-reg-ova
DC-reg
+--
++-
+-+
-+-
--+
* White = no APC
PI
Figure 15. DC-regs do not induce T cell death in mixed lymphocyte reactions. 1×105 OT-II T cells were stimulated with 1×104 DC-OVA +/- 1×104 DC-regs for 3 days. Propidium iodide (PI) staining of dead cells was assessed by flow cytometry. White histograms indicate un-stimulated cells. Results are representative of 3 independent experiments.
63
CD25
CD
4
CD25
Fo
xP3
DC
DC + DC-reg
1
0
66
64
CFSE
Res
po
nd
erT
cel
l No
.
Figure 16. DC-regs do not induce Foxp3+ Tregs in mixed lymphocyte cultures. CFSE-labelled 1×105 BALB/c T cells were stimulated with 1×104 C57BL/6 DC +/- C57BL/6 1×104 DC-regs for 7 days, harvested, stained for CD4, CD25, and FoxP3, and analyzed by flow cytometry. Results are representative of 3 independent experiments.
64
0 2000 4000 6000 8000 10000 12000
T + DC-ova
T + DC-ova + DC-reg-ova
T + DC-ova // DC-reg-ova
Cell number
Figure 17. Mechanism of DC-reg-mediated immuno-suppression involves both soluble and contact-dependent factors. 1×106 OT-II cells along with 1×105 DC-OVA were placed in the bottom chamber. 1×105 DC-reg-OVA were placed in the top chamber. Proliferation was assessed three days later by flow cytometry. Results represent data pooled from 3 independent experiments.
65
iNOS
DC
-reg
DC
-reg
+ LP
SS
ple
enS
ple
en +
LP
S
arginase1
FasL
PD-L1
IDO
ICOS-L
DC
-reg
DC
-reg
+ L
PS
Sp
leen
Sp
leen
+ L
PS
IL-10
Figure 18. RT-PCR expression of candidate molecules responsible for observed in vitro immuno-suppression. DC-regs or freshly isolated spleen mononuclear cells were pulsed with 2 µg/mL LPS overnight prior to gene expression evaluation.
66
02468
10121416
DC
DC
+ L
PS
DC
-reg
DC
-reg
+ L
PS
arg
inas
e ac
tivity
(mU
/ug
)
010
2030
4050
DC
DC
+ L
PS
DC
-reg
DC
-reg
+ L
PS
nitr
ite c
on
c (u
M)
A B
* * ** * *
Figure 19. DC-regs express high levels of arginase1 and iNOS (reflected by nitrite production) activity when stimulated with LPS. 1×106 cells/mL were pulsed with 2 µg/mL LPS overnight. A) Cells were harvested, washed and lysed. Arginase1 activity was determined by incubating cell lysate with L-arginine for 60 min and measuring resulting urea production. B) Supernatants were harvested and nitrite levels were detected by Griess reagent. * p<0.01 Data was pooled from 3 independent experiments.
67
DC DC-reg
DC + LPS DC-reg + LPS
PD-L1
Figure 20. Both DCs and DC-regs express high levels of PD-L1. 1×106 DC or DC-regs were pulsed with 2 µg/mL LPS overnight. Cells were then harvested, washed, stained, and assessed by flow cytometry. White histograms indicate isotype control. Result is representative of 3 independent experiments.
68
0
500
1000
1500
2000
2500
DC DC + LPS DC-reg DC-reg +LPS
IL10
(p
g/m
L)
ND ND ND
Figure 21. LPS-pulsed DC-regs express high levels of IL-10. Freshly isolated spleen DCs or DC-regs were pulsed with 2 µg/mL LPS overnight. Supernatants were harvested and IL-10 levels were detected by ELISA.
69
3 H-t
hym
idin
e in
corp
ora
tion
(cp
m)
1x03
DC-OVA + DC-reg
DC-OVA
L-NIL
Isoty
pe
Figure 22. DC-regs suppress T cell proliferation through an iNOS-dependent mechanism. 1×105 OT-I T cells were stimulated with 1×104 OVA peptide-pulsed DC +/- 1×104 DC-reg in the presence of IL-10R blocking Ab or chemical inhibitors Nor-NOHA and L-NIL that block arginase1 and iNOS activity, respectively. Cells were cultured for 3 days and proliferation was assessed by 3H- thymidine incorporation. Data is representative of 3 independent experiments.
70
Figure 23. DC-regs do not suppress T cell proliferation and activation in vivo. A) In vivo experimental scheme. CD45.1+ B6.SJL mice were i.v. injected with 1×106 CD45.2+ OT-II cells. The next day, mice were immunized with 1×106 DC +/- 1×106 DC-regs i.p. Mice were sacrificed on days 3 and 5. Cell proliferation was assessed by CFSE dilution. Spleen and lymph node cells were re-stimulated with OVA in vitro and cytokine production was assessed by flow cytometry 12 hours later. B) OT-II cells were injected into a mouse as in the experimental scheme above and primed with DC +/- DC-regs. Spleen and lymph nodes were harvested on days 3 and 5. OT-II cell proliferation was assessed by CFSE dilution. C) Spleen and lymph nodes were harvested on days 3 and 5 and re-stimulated with OVA in vitro. IFN-γ production was evaluated by flow cytometry.
CD45.2+ OT-II cells iv
Day 0
DC +/- DC- regip
Day 1
Assess T cell proliferation
+Cytokine
production
Day 3, 5
CD45.1+
A No APC
DAY 5
DAY 3
CFSE
CD
45.2
+ C
D4+
O
TII
Cel
l No
.
DC- ova
DC - ova + DC -reg- ova
B
0
10
20
30
40
50
-OVA DC-ova DC-ova +DC-reg-ova
DC-reg-ova
% IF
Ny
+ve
cells
Day 3
Day 5
C
71
No APCDC-ova
+ DC
DAY 5
DAY 3
CFSE
CD
45.2
+ C
D4+
O
TII
Cel
l No
.
DC-ova + DC-reg
A
05
1015
2025
3035
4045
-OVA DC-ova + DC DC-ova + DC-reg
% IF
Ny
+ve
cells
Day 3
Day 5
B
Figure 24. DC-regs fail to suppress T cell proliferation and activation in vivo. A) OT-II cells were injected into mouse as in the experimental scheme above and primed with DC +/- DC-regs. Number of APCs was normalized in both groups with co-injection of DCs that were not pulsed with OVA. Spleen and lymph nodes were harvested on days 3 and 5. OT-II cell proliferation was assessed by CFSE dilution. B) Spleen and lymph nodes were harvested on days 3 and 5 and re-stimulated with OVA in vitro. IFN-γ production was evaluated by flow cytometry.
72
No APC DC-reg-ovaDC-ova
CFSE
CD44
CD69
Figure 25. DC-regs induce OT-II cell activation in vivo. OT-II cells were injected into mouse as in the experimental scheme above and primed with DC +/- DC-regs. Spleens were harvested on day 3. Expression of activation markers (CD44, CD69) on CD45.2+ OT-II cells were evaluated by flow cytometry.
