1. health and fitness
2. disease

Activation of the Cellular Immune Response in
Drosophila melanogaster Larvae
Ines Anderl
Department of Molecular Biology
Umeå University, Umeå, Sweden 2015
Responsible publisher under swedish law: the Dean of the Medical Faculty
This work is protected by the Swedish Copyright Legislation (Act 1960:729)
ISBN: 978-91-7601-317-5
ISSN: 0346-6612-1741
Picture top left: Drosophila blood cells; top right: encapsulated wasp egg; bottom:
wasp-infected Drosophila larva; back cover: plasmatocytes (green) and
lamellocytes (red)
Electronic version available at http://umu.diva-portal.org/
Printed by: Print & Media, Umeå University
Umeå, Sweden 2015
“Time flies like an arrow; fruit flies like a banana.”
Groucho Marx
Table of Contents
Table of Contents............................................................. i Abstract ......................................................................... iii Papers included in the thesis ......................................... iv Background ..................................................................... 1 Introduction ................................................................................... 1 Drosophila blood cells ................................................................... 2 Drosophila hematopoiesis ............................................................. 3 Lymph gland hematopoiesis .............................................................................. 3 Peripheral hematopoiesis ................................................................................... 4 Adult hematopoiesis .............................................................................................. 6 Immune response to parasitoid wasps .......................................... 7 Drosophila as a research tool ........................................................ 8 Genetic Screens in Drosophila with focus on hemocytes ............. 9 Methods for hemocyte research .................................................. 10 Aims of the thesis .......................................................... 13 Article I ......................................................................... 14 “A directed screen for genes involved in Drosophila blood cell
activation.” ........................................................................................ 14 Preface .......................................................................................... 14 Summary of main results............................................................. 14 Discussion .................................................................................... 14 Article II ........................................................................ 16 “Infection-induced proliferation is a central hallmark of the
activation of the cellular immune response in Drosophila larvae.” 16 Preface and study design ............................................................. 16 Summary of main results............................................................. 17 Discussion .................................................................................... 17 Article III....................................................................... 21 “Genetic screen in Drosophila larvae links ird1 function to Toll
signaling in the fat body and hemocyte motility.” ........................... 21 Preface .......................................................................................... 21 Summary of main results............................................................. 21 Discussion .................................................................................... 21 Article IV ...................................................................... 24 “Control of Drosophila blood cell activation via Toll signaling in
the fat body.” ..................................................................................... 24 Preface .......................................................................................... 24 Summary of main results............................................................. 24 i
Discussion .................................................................................... 24 Acknowledgements .......................................................27 References ................................................................... 28 ii
Abstract
During the last 40 years, Drosophila melanogaster has become an invaluable tool
in understanding innate immunity. The innate immune system of Drosophila
consists of a humoral and a cellular component. While many details are known about
the humoral immune system, our knowledge about the cellular immune system is
comparatively small. Blood cells or hemocytes constitute the cellular immune system.
Three blood types have been described for Drosophila larvae. Plasmatocytes are
phagocytes with a plethora of functions. Crystal cells mediate melanization and
contribute to wound healing. Plasmatocytes and crystal cells constitute the blood cell
repertoire of a healthy larva, whereas lamellocytes are induced in a demand-adapted
manner after infection with parasitoid wasp eggs. They are involved in the melanotic
encapsulation response against parasites and form melanotic nodules that are also
referred to as tumors.
In my thesis, I focused on unraveling the mechanisms of how the immune system
orchestrates the cellular immune response. In particular, I was interested in the
hematopoiesis of lamellocytes.
In Article I, we were able to show that ectopic expression of key components of a
number of signaling pathways in blood cells induced the development of lamellocytes,
led to a proliferative response of plasmatocytes, or to a combination of lamellocyte
activation and plasmatocyte proliferation.
In Article II, I combined newly developed fluorescent enhancer-reporter
constructs specific for plasmatocytes and lamellocytes and developed a “dual reporter
system” that was used in live microscopy of fly larvae. In addition, we established
flow cytometry as a tool to count total blood cell numbers and to distinguish between
different blood cell types. The “dual reporter system” enabled us to differentiate
between six blood cell types and established proliferation as a central feature of the
cellular immune response. The combination flow cytometry and live imaging
increased our understanding of the tempo-spatial events leading to the cellular
immune reaction.
In Article III, I developed a genetic modifier screen to find genes involved in the
hematopoiesis of lamellocytes. I took advantage of the gain-of-function phenotype of
the Tl10b mutation characterized by an activated cellular immune system, which
induced the formation blood cell tumors. We screened the right arm of chromosome
3 for enhancers and suppressors of this mutation and uncovered ird1.
Finally in Article IV, we showed that the activity of the Toll signaling pathway in
the fat body, the homolog of the liver, is necessary to activate the cellular immune
system and induce lamellocyte hematopoiesis.
iii
Papers included in the thesis
This thesis is based on the following publications, which will be referred to
by roman numerals (I - IV). All publications are reproduced with the
permission from the publishers.
Article I
Zettervall, C.J., Anderl, I., Williams, M.J., Palmer, R., Kurucz, E., Ando, I.,
and Hultmark, D. (2004). A directed screen for genes involved in Drosophila
blood cell activation. Proc. Natl. Acad. Sci. USA 101, 14192-14197.
Article II
Anderl, I., Vesala, L., Ihalainen, T., Vanha-aho, L.-M., Rämet, M., and
Hultmark, D. (2015). Infection-induced proliferation is a central hallmark of
the activation of the cellular immunes response in Drosophila larvae.
(Manuscript, I. Anderl and L. Vesala share first authorship)
Article III
Schmid, M.R., Anderl, I., Vesala, L., Vo, H., Valanne, S., Yang, H.,
Kronhamn, J., Rämet, M., Rusten, T.E., Hultmark, D. (2015). Genetic screen
in Drosophila larvae links ird1 function to Toll signaling in the fat body and
hemocyte motility. (Manuscript)
Article IV
Schmid, M.R., Anderl, I., Vesala, L., Vanha-aho, L.-M., Deng, X.-J., Rämet,
M., and Hultmark, D. (2014). Control of Drosophila blood cell activation via
Toll signaling in the fat body. PLoS One 9, e102568. (M. Schmid and I.
Anderl share first authorship.)
iv
Background
Introduction
Every organism, regardless of size and complexity, must protect itself against
assaults from non-self to “live long and prosper”. The discipline of immunology was
founded by Elie Metchnikoff and Paul Ehrlich who were awarded the Nobel Prize in
1908 “in recognition for their work on immunity” Metchnikoff´s discovery of
phagocytosis spawned innate immunity and Ehrlich´s side-chain theory adaptive
immunity [1].
The first frontline to infection are physical barriers. The skin protects the outside
of our bodies and mucosal tissues, such as lung, airways, and gut the inside. Mucous
tissues secrete fluids like tears, saliva, or mucus that contain compounds that restrict
the growth of microorganisms. Moreover, mucosal tissues are colonized by immune
cells. Two forms of immunity exist - the innate and the adaptive immunity. Innate
immunity is fast and general and has a cellular and humoral component. Pattern
recognition receptors (PRRs) expressed by innate immune cells recognize pathogenassociated molecular patterns (PAMPs) on the surface of intruders [2], [3] and
phagocytize them. In mammals, the complement system constitutes the humoral arm
of innate immunity. Their adaptive immunity comprises B and T lymphocytes, is
highly specific, and confers immunological memory. Autoreactive B and T cells are
eliminated during their development. The innate and adaptive immune system
cooperate: “There is no antigen-specific immunity without “instruction” by the innate
immune system and no efficacious innate immune response without the guidance of
acquired immunity” [1].
Mammalian blood cells are born in primary lymphoid tissues, the bone marrow
and the thymus. They develop form multipotent hematopoietic stem cells by
hierarchical lineage restriction (reviewed in [4]). This view has recently been
challenged by the heterogeneity of tissue macrophages (reviewed in [5]). The adaptive
immune response is generated in the secondary lymphoid tissues, the lymph nodes,
the spleen, and the mucosa-associated lymphoid tissue (MALT).