73
References
1. Steinman, R. M. & Cohn, Z. A. Identification of a novel cell type in peripheral lymphoid organs of mice. The Journal of Experimental Medicine 137, 1142-1162 (1973).
2. Liu, K. & Nussenzweig, M. C. Origin and development of dendritic cells. Immunological Reviews 234, 45 (2010).
3. Inaba, K. et al. Generation of large numbers of dendritic cells from mouse bone marrow cultures supplemented with granulocyte/macrophage colony-stimulating factor. The Journal of Experimental Medicine 176, 1693-1702 (1992).
4. Lutz, M. B. et al. An advanced culture method for generating large quantities of highly pure dendritic cells from mouse bone marrow. J. Immunol. Methods 223, 77-92 (1999).
5. Asselin-Paturel, C. et al. Mouse type I IFN-producing cells are immature APCs with plasmacytoid morphology. Nat. Immunol. 2, 1144-1150 (2001).
6. Shortman, K. & Villadangos, A. J. in Handbook of dendritic cells. Bology, diseases, and therapeutics (eds Lutz, M. B., Romani, N. & Steinkasserer, A.) 199 (Wiley-VCH Verlag GmbH & Co., 2006).
7. Vremec, D., Pooley, J., Hochrein, H., Wu, L. & Shortman, K. CD4 and CD8 Expression by Dendritic Cell Subtypes in Mouse Thymus and Spleen. The Journal of Immunology 164, 2978-2986 (2000).
8. McLellan, A. D. et al. Anatomic location and T-cell stimulatory functions of mouse dendritic subsets defined by CD4 and CD8 expression. Blood 99, 2084 (2002).
9. Henri, S. et al. The Dendritic Cell Populations of Mouse Lymph Nodes. The Journal of Immunology 167, 741-748 (2001).
10. Kamath, A. T. et al. The Development, Maturation, and Turnover Rate of Mouse Spleen Dendritic Cell Populations. The Journal of Immunology 165, 6762-6770 (2000).
11. den Haan, J. M. M., Lehar, S. M. & Bevan, M. J. Cd8+ but Not Cd8− Dendritic Cells Cross-Prime Cytotoxic T Cells in vivo. The Journal of Experimental Medicine 192, 1685-1696 (2000).
12. Pooley, J. L., Heath, W. R. & Shortman, K. Cutting Edge: Intravenous Soluble Antigen Is Presented to CD4 T Cells by CD8- Dendritic Cells, but Cross-Presented to CD8 T Cells by CD8+ Dendritic Cells. The Journal of Immunology 166, 5327-5330 (2001).
13. Edwards, A. D. et al. Microbial Recognition Via Toll-Like Receptor-Dependent and -Independent Pathways Determines the Cytokine Response of Murine Dendritic Cell Subsets to CD40 Triggering. The Journal of Immunology 169, 3652-3660 (2002).
74
14. Maldonado-López, R. et al. CD8α+ and CD8α− Subclasses of Dendritic Cells Direct the Development of Distinct T Helper Cells In vivo. The Journal of Experimental Medicine 189, 587-592 (1999).
15. Boonstra, A. et al. Flexibility of Mouse Classical and Plasmacytoid-derived Dendritic Cells in Directing T Helper Type 1 and 2 Cell Development. The Journal of Experimental Medicine 197, 101-109 (2003).
16. Edwards, A. et al. Toll-like receptor expression in murine DC subsets: lack of TLR7 expression by CD8α+ DC correlates with unresponsiveness to imidazoquinolines. Eur. J. Immunol. 33, 827-833 (2003).
17. Sallusto, F. & Lanzavecchia, A. Understanding dendritic cell and T-lymphocyte traffic through the analysis of chemokine receptor expression. Immunological Reviews 177, 134 (2000).
18. Proietto, A. I. et al. Differential production of inflammatory chemokines by murine dendritic cell subsets. Immunobiology 209, 163-172 (2004).
19. Förster, R. et al. CCR7 Coordinates the Primary Immune Response by Establishing Functional Microenvironments in Secondary Lymphoid Organs. Cell 99, 23-33 (1999).
20. Ohl, L. et al. CCR7 Governs Skin Dendritic Cell Migration under Inflammatory and Steady-State Conditions. Immunity 21, 279-288 (2004).
21. Sozzani, S. et al. Cutting Edge: Differential Regulation of Chemokine Receptors During Dendritic Cell Maturation: A Model for Their Trafficking Properties. The Journal of Immunology 161, 1083-1086 (1998).
22. Dieu, M. et al. Selective Recruitment of Immature and Mature Dendritic Cells by Distinct Chemokines Expressed in Different Anatomic Sites. The Journal of Experimental Medicine 188, 373-386 (1998).
23. Merad, M., Ginhoux, F. & Collin, M. Origin, homeostasis and function of Langerhans cells and other langerin-expressing dendritic cells. Nat Rev Immunol 8, 935-947 (2008).
24. Valladeau, J. et al. Langerin, a Novel C-Type Lectin Specific to Langerhans Cells, Is an Endocytic Receptor that Induces the Formation of Birbeck Granules. Immunity 12, 71-81 (2000).
25. Ginhoux, F. et al. Blood-derived dermal langerin+ dendritic cells survey the skin in the steady state. J. Exp. Med. 204, 3133-3146 (2007).
26. Poulin, L. F. et al. The dermis contains langerin+ dendritic cells that develop and function independently of epidermal Langerhans cells. J. Exp. Med. 204, 3119-3131 (2007).
27. Wolff, K. The fine structure of the langerhans cell granule. J. Cell Biol. 35, 468-473 (1967).
75
28. Wang, L. et al. Langerin Expressing Cells Promote Skin Immune Responses under Defined Conditions. The Journal of Immunology 180, 4722-4727 (2008).
29. Merad, M. et al. Langerhans cells renew in the skin throughout life under steady-state conditions. Nat. Immunol. 3, 1135-1141 (2002).
30. Borkowski, T. A., Letterio, J. J., Farr, A. G. & Udey, M. C. A Role for Endogenous Transforming Growth Factor β1 in Langerhans Cell Biology:  The Skin of   Transforming Growth Factor β1 Null Mice Is Devoid of  Epidermal Langerhans Cells. J. Exp. Med. 184, 2417-2422 (1996).
31. Ginhoux, F. et al. Langerhans cells arise from monocytes in vivo. Nat. Immunol. 7, 265-273 (2006).
32. Kamath, A. T., Henri, S., Battye, F., Tough, D. F. & Shortman, K. Developmental kinetics and lifespan of dendritic cells in mouse lymphoid organs. Blood 100, 1734-1741 (2002).
33. Bursch, L. S. et al. Identification of a novel population of Langerin+ dendritic cells. J. Exp. Med. 204, 3147-3156 (2007).
34. Varol, C. et al. Intestinal Lamina Propria Dendritic Cell Subsets Have Different Origin and Functions. Immunity 31, 502-512 (2009).
35. Denning, T. L. et al. Functional Specializations of Intestinal Dendritic Cell and Macrophage Subsets That Control Th17 and Regulatory T Cell Responses Are Dependent on the T Cell/APC Ratio, Source of Mouse Strain, and Regional Localization. The Journal of Immunology 187, 733-747 (2011).