In the genomic era much effort has been dedicated to elucidating the origin of the
immune system. It has become clear that the immune system evolved in cooperation
with the commensal microbiota [6]. Notably, our gut microbiota shape our adaptive
immune system (reviewed in [7]). And also in the fly, the intestinal homeobox gene
Caudal represses the Toll-dependent expression of antimicrobial peptides. If this
regulation is abrogated, the commensal flora of the fly gut becomes unbalanced
resulting in the death of the animal [8]. Vertebrates, which comprise only a small
group within the animal kingdom, are so far the only group of animals known to have
adaptive immunity, but innate immunity is ubiquitious. From an evolutionary
perspective, insects are the most successful class in the animal kingdom representing
approximately half of all existing animal species [9], however, they only have an
innate immune system. It has been argued that the advantage of the specificity of the
immune reaction conferred by adaptive immunity might be in saving energy
resources [10].
The innate immune system of insects consists of a humoral and cellular arm. The
fat body, the homolog of the liver, is the central organ in secreting humoral
components. Blood cells or hemocytes are the effector cells of the cellular arm of the
immune system. Drosophila melanogaster has become the state of the art model
organism for innate immunity research during the last 40 years (reviewed in [11]).
Work on the humoral immune system of the cecropia moth led to the discovery of
antimicrobial peptides that are secreted by the fat body into the hemolymph in
response to bacterial or fungal infection (reviewed in [12]). In 1972, Bertil Rasmusons
lab showed for the first time that antimicrobial activity is inducible in Drosophila
1
[13]. Ingrid Faye´s group reported in 1992 that in the cecropia moth a nuclear
immuneresponsive factor binds to NFκB-like motifs [14]. In 1993 the Engström,
Faye, and Hultmark labs jointly reported for the first time that κB-like motifs regulate
the induction of immune genes in Drosophila [15]. And in 1995 the Hultmark lab
showed that Toll signaling is able to introduce Cecropin gene expression [16]. In 1996
Bruno Lemaitre in the Hoffmann lab reported that Micrococcus-induced Drosomycin
expression is dependent on Toll signaling [17]. These and other findings spawned a
renewed interest in innate immunity (reviewed in [11]). Jules Hoffman and Bruce
Beutler were jointly awarded the Nobel Prize for Physiology and Medicine in 2011 “for
their discoveries concerning the activation of innate immunity”
Analogous to our skin, the cuticle of the exoskeleton of insects is the first physical
barrier to infection. But in contrast to vertebrates, insects have an open circulatory
system, which needs quick repair when injured to prevent bleeding to death.
Therefore, the hemolymph harbors an efficient clotting or coagulation system to
minimize bleeding and to prevent invading bacteria from spreading through the
hemolymph [18], [19]. A quick and efficient wound healing response takes care of
tissue repair [20], [21]. Wound healing and hemolymph clotting are complemented
by melanization. Phenol oxidases (POs) induce the rapid polymerization of quinones
that result in the production of the black-brown pigment melanin at the site of injury.
Melanin traps bacteria in nodules and contributes to clot formation [22]. Melanin
itself and byproducts of the melanization cascade possess antimicrobial activity [23],
[22]. Finally, encapsulation is an immune response dedicated to the killing of
parasitoids that are too big for phagocytosis like the eggs of parasitoid wasps
(reviewed in [24], [25]). Melanization and encapsulation are immune reactions
specific to arthropods (reviewed in [19], [26]).
Blood cells constitute the cellular arm of innate immunity. This thesis is dedicated
to hemocytes, specifically to lamellocytes. Therefore I primarily focus on the
development and function of peripheral blood cells in the larva.
Drosophila blood cells
The cellular immune system of insects is represented by blood cells or hemocytes.
As insects do not have adaptive immunity, their hemocytes bear resemblance to
mammalian blood cells of the myloid lineage (reviewed in [27]). In Drosophila
melanogaster three different hemocyte types can be found: plasmatocytes, crystal
cells, and lamellocytes. Plasmatocytes and crystal cells are present in all
developmental stages of the fruit fly (reviewed in [28]), whereas lamellocytes are
induced only in response to infection by pathogens to large to phagocytize (reviewed
in [25]). In response to parasitoid wasp infection, lamellocyte numbers rise swiftly to
encapsulate and thereby kill the intruder [29].
In unchallenged and healthy animals, plasmatocytes are the most numerous cell
type constituting 90 to 95 % of all blood cells [30], [31]. They are small spherical cells
with a diameter up to 12 μm [30], [32]. Plasmatocytes are the workhorse of the
cellular immune systems with various functions in immunity and development. They
phagocytize apoptotic tissue and bacteria (reviewed in for example [28]), participate
in the encapsulation response (reviewed in [25]), patrol the hemolymph as sentinels
[33], synthesize antimicrobial peptides [34], cytokines [35], [36], [37], [38], [39], and
extracellular matrix [40], [41], [42], transdifferentiate into crystal cells [43] and
lamellocytes [44], [45], [46], are involved in the priming of the immune response
against bacteria [47], contribute to tissue remodeling [48], [35], regeneration, and
homeostatis [49], [50], [39]. Despite the plethora of functions, plasmatocyte
depletion in larvae is not necessary for survival [51].
The crystals within the cytoplasm of crystal cells were the inspiration for the
naming of this cell type. They are spherical, slightly bigger than plasmatocytes, and
have a golden hue in phase contrast microscopy. Crystal cells are dedicated to
2
melanization upon wounding and bacterial infection [52]. At the wound site, they
contribute to scab formation [20]. Crystal cells express two of the three PPO
(prophenol oxidase) genes of the Drosophila melanogaster genome. PPO1
(CG42639) is quickly released into the hemolymph after wounding. PPO2 (CG8913)
is stored within the crystals and only released after crystal cell rupture [52]. PPOrelease after crystal cells rapture is controlled by the Jun kinase basket (bsk),
RhoGTPases, and the Drosophila TNF homolog eiger (egr) [53]. PPO1 and PPO2
require proteolytic cleavage for their activation [54]. PPO1 is cleaved by the CLIP
serine protease Hayan [55]. The third PPO encoded in the Drosophila genome is
PPO3 (CG42640). PPO3 is lamellocyte-specific, does not require proteolytic cleavage
for activation, and is not sufficient to melanize encapsulated wasp eggs in a double
mutant background of PPO1 and PPo2 [52].
Lamellocytes are large discoid cells that are only 1 μm thick [30], [32], [31]. They
are effector cells only produced by the immune system when parasitoid wasps have
deposited their eggs inside the Drosophila host [56], or at aberrant conditions, when
they participate in forming melanotic nodules (reviewed in [57]).
Drosophila hematopoiesis
Hematopoiesis between Drosophila and vertebrates is evolutionary conserved
[27]. Blood cells hematopoiesis occurs in waves in the embryo [27], in the larva [58],
and in the adult fly [59]. In the Drosophila embryo, plasmatocytes and crystal cells
are born and proliferate in the head mesoderm. Crystal cells never leave this site
during embryonic development, whereas plasmatocytes migrate through the embryo
as macrophages and phagocytize apoptotic tissue [60], [61]. The embryonic
macrophages colonize the hemocoel of the Drosophila larva as peripheral hemocytes
and persist till the adult fly [62]. In the larval hemocoel, peripheral blood cells either
circulate or are sessile [31]. The sessile hemocyte compartment is now regarded as a
hematopoietic niche [63], [64], [43].
The lymph gland arises from the thoracic mesoderm [62]. The lymph glands have
long since been regarded the hematopoietic organ of the larva [65]. They grow
throughout larval development and disintegrate at the beginning of metamorphosis
[66]. Cells derived from the secondary lymph gland lobes home to hematopoitic
clusters in the abdomen of the fly [59].
Lymph gland hematopoiesis
During the last 15 years, the lymph glands have become an excellent model for
hematopoiesis. The lymph glands consist of several paired lobes adjacent to the heart
(Figure 1). The primary lymph gland lobes consist of distinct zones. Quiescent
hematopoietic stem cells, the prohemocytes, reside in the medullary zone (MZ), and
differentiated hemocytes are found in the cortical zone (CZ). The posterior signaling
center (PSC) constitutes the hematopoietic niche that controls the balance between
the prohemocytes and differentiating hemocytes in the medullary zone of the lymph
gland (Figure 1) [67].