36. Denning, T. L., Wang, Y., Patel, S. R., Williams, I. R. & Pulendran, B. Lamina propria macrophages and dendritic cells differentially induce regulatory and interleukin 17-producing T cell responses. Nat. Immunol. 8, 1086-1094 (2007).
37. Rescigno, M. et al. Dendritic cells express tight junction proteins and penetrate gut epithelial monolayers to sample bacteria. Nat. Immunol. 2, 361-367 (2001).
38. Iwasaki, A. & Kelsall, B. L. Localization of Distinct Peyer's Patch Dendritic Cell Subsets and Their Recruitment by Chemokines Macrophage Inflammatory Protein (Mip)-3α, Mip-3β, and Secondary Lymphoid Organ Chemokine. The Journal of Experimental Medicine 191, 1381-1394 (2000).
39. Iwasaki, A. & Kelsall, B. L. Unique Functions of CD11b+, CD8α+, and Double-Negative Peyer’s Patch Dendritic Cells. The Journal of Immunology 166, 4884-4890 (2001).
40. Iwasaki, A. & Kelsall, B. L. Freshly Isolated Peyer's Patch, but Not Spleen, Dendritic Cells Produce Interleukin 10 and Induce the Differentiation of T Helper Type 2 Cells. J. Exp. Med. 190, 229-240 (1999).
76
41. Wu, L. & Shortman, K. Heterogeneity of thymic dendritic cells. Semin. Immunol. 17, 304-312 (2007).
42. Steinman, R. M., Hawiger, D. & Nussenzweig, M. C. TOLEROGENIC DENDRITIC CELLS×. Annu. Rev. Immunol. 21, 685-711 (2003; 2003).
43. Li, J., Park, J., Foss, D. & Goldschneider, I. Thymus-homing peripheral dendritic cells constitute two of the three major subsets of dendritic cells in the steady-state thymus. J. Exp. Med. 206, 607-622 (2009).
44. Wilson, N. S. et al. Most lymphoid organ dendritic cell types are phenotypically and functionally immature. Blood 102, 2187-2194 (2003).
45. Nakano, H., Yanagita, M. & Gunn, D. M. CD11c+ B220+ Gr-1+ cells in mouse lymph nodes and spleen display characteristics of plasmacytoid dendritic cells. Journal of Experimental Medicine 194, 1171 (2001).
46. Colonna, M., Trinchieri, G. & Liu, Y. Plasmacytoid dendritic cells in immunity. Nature immunology 5, 1219 (2004).
47. O'Keeffe, M. et al. Mouse Plasmacytoid Cells. The Journal of Experimental Medicine 196, 1307-1319 (2002).
48. Liu, Y. IPC: Professional type I interferon-producing cells and plasmacytoid dendritic cell precursors. Annual Review of Immunology 23, 275 (2005).
49. Krug, A. et al. TLR9-Dependent Recognition of MCMV by IPC and DC Generates Coordinated Cytokine Responses that Activate Antiviral NK Cell Function. Immunity 21, 107-119 (2004).
50. Cella, M., Facchetti, F., Lanzavecchia, A. & Colonna, M. Plasmacytoid dendritic cells activated by influenza virus and CD40L drive a potent TH1 polarization. Nat. Immunol. 1, 305-310 (2000).
51. Martı�n, P. et al. Characterization of a new subpopulation of mouse CD8α+ B220+ dendritic cells endowed with type 1 interferon production capacity and tolerogenic potential. Blood 100, 383-390 (2002).
52. Dalod, M. et al. Dendritic Cell Responses to Early Murine Cytomegalovirus Infection. The Journal of Experimental Medicine 197, 885-898 (2003).
53. Ardavín, C. et al. Origin and differentiation of dendritic cells. Trends Immunol. 22, 691-700 (2001).
54. Ardavin, C., Wu, L., Li, C. & Shortman, K. Thymic dendritic cells and T cells develop simultaneously in the thymus from a common precursor population. Nature 362, 761-763 (1993).
77
55. Wu, L., Li, C. L. & Shortman, K. Thymic dendritic cell precursors: relationship to the T lymphocyte lineage and phenotype of the dendritic cell progeny. The Journal of Experimental Medicine 184, 903-911 (1996).
56. Traver, D. et al. Development of CD8α-Positive Dendritic Cells from a Common Myeloid Progenitor. Science 290, 2152-2154 (2000).
57. Martı�n, P. et al. Concept of lymphoid versus myeloid dendritic cell lineages revisited: both CD8α− and CD8α+dendritic cells are generated from CD4lowlymphoid-committed precursors. Blood 96, 2511-2519 (2000).
58. Manz, M. G., Traver, D., Miyamoto, T., Weissman, I. L. & Akashi, K. Dendritic cell potentials of early lymphoid and myeloid progenitors. Blood 97, 3333-3341 (2001).
59. Wu, L. et al. Development of thymic and splenic dendritic cell populations from different hemopoietic precursors. Blood 98, 3376-3382 (2001).
60. Fogg, D. K. et al. A Clonogenic Bone Marrow Progenitor Specific for Macrophages and Dendritic Cells. Science 311, 83-87 (2006).
61. Naik, S. H. et al. Development of plasmacytoid and conventional dendritic cell subtypes from single precursor cells derived in vitro and in vivo. Nat. Immunol. 8, 1217-1226 (2007).
62. Diao, J., Winter, E., Chen, W., Cantin, C. & Cattral, M. S. Characterization of Distinct Conventional and Plasmacytoid Dendritic Cell-Committed Precursors in Murine Bone Marrow. The Journal of Immunology 173, 1826-1833 (2004).
63. Diao, J. et al. In Situ Replication of Immediate Dendritic Cell (DC) Precursors Contributes to Conventional DC Homeostasis in Lymphoid Tissue. The Journal of Immunology 176, 7196-7206 (2006).
64. Naik, S. H. et al. Intrasplenic steady-state dendritic cell precursors that are distinct from monocytes. Nat. Immunol. 7, 663-671 (2006).
65. D'Amico, A. & Wu, L. The Early Progenitors of Mouse Dendritic Cells and Plasmacytoid Predendritic Cells Are within the Bone Marrow Hemopoietic Precursors Expressing Flt3. The Journal of Experimental Medicine 198, 293-303 (2003).
66. Karsunky, H., Merad, M., Cozzio, A., Weissman, I. L. & Manz, M. G. Flt3 Ligand Regulates Dendritic Cell Development from Flt3+ Lymphoid and Myeloid-committed Progenitors to Flt3+ Dendritic Cells In vivo. The Journal of Experimental Medicine 198, 305-313 (2003).
67. McKenna, H. J. et al. Mice lacking flt3 ligand have deficient hematopoiesis affecting hematopoietic progenitor cells, dendritic cells, and natural killer cells. Blood 95, 3489-3497 (2000).
78
68. Maraskovsky, E. et al. Dramatic increase in the numbers of functionally mature dendritic cells in Flt3 ligand-treated mice: multiple dendritic cell subpopulations identified. J. Exp. Med. 184, 1953-1962 (1996).