3
CZ
MZ
PSC
2nd
lobes
crystal cell
plasmatocyte
PSC cell
prohemocyte
pericardial cell
Figure 1 Mature lymph gland (CZ –
Cortical Zone, MZ – Medullary Zone,
PSC – Posterior Signaling Center)
Peripheral hematopoiesis
It has previously been assumed that lymph glands are the hematopoietic organ
that gives rise to all larval blood cells [27]. However, transplantation experiments
from the hemocyte anlage at the blastoderm stage revealed that embryonic
plasmatocytes are the founder population of peripheral larval hemocytes and that
they persist to the imagines [62]. Two peripheral larval blood cell populations have
been described, the circulating and the sessile cells (Figure 2).
Circulating hemocytes mostly consist of plasmatocytes. Depending on genetic
background, they are estimated to account for 90 to 95 % and crystal cells for the
remaining 5 to 10 % [30]). Also a low number of prohemocytes was observed in
circulation [32], [31], [63]. Circulating hemocyte numbers during the larval stage
increase from fewer than 200 at the beginning of the first instar to more than 6000
cells at the end of the third larval instar [30], [31], [64]. Sessile hemocytes account for
approximately the same number as circulating cells because in the eater1 mutant,
where sessile cells are absent, circulating cell numbers double in comparison to wildtype controls [68]. Whereas Lanot et al. estimated that at least one third of the blood
cells are attached to the integument [31]. Circulating hemocytes are regarded as
sentinels that passively patrol the hemocoel propelled by the hemolymph flow.
During wound healing, they are recruited by “direct capture“ from the hemolymph
[33].
Sessile cells attach to larval tissues and comprise plasmatocytes and crystal cells
(Figure 1A). Primarily they adhere to the dorsal integument in a repeated segmented
pattern [31], [69], [70] where they reside and proliferate in-between the epidermis
and muscles tightly associated with neurons of the peripheral nervous system. The
peripheral neurons are thought to provide a trophic environment of yet unknown
growth factors, and therefore these epidermal-muscular pockets are regarded as a
hematopoietic niche for circulating and sessile hemocytes [64]. When embryonic
hemocytes first colonize the larval hemocoel, they accumulate on the integument of
the two posteriormost abdominal segments. In later larval stages, the density of
4
A
Subepithelial
hemocyte patches
Lymph
glands
Posterior hematopoietic tissue (PHT)
non-infected
Plasmatocytes
Crystal cells
B
Secondary lymph
gland lobes
Encapsulated
parasite eggs
infected
Plasmatocytes
Lamellocytes
Figure 2 Hemocytes and hematopoietic compartments in the Drosophila larva
In (A) non-infected and (B) wasp-infected animals (published by Anderl &
Hultmark 2015 [71] and presented here with small modifications)
sessile cells is highest in this part of the larva [32], [72]. It has been suggested to refer
to these cells as the posterior hematopoietic tissue (PHT) [72]. In addition, hemocytes
can also be found adhering to imaginal discs [72], [62] and elsewhere.
Blood cells cycle between sessile and circulating states [33], [73], [64]. The
expression of the Nimrod receptor eater in plasmatocytes mediates the adhesion to
the sessile compartment, however the nature of the eater ligand is still elusive [68]. It
is possible that eater binds to a component of the basement membrane, as it was
shown that Kc167 cells bind to lamin in a syndecan-dependent manner and
hemocytes adhere to lamin in vivo [74].
Mature plasmatocytes proliferate by self-renewel in circulation [30], [64] and in
the epidermal-muscular pockets where their proliferation rate is higher [64].
Plasmatocytes express croquemort (crq), Peroxidasin (Pxn), eater (reviewed in [75],
and u-shaped (ush) [76], [45]. Misexpression of Pvf2 [77], Egfr, and Ras85D [70] in
hemocytes induce proliferation of plasmatocytes.
Crystal cell numbers also increase during larval stages. Paradoxically, all reports
agreed so far that mature crystal cells do not divide [30], [31]. Instead, plasmatocytes
transdifferentiate into crystal cells in the sessile compartment in a Notch-Serrate
dependent process that requires cell-to cell contacts [43], [78]. However, in eater1
mutants crystal cells do develop. Clumps of crystal cells and lamellocytes are
observed in tight association in circulation [68]. This suggests that hemocyte-tohemocyte contacts are more important than growth factors from the hematopoietic
niche.
The enormous plasticity of the cellular immune system becomes evident during
immune reactions to wasp infection or at aberrant genetic conditions that lead to socalled melanotic nodules. A new blood cell type, the lamellocyte, is generated.
5
Lamellocytes are not present in healthy Drosophila larvae. Therefore, they can only
be studied when the larva is immunocompromised. It has been debated whether
lamellocytes develop only in the lymph glands from prohemocytes and are released
into circulation, or if mature plasmatocytes of the peripheral hemocyte population
can transdifferentiate into lamellocytes. Rizki regarded lamellocytes as a specialized
type of plasmatocyte that develops in circulation by flattening via a transitional cell
type called podocyte [30], [79]. After wasp infection, circulating cell counts increase,
and it was therefore assumed that sessile hemocytes detach and develop into
lamellocytes [70]. In contrast, Meister and coworkers provided evidence that
lamellocytes develop in the lymph glands from prohemocytes and are released into
circulation, while not entirely excluding that lamellocytes can develop in the
periphery from prohemocytes [31]. The hypothesis in favor of lamellocyte
development in the lymph glands is corroborated by the observation that the primary
lymph gland lobes of some parasitized larvae disperse [80] and by the complete
absence of lamellocytes in a knot (collier) mutant where knot is acting in the PSC to
control lamellocyte differentiation [81].
These hypotheses were tested, and further investigations established that after
parasitization plasmatocytes and presumably prohemocytes, termed “double
negatives” for not expressing NimC1 (plasmatocyte antigen) or L1 (lamellocyte
antigen), left the sessile compartment. Lamellocytes appeared in circulation while the
primary lymph gland lobes were still intact. In an infected larva, where the anterior
half that contains the lymph glands was physically separated by ligation from the
posterior half where most of the sessile hemocytes reside, lamellocytes developed
only in the posterior ends of the larva [63]. Three independent lineage tracing studies
confirmed that peripheral plasmatocytes indeed differentiate into lamellocytes. Ando
and coworkers used two different drivers that are active in either the periphery (crqGAL4) or the lymph glands (Dot-GAL4). In non-infected larvae, the crq-lineage
derived plasmatocytes were only found in circulation, whereas Dot-lineage cells
resided only in the lymph glands. After parasitzation, lamellocytes in circulation
originated from both the lymph glands and the peripheral hemocyte compartments
[44]. Badenhorst and coworkers lineage traced the origin of lamellocytes after wasp
infection to plasmatocytes [46]. Avet-Rochex et al. knocked down ush in embryonic
plasmatocytes and were also able to show that plasmatocytes gave rise to lamellocytes
[45].
The presence of melanotic nodules in the absence of wasp infection in Drosophila
larvae has long since been associated with lamellocytes [82]. Mutations in genes of
two well-known pathways involved in immunity, the Toll pathway and the Jak/Stat
pathway, induce the formation of melanotic nodules [83], [84]. Two genetic screens,
a misexpression screen [85] and an in vivo RNAi screen [45] aimed to find new genes
involved in lamellocyte hematopoiesis. Lamellocyte hematopoiesis can be induced
genetically by overactivation of genes such as in the Toll10b (Tl10b) mutation [83],
[70], or by loss-of-function mutations in genes that repress lamellocyte
hematopoiesis such such as ush [76], [45]. Moreover, lamellocyte formation is also
controlled by epigenetic factors [85], [57]. It is noteworthy that melanotic nodules are
not always associated with blood cells [86].