69. Vremec, D. et al. The influence of granulocyte/macrophage colony-stimulating factor on dendritic cell levels in mouse lymphoid organs. Eur. J. Immunol. 27, 40-44 (1997).
70. Kabashima, K. et al. Intrinsic Lymphotoxin-β Receptor Requirement for Homeostasis of Lymphoid Tissue Dendritic Cells. Immunity 22, 439-450 (2005).
71. Diao, J., Winter, E., Chen, W., Xu, F. & Cattral, M. S. Antigen Transmission by Replicating Antigen-Bearing Dendritic Cells. The Journal of Immunology 179, 2713-2721 (2007).
72. Geissmann, F., Jung, S. & Littman, D. R. Blood Monocytes Consist of Two Principal Subsets with Distinct Migratory Properties. Immunity 19, 71-82 (2003).
73. Randolph, G. J., Inaba, K., Robbiani, D. F., Steinman, R. M. & Muller, W. A. Differentiation of Phagocytic Monocytes into Lymph Node Dendritic Cells In vivo. Immunity 11, 753-761 (1999).
74. Varol, C. et al. Monocytes give rise to mucosal, but not splenic, conventional dendritic cells. The Journal of Experimental Medicine 204, 171-180 (2007).
75. Inaba, K. et al. Identification of proliferating dendritic cell precursors in mouse blood. The Journal of Experimental Medicine 175, 1157-1167 (1992).
76. Brasel, K., De Smedt, T., Smith, J. L. & Maliszewski, C. R. Generation of murine dendritic cells from flt3-ligand–supplemented bone marrow cultures. Blood 96, 3029-3039 (2000).
77. Sato, K., Yamashita, N., Yamashita, N., Baba, M. & Matsuyama, T. Regulatory Dendritic Cells Protect Mice from Murine Acute Graft-versus-Host Disease and Leukemia Relapse. Immunity 18, 367-379 (2003).
78. Jenkinson, E. J., Jhittay, P., Kingston, R. & Owen, J. J. T. Studies of the role of thymic environment in the induction of tolerance to MHC antigens. Transplantation 39, 331 (1985).
79. Matzinger, P. & Guerder, S. Does T-cell tolerance require a dedicated antigen-presenting cell? Nature 338, 74-76 (1989).
80. Brocker, T., Riedinger, M. & Karjalainen, K. Targeted Expression of Major Histocompatibility Complex (MHC) Class II Molecules Demonstrates that Dendritic Cells Can Induce Negative but Not Positive Selection of Thymocytes In vivo. The Journal of Experimental Medicine 185, 541-550 (1997).
81. Anderson, G., Partington, K. M. & Jenkinson, E. J. Differential Effects of Peptide Diversity and Stromal Cell Type in Positive and Negative Selection in the Thymus. The Journal of Immunology 161, 6599-6603 (1998).
79
82. Gallegos, A. M. & Bevan, M. J. Central Tolerance to Tissue-specific Antigens Mediated by Direct and Indirect Antigen Presentation. J. Exp. Med. 200, 1039-1049 (2004).
83. Bonasio, R. et al. Clonal deletion of thymocytes by circulating dendritic cells homing to the thymus. Nat. Immunol. 7, 1092-1100 (2006).
84. Proietto, A. I. et al. Dendritic cells in the thymus contribute to T-regulatory cell induction. Proceedings of the National Academy of Sciences 105, 19869-19874 (2008).
85. Worbs, T. et al. Oral tolerance originates in the intestinal immune system and relies on antigen carriage by dendritic cells. J. Exp. Med. 203, 519-527 (2006).
86. Inaba, K. et al. High Levels of a Major Histocompatibility Complex II–Self Peptide Complex on Dendritic Cells from the T Cell Areas of Lymph Nodes. J. Exp. Med. 186, 665-672 (1997).
87. Hawiger, D. et al. Dendritic Cells Induce Peripheral T Cell Unresponsiveness under Steady State Conditions in vivo. The Journal of Experimental Medicine 194, 769-780 (2001).
88. Liu, K. et al. Immune Tolerance After Delivery of Dying Cells to Dendritic Cells In Situ. J. Exp. Med. 196, 1091-1097 (2002).
89. Jonuleit, H., Schmitt, E., Schuler, G., Knop, J. & Enk, A. H. Induction of Interleukin 10–Producing, Nonproliferating Cd4+ T Cells with Regulatory Properties by Repetitive Stimulation with Allogeneic Immature Human Dendritic Cells. J. Exp. Med. 192, 1213-1222 (2000).
90. Bonifaz, L. et al. Efficient Targeting of Protein Antigen to the Dendritic Cell Receptor DEC-205 in the Steady State Leads to Antigen Presentation on Major Histocompatibility Complex Class I Products and Peripheral CD8+ T Cell Tolerance. The Journal of Experimental Medicine 196, 1627-1638 (2002).
91. Yamazaki, S. et al. CD8+CD205+ Splenic Dendritic Cells Are Specialized to Induce Foxp3+ Regulatory T Cells. The Journal of Immunology 181, 6923-6933 (2008).
92. Coombes, J. L. et al. A functionally specialized population of mucosal CD103+ DCs induces Foxp3+ regulatory T cells via a TGF-β– and retinoic acid–dependent mechanism. The Journal of Experimental Medicine 204, 1757-1764 (2007).
93. Sun, C. et al. Small intestine lamina propria dendritic cells promote de novo generation of Foxp3 T reg cells via retinoic acid. The Journal of Experimental Medicine 204, 1775-1785 (2007).
94. Guilliams, M. et al. Skin-draining lymph nodes contain dermis-derived CD103− dendritic cells that constitutively produce retinoic acid and induce Foxp3+ regulatory T cells. Blood 115, 1958-1968 (2010).
80
95. Gordon, J. R., Li, F., Nayyar, A., Xiang, J. & Zhang, X. CD8α+, but Not CD8α , Dendritic Cells Tolerize Th2 Responses via Contact-Dependent and -Independent Mechanisms, and Reverse Airway Hyperresponsiveness, Th2, and Eosinophil Responses in a Mouse Model of Asthma. The Journal of Immunology 175, 1516-1522 (2005).
96. Huang, H., Dawicki, W., Zhang, X., Town, J. & Gordon, J. R. Tolerogenic Dendritic Cells Induce CD4+CD25hiFoxp3+ Regulatory T Cell Differentiation from CD4+CD25−/loFoxp3− Effector T Cells. The Journal of Immunology 185, 5003-5010 (2010).
97. Steinbrink, K., Wolfl, M., Jonuleit, H., Knop, J. & Enk, A. Induction of tolerance by IL-10-treated dendritic cells. The Journal of Immunology 159, 4772-4780 (1997).
98. Svensson, M., Maroof, A., Ato, M. & Kaye, P. M. Stromal Cells Direct Local Differentiation of Regulatory Dendritic Cells. Immunity 21, 805-816 (2004).