Adult hematopoiesis
Adult hemocytes consist of plasmatocytes and crystal cells. They are able to
phagocytose bacteria, but their cell number declines with age [87], [88]. It has been
generally accepted that the adult fly does not have a hematopoietic organ. It was
assumed that embryonic [62] and larval hemocytes are passed on and constitute the
adult hemocyte population where they stop proliferating (reviewed in [75]).
However, it has recently been shown that four hematopoietic clusters exist in the
abdomen of the fly adjacent to the heart. The hemocytes are embedded within a
6
network of extracellular matrix where they develop from precursors to either
plasmatocytes or crystal cells. Lineage tracing revealed that hemocytes of the
secondary lymph gland lobes give rise to the precursors in the hematopoietic clusters.
When the fly is challenged by bacterial infections, hemocytes divide in the
hematopoietic hub. The authors stress the similarity with human bone marrow and
suggest using the adult fly´s hematopoietic hub as a model system for human
hematopoiesis [59].
Immune response to parasitoid wasps
1.
1. wasps inject eggs
+ venom
3.
2.
5.
4.
2. layer of on unknown compound is deposited on the
egg
3. plasmatocytes are recruited
and spread on the egg
4. lamellocytes are recruited
to the egg and encapsulation ensues
unknown
compound
plasmatocytes
lamellocytes
melanin
deposit
5. melanin is deposited on
the egg to seal it off
Figure 3 The melanotic encapsulation response against wasp eggs.
Long since have scientist been fascinated by the interaction of parasites with their
hosts. Like many other insects, Drosophila is frequently infected by parasitoid wasps.
Wasps from at least five genera parasitize Drosophilids. The figitids Leptopilina spp.
and Ganaspis spp. as well as the braconids Asobara spp are cosmopolitan larval
parasitoids, while Trichopria spp. (Diapriidae) and Pachycrepoideus vindemmiae
(Pteromalidae) attack the pupal stages of fruit flies [89], [90]. In particular
representatives of the genera Leptopilina, Ganaspis, and Asobara have been studied
in the context of immunology.
Melanotic encapsulation is evolutionary conserved in insects and is the visible
evidence of the vigorous immune response that the host mounts in response to
parasite infection. It is a stepwise process that comprises recognition of non-self
surfaces, recruitment of blood cells, transcriptional, translational, and biochemical
changes in blood cells and fat bodies, as well as alterations in the metabolism and
physiology of host larvae (reviewed in [91], [25], and Figure 3). The female wasp
pierces the cuticle of the host with her ovipositor and injects an egg along with venom
into the hemocoel of the Drosophila larva. The venom often contains virus or viruslike particles and additional compounds that attempt to attenuate the immune
response of the host [92]. In the larva, the parasitoid egg is recognized by an
unknown mechanism. The first visible event in capsule formation is the deposition of
an electron-dense layer of unknown material onto the chorion of the wasp egg as
early as six hours after infection that is recognized by plasmatocytes [93]. This layer
might consist of extracellular matrix because plasmatocytes of larvae mutated in
LaminA were not able to adhere to wasp eggs [94]. Plasmatocytes are recruited from
the sessile compartment [70], [63] and the lymph glands [31], [44]. They adhere to
the electron dense layer, spread over the egg surface, and form septate junctions to
seal of the egg [93]. Plasmatocyte recruitment after infection by Ganaspis sp.1 is
dependent on a calcium burst that is controlled by a calcium channel encoded by
7
Rya-r44F (Ryanodine receptor 44F) [95]. Secretion of the cytokine edin (elevated
during infection) from the fat body activates plasmatocytes to leave the sessile
hemocyte compartment [96]. Spreading of plasmatocytes on the wasp egg and
formation of septate junctions between plasmatocytes is mediated by the RhoGTPase
Rac2 [97]. Twenty-four hours after infection, lamellocytes appear in circulation and
start to encapsulate the egg [93]. RhoGTPases, RhoGEFs, and integrins are needed
for proper lamellocyte function [98], [94] . Adhesion of lamellocytes to the
plasmatocyte-covered wasp egg is controlled by the cellular adhesion molecule
Neuroglian [99]. The RhoGEF Zizimin-related (Zir) mediates spreading of
lamellocytes via the RhoGTPases Rac2 and Cdc42 [97], [100]. Rac1 in combination
with the heat shock protein Hsp83 mediates the relocalization of the beta-Integrin
Mysopheroid (Mys) to the periphery of spreading lamellocytes [101]. In addition,
Rac1 signals via the Jun Kinase basket (bsk) to regulate the turnover of focal
adhesions [102]. N-glycosylation on membrane proteins of lamellocytes is needed to
consolidate the capsule [103]. In addition, Hemese (He), a membrane glycoprotein
that is predicted to be O-glycosylated, is a negative regulator of encapsulation [104].
Finally, melanization of the encapsulated egg ensues, and the egg is completely
entombed within several layers of lamellocytes and encrusted by melanin deposits
forty hours after infection [105],[93],[97]. Rac1 and Rac2 mutant hemocytes, as well
as RNAi of bsk in blood cells display defects in melanizing wasp eggs [97], [102].
Hemocytes are the key players in the encapsulation response. Many studies found
that high larval blood cell numbers increased the success in killing the parasite [29],
[106],[107],[108],[109]. Only one study that used 24 european field lines of D.
melanogaster was not able to find that resistance to A. tabida was correlated with a
high hemocyte load [110]. The importance of hemocytes for the encapsulation
reaction becomes evident as D. obscura as is not able to generate lamellocytes in
response to parasitization [111].
Drosophila as a research tool
Next to the commonly advertised advantages of ideal model organism like
comparatively low maintenance costs due to small size, simple diet, short life cycle,
and fast reproduction, the true power of the fruit fly lies in its long and successful
history in research and in a fly community eager to develop new tools and make new
discoveries. It started in 1910, when T. H. Morgan introduced Drosophila into his
laboratory to study heredity. Only within five years, Morgan and his students Alfred
Sturtevant, Calvin Bridges, and Hermann Muller developed the theory of heredity
(reviewed in [112]), [113]. During these five years, more discoveries were made that
greatly contributed to the understanding of the principles of genetics for which
Morgan was awarded the 1933 Nobel Prize in Physiology or Medicine. From a lab
worker´s perspective, the introduction of balancer chromosomes by Muller in 1918 is
one of the highlights (reviewed in [112]). Muller´s research on mutagenesis earned
him the Nobel Prize in Physiology or Medicine “for the discovery of the production of
mutations by means of X-ray irradiation" .
Christiane Nüsslein-Volhard and Eric Wieschaus systematically mutated the
Drosophila genome and screened for genes impacting early development of the fly
[114], which won them the Nobel Prize in 1995 together with Edward Lewis "for their
discoveries concerning the genetic control of early embryonic development".
Traditional mutagenesis only permitted forward genetics. Transgenesis by
transposable elements opened the door to reverse genetics. Allan Spradling and Gerry
Rubin used the P-element as a “gene ferry” to artificially introduce extraneous DNA
into the fly´s genome [115], [116], [115]. P-element-mediated transgenesis fueled the
further development of additional techniques for gene manipulation. P-elements and
piggyBacs are the most widely used transposons [117]. P-element or piggyBack
insertions are available for more than 65 % of Drosophila genes. These transposons
8
can be directly used for gene disruption or to introduce targeted expression systems
into the genome [117], [118], [119]. The FRT/FLP system adapted from yeast is based
on the recombinase Flippase (FLP) to act on the Flippase Recombination Target
(FRT). FRT/FLP is used to create deletions [120] and to induce mitotic homozygous
clones in otherwise heterozygous animals [121], [122]. The GAL4-UAS system for
targeted gene expression uses the yeast transcription factor GAL4 to bind to the
promoter sequence UAS (Upstream Activation Sequence). GAL4 expression is
regulated by tissue-specific promoters that control UAS-mediated gene expression
[123]. GAL80, a negative regulator of GAL4, permits the tempo-spatial control of
gene expression by the GAL4-UAS system [124], [125], [126]. MARCM (Mosaic
Analysis with a Repressible Cell Marker) synergistically combines FLP/FRT, GAL4UAS, and GAL80 to mark homozygous clones [127], [128]. The Phi-C31 integrase is
facilitates target-specific genome engineering [129]. Finally, the CRISPR/CAS
technology has further eased genome engineering and introduced gene editing [130],
[131].