99. Wakkach, A. et al. Characterization of Dendritic Cells that Induce Tolerance and T Regulatory 1 Cell Differentiation In vivo. Immunity 18, 605-617 (2003).
100. Pulendran, B., Tang, H. & Manicassamy, S. Programming dendritic cells to induce TH2 and tolerogenic responses. Nat. Immunol. 11, 647-655 (2010).
101. Brown, J. A. et al. Blockade of Programmed Death-1 Ligands on Dendritic Cells Enhances T Cell Activation and Cytokine Production. The Journal of Immunology 170, 1257-1266 (2003).
102. Freeman, G. J. et al. Engagement of the Pd-1 Immunoinhibitory Receptor by a Novel B7 Family Member Leads to Negative Regulation of Lymphocyte Activation. The Journal of Experimental Medicine 192, 1027-1034 (2000).
103. Keir, M. E. et al. Tissue expression of PD-L1 mediates peripheral T cell tolerance. J. Exp. Med. 203, 883-895 (2006).
104. Latchman, Y. et al. PD-L2 is a second ligand for PD-1 and inhibits T cell activation. Nat. Immunol. 2, 261-268 (2001).
105. Carter, L. L. et al. PD-1:PD-L inhibitory pathway affects both CD4+ and CD8+ T cells and is overcome by IL-2. European Journal of Immunology 32, 634 (2002).
106. Agata, Y. et al. Expression of the PD-1 antigen on the surface of stimulated mouse T and B lymphocytes. International Immunology 8, 765-772 (1996).
107. Bronte, V. & Zanovello, P. Regulation of immune responses by L-arginine metabolism. Nat Rev Immunol 5, 641-654 (2005).
108. MacMicking, J., Xie, Q. & Nathan, C. NITRIC OXIDE AND MACROPHAGE FUNCTION. Annu. Rev. Immunol. 15, 323-350 (1997; 1997).
81
109. Modolell, M., Corraliza, I. M., Link, F., Soler, G. & Eichmann, K. Reciprocal regulation of the nitric oxide synthase/arginase balance in mouse bone marrow‐derived macrophages by TH 1 and TH 2 cytokines. Eur. J. Immunol. 25, 1101-1104 (1995).
110. Munder, M., Eichmann, K. & Modolell, M. Alternative Metabolic States in Murine Macrophages Reflected by the Nitric Oxide Synthase/Arginase Balance: Competitive Regulation by CD4+ T Cells Correlates with Th1/Th2 Phenotype. The Journal of Immunology 160, 5347-5354 (1998).
111. Munder, M. et al. Th1/Th2-Regulated Expression of Arginase Isoforms in Murine Macrophages and Dendritic Cells. The Journal of Immunology 163, 3771-3777 (1999).
112. Bogdan, C. Nitric oxide and the immune response. Nat. Immunol. 2, 907-916 (2001).
113. Wei, X. et al. Altered immune responses in mice lacking inducible nitric oxide synthase. Nature 375, 408-411 (1995).
114. Gregory, S., Wing, E., Hoffman, R. & Simmons, R. Reactive nitrogen intermediates suppress the primary immunologic response to Listeria. The Journal of Immunology 150, 2901-2909 (1993).
115. Albina, J., Abate, J. & Henry, W. Nitric oxide production is required for murine resident peritoneal macrophages to suppress mitogen-stimulated T cell proliferation. Role of IFN-gamma in the induction of the nitric oxide-synthesizing pathway. The Journal of Immunology 147, 144-148 (1991).
116. Mazzoni, A. et al. Myeloid Suppressor Lines Inhibit T Cell Responses by an NO-Dependent Mechanism. The Journal of Immunology 168, 689-695 (2002).
117. Bingisser, R. M., Tilbrook, P. A., Holt, P. G. & Kees, U. R. Macrophage-Derived Nitric Oxide Regulates T Cell Activation via Reversible Disruption of the Jak3/STAT5 Signaling Pathway. The Journal of Immunology 160, 5729-5734 (1998).
118. Holt, P. G. et al. Downregulation of the antigen presenting cell function(s) of pulmonary dendritic cells in vivo by resident alveolar macrophages. J. Exp. Med. 177, 397-407 (1993).
119. Dinapoli, M. R., Calderon, C. L. & Lopez, D. M. The altered tumoricidal capacity of macrophages isolated from tumor-bearing mice is related to reduce expression of the inducible nitric oxide synthase gene. J. Exp. Med. 183, 1323-1329 (1996).
120. Hung, K. et al. The Central Role of CD4+ T Cells in the Antitumor Immune Response. J. Exp. Med. 188, 2357-2368 (1998).
121. Farias-Eisner, R., Sherman, M. P., Aeberhard, E. & Chaudhuri, G. Nitric oxide is an important mediator for tumoricidal activity in vivo. Proceedings of the National Academy of Sciences 91, 9407-9411 (1994).
82
122. Rodriguez, P. C. et al. Arginase I Production in the Tumor Microenvironment by Mature Myeloid Cells Inhibits T-Cell Receptor Expression and Antigen-Specific T-Cell Responses. Cancer Res. 64, 5839-5849 (2004).
123. Rodriguez, P. C. et al. L-Arginine Consumption by Macrophages Modulates the Expression of CD3ζ Chain in T Lymphocytes. The Journal of Immunology 171, 1232-1239 (2003).
124. Otsuji, M., Kimura, Y., Aoe, T., Okamoto, Y. & Saito, T. Oxidative stress by tumor-derived macrophages suppresses the expression of CD3 ζ chain of T-cell receptor complex and antigen-specific T-cell responses. Proceedings of the National Academy of Sciences 93, 13119-13124 (1996).
125. Rodriguez, P. C., Quiceno, D. G. & Ochoa, A. C. L-arginine availability regulates T-lymphocyte cell-cycle progression. Blood 109, 1568-1573 (2007).
126. Corzo, C. A. et al. HIF-1α regulates function and differentiation of myeloid-derived suppressor cells in the tumor microenvironment. The Journal of Experimental Medicine 207, 2439-2453 (2010).
127. Hwu, P. et al. Indoleamine 2,3-Dioxygenase Production by Human Dendritic Cells Results in the Inhibition of T Cell Proliferation. The Journal of Immunology 164, 3596-3599 (2000).
128. Uyttenhove, C. et al. Evidence for a tumoral immune resistance mechanism based on tryptophan degradation by indoleamine 2,3-dioxygenase. Nat. Med. 9, 1269-1274 (2003).
129. Munn, D. H. et al. Inhibition of  T Cell Proliferation by Macrophage Tryptophan Catabolism. J. Exp. Med. 189, 1363-1372 (1999).
130. Mellor, A. L. & Munn, D. H. Ido expression by dendritic cells: tolerance and tryptophan catabolism. Nat Rev Immunol 4, 762-774 (2004).
131. Mellor, A. L. et al. Cutting Edge: Induced Indoleamine 2,3 Dioxygenase Expression in Dendritic Cell Subsets Suppresses T Cell Clonal Expansion. The Journal of Immunology 171, 1652-1655 (2003).
132. Munn, D. H. et al. Expression of indoleamine 2,3-dioxygenase by plasmacytoid dendritic cells in tumor draining lymph nodes. the journal of clinical investigation (2004).
133. Munn, D. H. et al. GCN2 Kinase in T Cells Mediates Proliferative Arrest and Anergy Induction in Response to Indoleamine 2,3-Dioxygenase. Immunity 22, 633-642 (2005).
134. Frumento, G. et al. Tryptophan-derived Catabolites Are Responsible for Inhibition of T and Natural Killer Cell Proliferation Induced by Indoleamine 2,3-Dioxygenase. J. Exp. Med. 196, 459-468 (2002).
135. Saraiva, M. & O'Garra, A. The regulation of IL-10 production by immune cells. Nat Rev Immunol 10, 170-181 (2010).
83
136. Dillon, S. et al. A Toll-Like Receptor 2 Ligand Stimulates Th2 Responses In vivo, via Induction of Extracellular Signal-Regulated Kinase Mitogen-Activated Protein Kinase and c-Fos in Dendritic Cells. The Journal of Immunology 172, 4733-4743 (2004).
137. Ouyang, W., Rutz, S., Crellin, N. K., Valdez, P. A. & Hymowitz, S. G. Regulation and Functions of the IL-10 Family of Cytokines in Inflammation and Disease. Annu. Rev. Immunol. 29, 71-109 (2011; 2011).
138. Enk, A., Angeloni, V., Udey, M. & Katz, S. Inhibition of Langerhans cell antigen-presenting function by IL-10. A role for IL-10 in induction of tolerance. The Journal of Immunology 151, 2390-2398 (1993).
139. Del Prete, G. et al. Human IL-10 is produced by both type 1 helper (Th1) and type 2 helper (Th2) T cell clones and inhibits their antigen-specific proliferation and cytokine production. The Journal of Immunology 150, 353-360 (1993).
140. Groux, H., Bigler, M., de Vries, J. E. & Roncarolo, M. G. Interleukin-10 induces a long-term antigen-specific anergic state in human CD4+ T cells. J. Exp. Med. 184, 19-29 (1996).
141. de Waal Malefyt, R. et al. Interleukin 10 (IL-10) and viral IL-10 strongly reduce antigen-specific human T cell proliferation by diminishing the antigen-presenting capacity of monocytes via downregulation of class II major histocompatibility complex expression. J. Exp. Med. 174, 915-924 (1991).
142. Creery, W. D., Diaz-Mitoma, F., Filion, L. & Kumar, A. Differential modulation of B7-1 and B7-2 isoform expression on human monocytes by cytokines which influence the development of T helper cell phenotype. Eur. J. Immunol. 26, 1273-1277 (1996).
143. D'Andrea, A. et al. Interleukin 10 (IL-10) inhibits human lymphocyte interferon gamma-production by suppressing natural killer cell stimulatory factor/IL-12 synthesis in accessory cells. J. Exp. Med. 178, 1041-1048 (1993).
144. Fiorentino, D., Zlotnik, A., Mosmann, T., Howard, M. & O'Garra, A. IL-10 inhibits cytokine production by activated macrophages. The Journal of Immunology 147, 3815-3822 (1991).
145. Hart, P., Hunt, E., Bonder, C., Watson, C. & Finlay-Jones, J. Regulation of surface and soluble TNF receptor expression on human monocytes and synovial fluid macrophages by IL-4 and IL-10. The Journal of Immunology 157, 3672-3680 (1996).
146. Allavena, P. et al. IL-10 prevents the differentiation of monocytes to dendritic cells but promotes their maturation to macrophages. Eur. J. Immunol. 28, 359-369 (1998).
147. Takeda, K. et al. Enhanced Th1 Activity and Development of Chronic Enterocolitis in Mice Devoid of Stat3 in Macrophages and Neutrophils. Immunity 10, 39-49 (1999).
148. Gabrilovich, D. Mechanisms and functional significance of tumour-induced dendritic-cell defects. Nat Rev Immunol 4, 941-952 (2004).
84
149. Almand, B. et al. Clinical Significance of Defective Dendritic Cell Differentiation in Cancer. Clinical Cancer Research 6, 1755-1766 (2000).
150. Thurnher, M. et al. Human renal-cell carcinoma tissue contains dendritic cells. Int. J. Cancer 68, 1-7 (1996).
151. Palucka, K., Ueno, H. & Banchereau, J. Recent Developments in Cancer Vaccines. The Journal of Immunology 186, 1325-1331 (2011).
152. Troy, A. J., Summers, K. L., Davidson, P. J., Atkinson, C. H. & Hart, D. N. Minimal recruitment and activation of dendritic cells within renal cell carcinoma. Clinical Cancer Research 4, 585-593 (1998).
153. Gabrilovich, D. I., Corak, J., Ciernik, I. F., Kavanaugh, D. & Carbone, D. P. Decreased antigen presentation by dendritic cells in patients with breast cancer. Clinical Cancer Research 3, 483-490 (1997).
154. Gabrilovich, D. I., Ciernik, I. F. & Carbone, D. P. Dendritic Cells in Antitumor Immune Responses: I. Defective Antigen Presentation in Tumor-Bearing Hosts. Cell. Immunol. 170, 101-110 (1996).
155. Vicari, A. P. et al. Reversal of Tumor-induced Dendritic Cell Paralysis by CpG Immunostimulatory Oligonucleotide and Anti–Interleukin 10 Receptor Antibody. J. Exp. Med. 196, 541-549 (2002).
156. Gabrilovich, D. I. et al. Production of vascular endothelial growth factor by human tumors inhibits the functional maturation of dendritic cells. Nat. Med. 2, 1096-1103 (1996).
157. Labeur, M. S. et al. Generation of Tumor Immunity by Bone Marrow-Derived Dendritic Cells Correlates with Dendritic Cell Maturation Stage. The Journal of Immunology 162, 168-175 (1999).
158. Chaux, P., Favre, N., Martin, M. & Martin, F. Tumor-infiltrating dendritic cells are defective in their antigen-presenting function and inducible B7 expression in rats. Int. J. Cancer 72, 619-624 (1997).
159. Kiertscher, S. M., Luo, J., Dubinett, S. M. & Roth, M. D. Tumors Promote Altered Maturation and Early Apoptosis of Monocyte-Derived Dendritic Cells. The Journal of Immunology 164, 1269-1276 (2000).
160. Diao, J., Zhao, J., Winter, E. & Cattral, M. S. Recruitment and Differentiation of Conventional Dendritic Cell Precursors in Tumors. The Journal of Immunology 184, 1261-1267 (2010).
161. Diao, J., Zhao, J., Winter, E. & Cattral, M. S. Tumors Suppress In Situ Proliferation of Cytotoxic T Cells by Promoting Differentiation of Gr-1+ Conventional Dendritic Cells through IL-6. The Journal of Immunology 186, 5058-5067 (2011).
85
162. Park, S. et al. IL-6 Regulates In vivo Dendritic Cell Differentiation through STAT3 Activation. The Journal of Immunology 173, 3844-3854 (2004).