Article III of this thesis is based on the development of a new generation of
deficiency kits and the development of in vivo RNAi technology. Deficiencies are
delations that span a few to some hundred genes. They can be maintained in
Drosophila over balancer chromosomes. Deficiencies are Null alleles of the genes
they cover. They have been used to map mutations and to carry out modifier screens
[132]. Through the years, the Bloomington stock center has assembled a core
collection of 270 deletion strains spanning 70 % of the genome. The disadvantages of
these deletions are their heterogenous genetic backgrounds due to their diverse
origins and their imprecise mapping to the genome sequence [133]. The DrosDel
[134], [135] and Exelixis consortia [136] used genome engineering based on targeted
recombination mediated by the FRT/FLP system carried in P element [120], [137]
and piggyBac vectors [138] to overcome these limitations. The new deficiency kits
were generated in isogenized genetic backgrounds and the breakpoints are precisely
mapped to genome sequence [133]. The DrosDel deficiencies cover 77 % and the
Exelixis collection 56 % of the genome. The Bloomington stock center created and
additional set of deficiencies based on the Exelixis collection. In combination, the
DrosDel, Exelixis, and the Bloomington stock center deficiencies cover 98.4 % of the
genome [139].
RNA interference (RNAi) by double-stranded RNA is an efficient method of gene
silencing [140]. It is widely used in many species for in vitro and in vivo RNAi. In
Drosophila, in vivo RNAi is delivered by UAS-hairpin constructs that provide the
siRNAs necessary for gene silencing. GAL4-directed expression of the UAS-hairpin
constructs allows tempo-spatial control of RNAi. Drawbacks of RNAi are false
positive and false negative results. Off-target effects that lead to false positives are
caused by siRNAs that recognize and destroy mRNA of genes with similar nucleotide
sequence, and weak GAL4 driver expression insufficiently knocks down genes what
may result in false negatives [141], [142]. Three independent RNAi libraries approach
the problem of off-target effects in different ways. Currently approximately 30000
UAS-hairpin constructs are available [141].
Genetic Screens in Drosophila with focus on hemocytes
The purpose of a genetic screen is to identify new genes and their functions in a
particular process. Screening strategies follow two approaches: forward and reverse
genetics. In forward genetic screens, the wild-type function of a gene is inferred from
its mutant phenotype. Mutations can be induced by radiation, chemical mutagens, or
genetic techniques [132]. The genotypes of mutations fall into two classes: loss-offunction and gain-of-function mutations. Loss-of-function mutations are either
amorphs with complete loss of gene function (also called Null mutations), or
hypomorphs with reduced gene function. Gain-of-function mutations are
9
hypermorphs with enhanced gene function, neomorphs with novel gene functions,
and antimorphs with gene function opposite to the original function (also called
dominant negative mutations).
Genetic screens in Drosophila have a successful tradition. In 1995 Christiane
Nüsslein-Volhard and Eric Wieschhaus received a Nobel Prize for “Genetic control of
early structural development”. They investigated mutations of the embryonic
development of cuticular structures and identified segment polarity, pair-rule, and
gap genes by aforward genetic screening strategy [114]. Several types of screens have
been developed during the years (reviewed in [132], [143]). I will describe screening
strategies that have been used to find unknown genes in Drosophila immunity and
hematopoiesis. The first forward genetic screen to discover immunity genes that
respond to bacterial infection was carried out by Wu et al. [144].
In reverse genetic screening, known mutations of a genotype can be used to
screen for alterations in a specific phenotype. In vivo RNAi is a tool predestined to
carry out reverse genetic screens. In blood cell research, in vivo RNAi was used to
identify genes that induce lamellocyte formation [45] or regulate lymph gland
hematopoiesis [145], [146], [147].
A particularly useful approach to decipher unknown genes in a signaling pathway
is a genetic modifier screen. The key element of a modifier screen is a sensitized
genetic background in the signaling pathway or developmental process under
investigation. In a sensitized genetic background, an allele of a gene is used that
partially disrupts gene function, either enhancing (hypermorph) or reducing
(hypomorph) it. Amorphic mutations in most genes are recessive therefore only one
gene copy is needed to secure gene function. Under this condition, the sensitized
genetic background becomes susceptible to the signaling activity of downstream
genes, which may cause phenotypically detectable changes. A gene rendering the
phenotype of a sensitized genetic background more severe is called enhancer, a gene
rendering it more wild-type-like is called suppressor. This approach was used to find
genes involved in lymph gland hematopoiesis. A Zfrp8-Null allele with a lymph gland
overgrowth phenotype was used as the sensitized genetic background and deficiencies
were used to screen for enhancers and suppressors [146].
A misexpression screen is based on the assumption that also ectopic activation of
genes can help to elucidate gene function. Transposons have been used to introduce
promoter or enhancer elements into the fly genome that induce gene expression at
the integration site. Pernille Rørth created the first misexpression library called EP
(Enhancer Promoter) [148], additional libraries were created later [118]. Pxn-GAL4
induced overexpression of such enhancer elements was used to screen for genes
involved in peripheral larval blood cell hematopoiesis and localization [85].
Methods for hemocyte research
Originally, hemocyte types have been characterized by morphology and
ultrastructure with the help of light and electron microscopy. Research on Drosophila
blood cells started in larvae, presumably that is the developmental stage where
hemocytes are most easily accessible. Plasmatocytes and crystal cells are the basic
hemocyte types in all developmental stages of Drosophila [59], [149]. They are round
cells with a diameter of 7 to 12 μm. Plasmatocytes have a smooth cytoplasm with
primary lysosomes, and crystal cells harbor crystalline inclusions in their cytoplasm.
In addition, smaller cells with a diameter of 4 to 8 μm are referred to as
prohemocytes. Lamellocytes are large flat cells with a diameter of up to 50 μm but
with a thickness of only 1 μm. Their cytoplasm contains only few organelles, and their
plasma membrane often has microridges [30], [32], [31].
Advancements in molecular biological and genetical techniques enabled the
systematic identification of molecular markers for hemocytes. Screening of enhancer
trap libraries for β-galactosidase expression in circulating and lymph gland
10
hemocytes in the embryo and larva identified some of the first genes expressed in
blood cells. However, hemocytes were often not the only β-galactosidase-expressing
tissue present in individual fly lines [150], [151]. Kimbrell and coworkers screened for
reporter expression in immune tissues and for reporter lines with increased
expression induced by bacterial infection. They found Dorothy (Dot), escargot (esg),
and Cytochrome b5 (Cyt-b5) to be expressed in lymph glands as well as Collagen 25C
(Cg25C) and viking (vkg) in hemocytes and fat body. Esg and Cyt-b5 lines had
melanotic tumors. Thor expression increased after bacterial challenge [150]. Meister
and coworkers looked for enhancer trap lines with activity predominantly in
lamellocytes. They discovered 21 lines with hemocyte expression patterns, and five of
these could be used as lamellocyte reporters. The lamellocyte-specific reporter lines
were mapped to misshapen (msn), cAMP-dependent protein kinase A (Pka-C1), Chip
(Chi), and CG9932. An addition, they found domino [151]. The insertion close to msn
has been used as a lamellocyte reporter.
Another approach was to generate monoclonal antibodies against hemocyte
antigens that were used to identify hemocyte-specific genes. PDGF- and VEGFrelated factor 2 (Pvf2) was discovered by purifying the antigen with the help of
affinity chromatography and protein sequencing [77]. The Ando and Hultmark labs
generated pan-hemocyte antibodies, and antibodies specific for plasmatocytes and
plasmatocyte subtypes, lamellocytes and lamellocyte subtypes, as well as crystal cells
[72]. They identified five genes by using these antibodies to screen cDNA expression
libraries [104]. Hemese (He) is expressed by all hemocyte types in the periphery and
in the lymph glands. It is predicted to be glycosylated and has sequence similarity
with Glycophorins on the cell membranes of erythrocytes. He is a negative regulator
of the encapsulation response to parasitoid wasps and of tumor severity of l(3)mbn
[104]. NimC1 is expressed on all plasmatocytes. It is a phagocytosis receptor and the
founding member of a cluster of ten structurally related nimrod-like genes on
chromosome arm 2L [152]. Atilla is expressed by all lamellocytes. Its expression is
upregulated twofold in Ras-induced overproliferating hemocytes [153]. Cheerio
(cher) codes for Filamin-240, which is involved in the reorganization of the actin
cytoskeleton during the flattening of lamellocytes [154]. The L4 antigene was
identified as a beta-PS integrin (Kurucz et al. unpublished in [75]). In the fly
community, the He/H2, NimC1/P1, and atilla/L1 antibodies have become the
defining tool for blood cells in general, as well as plasmatocytes and lamellocytes in
particular.
The characterization of molecular markers for hemocytes permitted the
construction of hemocyte-specific GAL4 driver lines. He-GAL4 was the first driver
expressed in all larval hemocyte types, but only in approximately 80 % of all
circulating hemocytes. In contrast to its antigen, it is not active in the lymph glands of
unchallenged larvae [70]. Hemolectin-GAL4 (Hml-GAL4) [69], its truncated version
HmlΔ-GAL4 [155], Peroxidasin-GAL4 (Pxn-GAL4)[156], eater-GAL4 [157], srpHemoGAL4 [158], and Croquemort-GAL4 (crq-GAL4)[159] are expressed in embryonic
plasmatocytes, in larval plasmatocytes and crystal cells in circulation and in the
lymph glands, whereas eater is not to to expressed in the embryo [160]. Cg-GAL4 is
expressed in plasmatocytes and the fat body [153]. Lz-GAL4 is specific for crystal cells
[161], [43]. He-GAL4, Hml-GAL4, Pxn-GAL4, and crq-GAL4 are also expressed in the
hematopoietic hub of the adult fly [59]. Hml has sequence homologies with the von
Willebrand factor and is involved in hemolymph clotting [69]. Pxn [162] and Cg25C
[40] are extracellular matrix molecules produced by hemocytes and the fat body. Crq
and eater are phagocytosis receptors of which Crq mediates the phagocytosis of
apoptotic bodies [163] and eater of bacteria [160]. Srp [164] and lz [161] are
transcription factors involved in hematopoiesis. Besides ectopic expression of
transgenes, these GAL4 drivers enable the targeted expression of fluorescent proteins
11
in hemocytes to study expression patterns of hemocyte compartments and to monitor
hemocyte behavior in vivo.
In addition to GAL4 drivers, hemocyte-specific enhancer-reporter transgenes
were generated that decouple hemocyte visualization from driver expression. These
constructs facilitate the investigation of signals from other immune tissues in
hematopoiesis and immunity. Fly lines are available for each blood cell type and
tagged with different fluorescent proteins. This makes it possible to visualize several
blood cell types in combination [157]. The eater [165] and Hml [64] enhancers were
used to construct plasmatocyte-specific reporter constructs. The MSNF9mo enhancer
within msn [166] and the Bcf6 enhancer upstream of PPO1 [167] direct expression
specifically to lamellocytes and crystal cells, respectively. An insertion of a Minos
element into the atilla gene can be used as a lamellocyte reporter [168]. Recently,
Ando and coworkers combined in vivo antibody staining with blood cell reporter
expression for live imaging of hemocytes in Drosophila larvae [169]. Many more tools
are available for the lymph glands and are summarized in a current review [170].
Circulating larval hemocytes have generally been counted with the help of
hemocytometers (for example [70]) which is tedious, time consuming, and errorprone. A flow cytometry-based approach for circulating and lymph gland blood cells
was introduced [171], but only one report so far used flow cytometry for quantifying
blood cells [43]. In the adult fly, FACS was applied to sort GFP-expressing blood cells
[172], [38]. Moreover, an antibody-based rosetting method to separate hemocyte
subpopulations was developed [173].
12
Aims of the thesis
The general aim of this thesis has been to find out how the immune system
controls and orchestrates the hematopoiesis of lamellocytes. Lamellocytes constitute
a unique cell type in the respect that they are not present in the blood cell repertoire
of healthy larvae. Instead, their development is induced “naturally” - by wasp
infection, or “artificially” - under aberrant genetic conditions that lead to melanotic
nodules also referred to as tumors. Anyhow, their appearance signals that the
individual is immune-compromised. I used genetics, imaging, and flow cytometry to
find out more about lamellocyte hematopoiesis.
In Article I we used the blood cell driver He-GAL4 in a directed overexpression
screen to investigate which signaling pathways are involved in the activation of the
cellular immune system. Specifically, we looked for changes in numbers of peripheral
plasmatocytes and crystal cells and for the induction of lamellocytes.
In Article II we aimed to develop methodology to automate hemocyte counting
and facilitate live imaging of hemocytes in Drosophila larvae. We focused on a
reporter construct-based approach that enabled counting of hemocytes and
distinguishing between hemocyte subtypes by flow cytometry and live imaging.
In Article III we developed a genetic modifier screen to identify new genes that
control the activation of the cellular immune system.
In Article IV we studied the mechanism of Toll-dependent activation of blood
cells.
13
Article I
“A directed screen for genes involved in Drosophila blood cell
activation.”
Preface
In parallel to their work on humoral immunity, the Hultmark lab was aiming to
create antibodies against hemocytes to study cellular immunity. Istvan Ando and Eva
Kurucz had already generated some monoclonal antibodies and joined the Hultmark
lab, where a larger set was generated and characterized. This collaboration led to the
discovery of Hemese [104]. Since the Hemese antibody stains all hemocyte types [72]
it was decided to generate a Hemese-GAL4 driver assuming it would be expressed in
all hemocytes.
I joined the lab at the end of 2002 when Calle Zettervall had already cloned HeGAL4 and had started to overexpress genes relevant for human cancers. My
contribution to Article I was setting up a method for differential hemocyte counting
and then counting larval hemocytes of the F1 generation of He-GAL4-induced ectopic
expression of the transgenes that induced visible immune phenotype. Michael
Williams counted sessile crystal cells.
Summary of main results
1. The Hemese-GAL4 driver can be used to ectopically express genes in blood
cells.
2. Ectopic expression of activated and wild-type constructs of different signaling
pathways induce fate changes of hemocytes and affect proliferation of plasmatocytes
and activation of lamellocytes.
Discussion
1. The He gene was identified by using monoclonal antibodies previously
generated against larval hemocyte to screen a cDNA expression library made from
l(3)mbn-1 larval blood cells [104]. The Hemese antibody is specific for all sessile and
circulating hemocyte types as well as for hemocytes residing in the lymph glands [72].
The He-GAL4 driver was the first pan-hemocyte driver generated. Its expression
starts during the second larval instar (IA unpublished observation). The He-GAL4
driver is active in all blood cell types, but only in 75-80 % of all circulating blood cells.
It is also expressed in sessile hemocytes. As circulating and sessile hemocytes seem to
be one interchangeable population [64], it is likely that the percentage of He-GAL4expressing cells is equal in both compartments. In contrast to the antibody, He-GAL4
is not expressed in lymph glands of non-immune challenged larvae. We used HeGAL4 to overexpress genes that have been identified in human cancers.
2. We have started to use the terminology proliferation for increase of
plasmatocytes and activation for the de novo generation of lamellocytes. Several
models for lamellocyte activation have been proposed (see introduction and
discussion of Article II). According to our model in Article II, two lineages exist in
peripheral hemocytes: the plasmatocyte and lamellocyte lineage. Plasmatocytes are
extraordinarily plastic. After wasp parasitization three types exists. Lamellocytes are
generated in a demand-adapted manner from what we assume are prohemocytes.
Prohemocytes that undergo avid proliferation develop via prelamellocytes to mature
lamellocytes. Increase in total hemocyte count during the beginning of the
encapsulation response is almost entirely due to the proliferation of the lamellocyte
lineage (see discussion of Article II). Expression of a wild-type form of Alk, an active
14
form of Pvr, and the Tl10b mutant induce similar phenotypes when monitored with
the double hemocyte reporter construct eaterGFP,msnCherry (IA unpublished
results). As the blood cell phenotypes are similar after parasitization and genetic
manipulation, the results obtained in Article III can be used to reinterpret the blood
cell counts in Article I. Forced expression of constructs that represent different
signaling pathways turn on either a proliferation program, an activation program, or
a combination of both.