163. Gabrilovich, D. I., Ishida, T., Nadaf, S., Ohm, J. E. & Carbone, D. P. Antibodies to Vascular Endothelial Growth Factor Enhance the Efficacy of Cancer Immunotherapy by Improving Endogenous Dendritic Cell Function. Clinical Cancer Research 5, 2963-2970 (1999).
164. Herber, D. L. et al. Lipid accumulation and dendritic cell dysfunction in cancer. Nat. Med. 16, 880-886 (2010).
165. Bronte, V. et al. Identification of a CD11b+/Gr-1+/CD31+ myeloid progenitor capable of activating or suppressing CD8+T cells. Blood 96, 3838-3846 (2000).
166. Gabrilovich, D. I. & Nagaraj, S. Myeloid-derived suppressor cells as regulators of the immune system. Nat Rev Immunol 9, 162-174 (2009).
167. Kusmartsev, S. & Gabrilovich, D. I. Inhibition of myeloid cell differentiation in cancer: the role of reactive oxygen species. J. Leukoc. Biol. 74, 186-196 (2003).
168. Almand, B. et al. Increased Production of Immature Myeloid Cells in Cancer Patients: A Mechanism of Immunosuppression in Cancer. The Journal of Immunology 166, 678-689 (2001).
169. Youn, J., Nagaraj, S., Collazo, M. & Gabrilovich, D. I. Subsets of Myeloid-Derived Suppressor Cells in Tumor-Bearing Mice. The Journal of Immunology 181, 5791-5802 (2008).
170. Movahedi, K. et al. Identification of discrete tumor-induced myeloid-derived suppressor cell subpopulations with distinct T cell–suppressive activity. Blood 111, 4233-4244 (2008).
171. Nagaraj, S., Schrum, A. G., Cho, H., Celis, E. & Gabrilovich, D. I. Mechanism of T Cell Tolerance Induced by Myeloid-Derived Suppressor Cells. The Journal of Immunology 184, 3106-3116 (2010).
172. Kusmartsev, S. & Gabrilovich, D. I. STAT1 Signaling Regulates Tumor-Associated Macrophage-Mediated T Cell Deletion. The Journal of Immunology 174, 4880-4891 (2005).
173. Gallina, G. 1. et al. Tumors induce a subset of inflammatory monocytes with immunosuppressive activity on CD8+ T cells. [Article].
174. Sinha, P., Clements, V. K., Bunt, S. K., Albelda, S. M. & Ostrand-Rosenberg, S. Cross-Talk between Myeloid-Derived Suppressor Cells and Macrophages Subverts Tumor Immunity toward a Type 2 Response. The Journal of Immunology 179, 977-983 (2007).
86
175. Corzo, C. A. et al. Mechanism Regulating Reactive Oxygen Species in Tumor-Induced Myeloid-Derived Suppressor Cells. The Journal of Immunology 182, 5693-5701 (2009).
176. Bronte, V. et al. IL-4-Induced Arginase 1 Suppresses Alloreactive T Cells in Tumor-Bearing Mice. The Journal of Immunology 170, 270-278 (2003).
177. Kusmartsev, S., Nefedova, Y., Yoder, D. & Gabrilovich, D. I. Antigen-Specific Inhibition of CD8+ T Cell Response by Immature Myeloid Cells in Cancer Is Mediated by Reactive Oxygen Species. The Journal of Immunology 172, 989-999 (2004).
178. Sica, A., Allavena, P. & Mantovani, A. Cancer related inflammation: The macrophage connection. Cancer Lett. 267, 204-215 (2008).
179. Lin, E. Y., Nguyen, A. V., Russell, R. G. & Pollard, J. W. Colony-Stimulating Factor 1 Promotes Progression of Mammary Tumors to Malignancy. J. Exp. Med. 193, 727-740 (2001).
180. Goswami, S. et al. Macrophages Promote the Invasion of Breast Carcinoma Cells via a Colony-Stimulating Factor-1/Epidermal Growth Factor Paracrine Loop. Cancer Res. 65, 5278-5283 (2005).
181. Smith, H. O. et al. The role of colony-stimulating factor 1 and its receptor in the etiopathogenesis of endometrial adenocarcinoma. Clinical Cancer Research 1, 313-325 (1995).
182. Biswas, S. K. & Mantovani, A. Macrophage plasticity and interaction with lymphocyte subsets: cancer as a paradigm. Nat. Immunol. 11, 889-896 (2010).
183. Schutyser, E. et al. Identification of Biologically Active Chemokine Isoforms from Ascitic Fluid and Elevated Levels of CCL18/Pulmonary and Activation-regulated Chemokine in Ovarian Carcinoma. J. Biol. Chem. 277, 24584-24593 (2002).
184. Giaccia, A., Siim, B. G. & Johnson, R. S. HIF-1 as a target for drug development. Nat Rev Drug Discov 2, 803-811 (2003).
185. Colombo, M. P. & Mantovani, A. Targeting Myelomonocytic Cells to Revert Inflammation-Dependent Cancer Promotion. Cancer Res. 65, 9113-9116 (2005).
186. Kuang, D. et al. Activated monocytes in peritumoral stroma of hepatocellular carcinoma foster immune privilege and disease progression through PD-L1. J. Exp. Med. 206, 1327-1337 (2009).
187. Schioppa, T. et al. Regulation of the Chemokine Receptor CXCR4 by Hypoxia. J. Exp. Med. 198, 1391-1402 (2003).
188. Talks, K. L. et al. The Expression and Distribution of the Hypoxia-Inducible Factors HIF-1α and HIF-2α in Normal Human Tissues, Cancers, and Tumor-Associated Macrophages. The American Journal of Pathology 157, 411-421 (2000).
87
189. Lewis, J. S., Landers, R. J., Underwood, J. C. E., Harris, A. L. & Lewis, C. E. Expression of vascular endothelial growth factor by macrophages is up‐regulated in poorly vascularized areas of breast carcinomas. J. Pathol. 192, 150-158 (2000).
190. Eubank, T. D., Galloway, M., Montague, C. M., Waldman, W. J. & Marsh, C. B. M-CSF Induces Vascular Endothelial Growth Factor Production and Angiogenic Activity From Human Monocytes. The Journal of Immunology 171, 2637-2643 (2003).
191. Clear, A. J. et al. Increased angiogenic sprouting in poor prognosis FL is associated with elevated numbers of CD163+ macrophages within the immediate sprouting microenvironment. Blood 115, 5053-5056 (2010).
192. Lin, E. Y. et al. Macrophages Regulate the Angiogenic Switch in a Mouse Model of Breast Cancer. Cancer Res. 66, 11238-11246 (2006).
193. Grivennikov, S. I., Greten, F. R. & Karin, M. Immunity, Inflammation, and Cancer. Cell 140, 883-899 (2010).
194. McLellan, A. D., Starling, G. C. & Hart, D. N. J. Isolation of human blood dendritic cells by discontinuous Nycodenz gradient centrifugation. J. Immunol. Methods 184, 81-89 (1995).