When plasmatocytes strongly proliferate, normally lamellocyte activation does
not occur like in the case of Egfr, Ras85D. Plasmatocyte diameters after
overexpression of Egfr and Ras85D visibly decreased. Small plasmatocytes were also
observed when Cg-GAL4 was used to express Ras85D.V12 [153]. The small size of
these cells is compatible with a hypothesis that these cells are prohemocytes and
constitutive Egfr-Ras85D signaling maintains prohemocyte fate and is necessary for
prohemocyte proliferation. Also adult midgut progenitor cells require Egfr signaling
for proliferation [174].
If the immune response favors activation as after the expression of an activated
form of hemipterous (hep.CA), the Drosophila homolog of Jun kinase kinase
(JunKK), or hopscotch (hopTum), the Drosophila homolog of Jak, only the lamellocyte
lineage is activated. Jak/Stat signaling is activated during the encapsulation response
[175], [176], [177]. Negative regulation of the wingless pathway also mainly activates
lamellocytes without a strong proliferative response of plasmatocytes. This is in line
with the role of wingless signaling in stem cell maintenance in the lymph gland [178].
The expression of a gain-of-function construct of shaggy (sggS9E), the Drosophila
homolog of Gsk3, had a particularly interesting overgrowth phenotype of the
secondary lymph gland lobes.
Overexpession of Alk, Pvr, and Toll activates the lamellocyte lineage and induces
proliferation of plasmatocytes. However, in comparison to Egfr and Ras85D
expression, plasmatocytes are diverse. High numbers of activated plasmatocytes type
II (see Article III) are in circulation. These cells are very reactive. And it is therefore
possible that the high degree of larval and pupal lethality is due to the presence of this
cell type.
Notably, several signaling pathways induce similar phenotypes. This suggests that
signaling pathways either act in parallel or sequentially. Sequential activation can
occur within the same cell or tissue or between cells or tissues. Cytokines are needed
for tissue interaction, which I discuss under Article IV. As overexpression of the Junk
and the Jak/Stat signaling pathway components had similar phenotypes, we tested if
these two pathways interacted genetically. We found that the Jak/Stat pathway
signals downstream of the Junk pathway to activate lamellocytes, and that the
Jak/Stat pathway suppresses all the effects of activated Junk signaling such as
lamellocyte activation, loss of sessile cells, and melanotic nodules (Kronhamn et
al.unpublished manuscript). This is supported by a report of the Schulz lab [166].
Moreover, serpent (srp) activity promotes lamellocyte activation downstream of the
activated Jak/Stat pathway, whereas u-shaped (ush) suppresses lamellocyte fate [76].
Some signaling pathway interactions are known for the humoral immune response
[179], [180], [181].
15
Article II
“Infection-induced proliferation is a central hallmark of the
activation of the cellular immune response in Drosophila larvae.”
Preface and study design
The quantification of hemocytes for Article I in the traditional manner by
hemocytometer led to the insights that blood cell numbers are variable, thus many
individual larvae need to be counted to obtain reliable results. Moreover, cell
counting in that way is not an entirely objective method, because it depends to a great
extent on the skill and patience of the experimenter. To circumvent these obvious
flaws, in Article II, we combined fluorescent plasmatocyte- and lamellocyte-specfic
enhancer-reporter constructs and developed a flow cytometry-based approach for cell
counting that we verified by confocal microscopy and cell sorting. In addition, these
reporters facilitated live imaging of Drosophila larvae.
Fluorescent hemocyte type-specific enhancer-reporter constructs decouple GAL4UAS driven gene expression from the mere observation of hemocytes. We had started
to design our own enhancer-reporter constructs just prior to the Schulz lab´s report
of fluorescently tagged plasmatocyte, crystal cell, and lamellocyte constructs [157].
We abandoned our own approach and instead used their constructs. I meiotically
recombined plasmatocyte and lamellocyte reporters tagged with different fluorescent
proteins onto the same chromosome. I also generated fly lines of plasmatocytelamellocyte reporter constructs in combination with several independent tissuespecific drivers and fluorescent constructs reporting other cell functions.
In all experiments, we were careful to have only one copy of each enhancerreporter, UAS-, and GAL4 construct in the F1 generation to warrant identical genetic
background compositions. This was necessary because differing genetic backgrounds
add another layer of complication to the already variable blood cell counts and
influence the speed of the immune reaction. For example in Fig. 9 of Article II, the
appearance of eaterGFP-low plasmatocytes in the RNAi-mediated knockdown of edin
in the fat body was delayed by three to four hours in a homozygous enhancer-reporter
genetic background, compared to the heterozygous genetic background.
Lamellocyte activation is part of the immune response induced after
parasitization by parasitoid wasps. The study of the virulence factors wasp females
inject alongside the egg into fly larvae bring to light the importance of the
encapsulation reaction for fly survival, as virulence factors attempt to abrogate this
immune response to ensure the survival of wasp larvae (reviewed in [25]). Because
we wanted to study lamellocyte hematopoiesis with a combined approach of flow
cytometry and imaging, we deliberately chose three wasp species of the genus
Leptopilina that affect the cellular immune response in different ways. L. boulardi is
a specialist for D. melanogaster and kills approximately half of the infected
individuals. L. heterotoma is a generalist species that kills almost all D. melanogaster
[177] by lysing lamellocytes [182], [183]. D. melanogaster is not a host-species for L.
clavipes and hence kills L. clavipes [184]. We expected that the predispositions of
these wasp species would be reflected in the dynamics and the cell types participating
in the induced immune response in fly larvae. Many features of the encapsulation
response have been described (reviewed in [25]), however, there has never been a
study that takes a holistic view of the dynamics of blood cells at the different sites in
the larva. We therefore decided to investigate circulating and sessile blood cells, the
blood cell response on the wasp egg, and the stability of the lymph glands at different
time points in non-infected and infected Drosophila larvae. The time points for the
survey of sessile blood cells and for the blood cell response on the wasp egg were
determined by time points of the timeline of circulating cells that mark major
16
changes. I constructed all the genetic tools used in this project. Laura Vesala and I
conducted all experiments together.
Summary of main results
1. Flow cytometry with two different reporter constructs detects six different
blood cell types of two functionally different lineages and a negative cell population:
Plasmatocyte lineage: eaterGFP-high plasmatocytes, activated plasmatocytes type
I and type II
Lamellocyte lineage: eaterGFP-low plasmatocytes, prelamellocytes, and
lamellocytes
2. Wasp parasitization induced a demand-adapted hematopoiesis in circulating
hemocytes that was based nearly only on the proliferation of eaterGFP-low
plasmatocytes and prelamellocytes of the lamellocyte lineage.
3. The origin of eaterGFP-low plasmatocytes remains elusive.
4. The encapsulation response against all wasp species is identical. It seems to be
the virulence factors of the wasp species rather than the immune response of the fly
that determine the outcome of infection.
5. The concerted action of hemocytes of all compartments (circulating and sessile
hemocytes, hemocytes on the wasp egg) contributes to a successful immune reaction.
6. The depletion of the cytokine edin in the fat body reduced eaterGFP-low
plasmatocyte numbers.
Discussion
1. Flow cytometry has a long history in human blood cell research [185]. The
development of flow cytometers that can distinguish between 12 different
fluorophores plus forward and side scatter was instrumental for modern blood cell
research and diagnostics. More than 100 blood cell populations can be identified
from human blood [186]. In Drosophila, only one report attempted to characterize
different blood cell populations simultaneously [171]. In this report, the distinction
between different blood cell types relied to a great extent on immunohistochemical
methods. Considering the fact that individual larvae have a low volume of
hemolymph, immunohistochemical methods for blood cell type differentiation and
counting are not suitable for high-throughput analysis on the level of individual
larvae. We are the first laboratory to develop a flow cytometry technique based on two
fluorescent in vivo reporter constructs [166], [165]. This enabled the detection and
counting of six different blood cell populations. In addition, we used flow cytometry
for detection and counting of proliferating cell types with an experimental set-up of
three fluorescent proteins.