195. Selenko-Gebauer, N. et al. B7-H1 (Programmed Death-1 Ligand) on Dendritic Cells Is Involved in the Induction and Maintenance of T Cell Anergy. The Journal of Immunology 170, 3637-3644 (2003).
196. Tuettenberg, A. et al. The Role of ICOS in Directing T Cell Responses: ICOS-Dependent Induction of T Cell Anergy by Tolerogenic Dendritic Cells. The Journal of Immunology 182, 3349-3356 (2009).
197. RodrÃ-guez, P. C. & Ochoa, A. C. Arginine regulation by myeloid derived suppressor cells and tolerance in cancer: mechanisms and therapeutic perspectives. Immunol. Rev. 222, 180-191 (2008).
198. Chen, M. & Wang, J. Programmed cell death of dendritic cells in immune regulation. Immunol. Rev. 236, 11-27 (2010).
199. Chen, M. et al. Dendritic Cell Apoptosis in the Maintenance of Immune Tolerance. Science 311, 1160-1164 (2006).
200. Lee, Y. K. et al. Late Developmental Plasticity in the T Helper 17 Lineage. Immunity 30, 92-107 (2009).
201. Sharma, M. D. et al. Indoleamine 2,3-dioxygenase controls conversion of Foxp3+ Tregs to TH17-like cells in tumor-draining lymph nodes. Blood 113, 6102-6111 (2009).
202. Xu, L., Kitani, A., Fuss, I. & Strober, W. Cutting Edge: Regulatory T Cells Induce CD4+CD25–Foxp3– T Cells or Are Self-Induced to Become Th17 Cells in the Absence of Exogenous TGFbeta. The Journal of Immunology 178, 6725-6729 (2007).
88
203. Koenen, H. J. P. M. et al. Human CD25highFoxp3pos regulatory T cells differentiate into IL-17–producing cells. Blood 112, 2340-2352 (2008).
204. Watkins, S. K., Egilmez, N. K., Suttles, J. & Stout, R. D. IL-12 Rapidly Alters the Functional Profile of Tumor-Associated and Tumor-Infiltrating Macrophages In vitro and In vivo. The Journal of Immunology 178, 1357-1362 (2007).
205. Heusinkveld, M. et al. M2 Macrophages Induced by Prostaglandin E2 and IL-6 from Cervical Carcinoma Are Switched to Activated M1 Macrophages by CD4+ Th1 Cells. The Journal of Immunology 187, 1157-1165 (2011).
206. Caux, C., Dezutter-Dambuyant, C., Schmitt, D. & Banchereau, J. GM-CSF and TNF-[alpha] cooperate in the generation of dendritic Langerhans cells. Nature 360, 258-261 (1992).
207. Auffermann-Gretzinger, S., Keeffe, E. B. & Levy, S. Impaired dendritic cell maturation in patients with chronic, but not resolved, hepatitis C virus infection. Blood 97, 3171-3176 (2001).
208. Serhan, C. N. & Savill, J. Resolution of inflammation: the beginning programs the end. Nat. Immunol. 6, 1191-1197 (2005).
209. Zorro Manrique, S. et al. Foxp3-positive macrophages display immunosuppressive properties and promote tumor growth. J. Exp. Med. 208, 1485-1499 (2011).
210. Shortman, K. & Liu, Y. Mouse and human dendritic cell subtypes. Nat Rev Immunol 2, 151-161 (2002).
211. Gordon, S. & Taylor, P. R. Monocyte and macrophage heterogeneity. Nat Rev Immunol 5, 953-964 (2005).
212. Zhang, M. et al. Splenic stroma drives mature dendritic cells to differentiate into regulatory dendritic cells. Nat Immunology 5, 1124-1133 (2004).
213. Jameson, S. Maintaining the norm: T cell homeostasis. Nat Rev Immunol 2, 547-556 (2002).
214. Huang, B. et al. GR-1+ CD115+ immature myeloid suppressor cells mediate the development of tumor-induced T regulatory cells and T cell anergy in tumor-bearing hosts. Cancer Research 66, 1123-1131 (2006).
215. Gabrilovich, D. et al. Mechanism of immune dysfunction in cancer mediated by immature GR-1+ myeloid cells. The Journal of Immunology 166, 5398-5406 (2001).
216. Herber, D. et al. Lipid accumulation and dendritic cell dysfunction in cancer. Nature Medicine 16, 880-887 (2010).
217. Wirtz, S. et al. Chemically induced mouse models of intestinal inflammation. Nature Protocols 2, 541-546 (2007).
89
218. Allan, R. et al. Epidermal viral immunity is induced by CD8α+ dendritic cells but not by langerhan cells. Science 301, 1925-1928 (2003).
219. Galli, S. J. et al.Phenotypic and functional plasticity of cells of innate immunity: macrophages, mast cells and neutrophils. Nature Immunology 12, 1035-1044 (2011).
220. Dranoff, G. Cytokines in cancer pathogenesis and cancer therapy. Nature Reveiws Cancer 4, 11-22 (2004).
221. CML Trialists Colaborative Group. Interferon alpha versus chemotherapy for chronic myeloid leukemia: a meta-analysis of seven randomized trials. Journal of the National Cancer Institute. 89, 1616-1620 (1997).
222. Fyfe, G. et al. Results of treatment of 255 patients with metastatic renal cell carcinoma who received high-dose recombinant interleukin-2 therapy. Journal of Clinical Oncology. 13, 688-696 (1995).
223. Lukacs-Kornek, V. et al. Regulated release of nitric oxide by non-hematopoietic stroma controls expansion of activated T cell pool in lymph nodes. Nat Imm. 12, 1096-1105 (2011).
224. Soiffer, R. et al. Vaccination with autologous melanoma cells engineered to secrete human granulocyte–macrophage colony-stimulating factor generates potent antitumor immunity in patients with metastatic melanoma. Proc. Natl. Acad. Sci. USA. 95, 13141-13146 (1998).
225. Soiffer, R. et al. Vaccination with irradiated, autologous melanoma cells engineered to secrete granulocyte-macrophage colony-stimulating factor by adenoviral-mediated gene transfer augments antitumor immunity in patients with metastatic melanoma. J. Clin. Oncol. 21, 3343-50 (2003).
226. Salgia, R. et al. Vaccination with irradiated autologous tumor cells engineered to secrete granulocyte-macrophage colony-stimulating factor augments antitumor immunity in some patients with metastatic non-small-cell lung carcinoma. J. Clin. Oncol. 21, 624-630 (2003).
227. Emens, A. L. et al. Timed sequential treatment with cyclophosphamide, doxorubicin and an allogeneic granulocyte-macrophage colony-stimulating factor-secreting breast tumor vaccine: a chemotherapy dose-ranging factorial study of safety and immune activation. J. Clin. Oncol. 27, 5911-5918 (2009).
228. Gupta, A. and Emmens, A. L. GM-CSF-secreting vaccines for solid tumors: moving forward. Discov Med. 10, 52-60 (2010).
229. Kusmartsev, S. et al. Tumor-associated CD8+ T cell tolerance induced by bone marrow derived immature myeloid cells. J. Immunol. 175, 4583-4592 (2005).