Key to using flow cytometry is not only the development of adequate genetic and
other tools to investigate cell functions, but also the appropriate flow cytometer. We
were lucky to have an AccuriC6 flow cytometer at the institute. The advantage of the
AccuriC6 is the low volume of fluid it requires. We routinely prepared 100 µl of a
blood cell suspension of which we analysed 30 µl. This small amount of fluid was
enough to obtain reliable results.
Flow cytometry-based counting of distinct hemocyte populations can still be
improved. The AccuriC6 is also available with three lasers, which would allow the
detection and separation of additional fluorescent reporters. It would be desirable to
better characterize the hemocyte subtypes and create additional fluorescent in vivo
enhancer constructs.
2. In accordance with our findings, I suggest to distinguish between steady-state
hematopoiesis in healthy and demand-adapted hematopoiesis in immune-challenged
animals because these processes need to meet different requirements and are
functionally different. Steady-state hematopoiesis provides the animal with blood
17
cells for the daily wear-and-tear, whereas under circumstances of demand, new
effector cells have to be created and cells need to proliferate quickly (reviewed in
[187]). Immune responses require extra energy [188]. In addition, effector cells are
potentially harmful to the animal itself. Thus restricting the hematopoiesis of effector
cells such as lamellocytes contributes to immune and energy homeostasis.
In this study, we show that plasmatocytes develop by steady-state hematopoiesis
regardless of the state of the immune system, and lamellocytes by demand-adapted
hematopoiesis. However, after wasp infection eaterGFP-high plasmatocytes displayed
an extraordinary plasticity. In unchallenged larvae, eaterGFP-high plasmatocytes
were the predominant plasmatocyte type, thus the plasmatocyte population appeared
very homogenous. Plasmatocyte counts increased steadily during the time course of
our experiment and at the end of the time line, a sudden increase presumably due to
changes in the hormonal status just before metamorphosis occurred [189]. We were
not able to follow crystal cell counts with our experimental set up. After infect by
parasitoid wasps, plasmatocytes display a remarkable plasticity in circulation and on
the wasp egg. Activated plasmatocytes type II developed in contact with the wasp egg
and activated plasmatocytes type I in circulation and within the sessile islets.
Activated plasmatocytes type II retain eaterGFP and NimC1 expression while also
requiring lamellocytes markers. They seem to represent a new cell type that unifies
plasmatocyte and lamellocyte features.
Lamellocytes are immune-effector cells that only develop under certain
conditions. The mode and place of their hematopoiesis remains controversial. Rizki
described lamellocytes as morphological variants of circulating plasmatocytes that
emerge at the end of the third larval instar [30]. This view was corroborated by three
independent lineage tracing experiments that show that lamellocytes develop from
plasmatocytes [45], [44], [46] and one publication reporting that lamellocytes
develop from sessile plasmatocytes [63]. In contrast, other reports claim that the
lymph glands are the only source of lamellocytes. In the lymph glands, lamellocytes
develop from prohemocytes controlled by the actions of cells in the posterior
signaling center. The lymph glands disintegrate and the lamellocyts are released into
the hemolymph [81]. Yet another publication assumes that lamellocytes can develop
from prohemocytes also in circulation, but they are mainly released from the lymph
glands [31].
In our study, we show that lamellocytes develop in a demand-adapted fashion.
Early after infection eaterGFP-low plasmatocytes appear in circulation and divide
avidly. They develop further into prelamellocytes that give rise to mature
lamellocytes. Because of their sudden appearance, the low and transient expression of
eaterGFP, and the high proliferation rate, we assume that these cells are
prohemocytes or derive from prohemocyte.
3. The appearance of eaterGFP-low plasmatocytes is our most interesting
discovery. We tried hard to unravel their origin, but were not able to solve that quest.
As described before, two experimentally based hypotheses of the origin of
lamellocytes have been discussed in literature. Therefore we considered several
possibilities for the origin of eaterGFP-low plasmatocytes Firstly, eaterGFP-high
plasmatocytes give rise to eaterGFP-low plasmatocytes by asymmetric cell division,
by dilution of the eaterGFP content due to an accelerated cell cycle accompanied by a
strong increase in proliferation rate, or by simply down regulating their eaterGFP
while directly transdifferentiating into eaterGFP-low plasmatocytes. We tested the
first option by imaging blood cells of infected larvae early during the immune
response in order to find cells with high and low eaterGFP expression that had just
undergone mitosis. We saw many dividing cells, but none met the requirement of
asymmetric cell division. Next, we considered the dilution effect of eaterGFP after cell
division. The fluorescent signal emitted by eaterGFP-high plasmatocytes is
approximately ten times higher than that of eaterGFP-low plasmatocytes. This
18
implies that eaterGFP-high plasmatocytes have to go through three to four rounds of
cell division within only eight hours (the time of the first appearance of eaterGFP-low
plasmatocytes until their highest peak). This also implies that eaterGFP-high
plasmatocytes must quadruplicate their division rate to keep their own numbers
constant while serving as a pool for eaterGFP-low plasmatocytes. Alternatively, the
counts of eaterGFP-high plasmatocytes should drop drastically in favor of eaterGFPlow plasmatocytes during the early demand-adapted phase of the immune reaction.
None of the listed options is in agreement with our experimental data. Finally, the
half-life of enhanced GFP is approximately 24 hours [190], [191]. Down regulating
enhanced GFP in only eight hours is impossible.
Secondly, eaterGFP-low plasmatocytes are prohemocytes or derive from
prohemocytes only transiently expressing low levels of eaterGFP during their
maturation step to become lamellocytes. The most compelling support for this
hypothesis is that plasmatocyte and lamellocyte lineages are strictly separated on our
FACS plots. Moreover, several publications report the presence of prohemocytes [31],
[192], [193] in circulation.
4. The interaction of a parasite with its host has always been described as an
evolutionary arms race. Some of the virulence mechanisms of the wasp parasitoids
have been described and the encapsulation response has been studied (reviewed in
[25]). When comparing the differential hemograms of circulating hemocytes of
animals infected by the three different wasp species, it becomes evident that the
encapsulation response is orchestrated in a similar manner with slight delays in the
timing. It rather seems that the virulence factors of the wasps shape the immune
response. In the lamellocyte lineage after a lapse phase with no apparent action,
eaterGFP-low plasmatocytes, prelamellocytes, and lamellocytes appear stepwise in
circulation. After all these cell types have reached their peaks, their numbers drop
and even out at a lower level. The timing of eaterGFP-low plasmatocyte emergence in
the hemolymph and the appearance of the first blood cells on the wasp eggs of L.
boulardi coincide. Blood cells do not adhere to eggs of L. clavipes and L. heterotoma.
As aforementioned, L. heterotoma kills as good as all infected Drosophila larvae. L.
heterotoma females inject lamellolysin alongside the egg into the host. Lamellolysin
is a compound that affects lamellocyte morphology and finally kills them [194]. In our
experiment, lamellocyte precursors, the eaterGFP-low cells, appear but the more
mature lamellocyte types are depleted after infection with L. heterotoma. L. boulardi
injects three venoms with know function into the host larva: A RhoGAP [195], [196]
which affects hemocyte spreading behavior, a serpin [197], and a superoxide
dismutase [198] both of which affect the melanization reaction. Larvae infected by L.
boulardi have a clear delay in melanization compared to infection by L. clavipes.
After L. clavipes infection, melanization of the wasp egg starts as soon as lamellocytes
appear.
5. Simultaneously with the first eaterGFP-low plasmatocytes in circulation,
eaterGFP-high and activated plasmatocytes type II are found on the wasp egg. This
may suggest that recognition of the wasp egg by eaterGFP-high plasmatocytes
induces the demand adapted-hematopoiesis of lamellocytes. In the wound healing
response of the embryo, macrophages in the vicinity of the wound migrate towards
wound edges [156] while in the wound healing response of third instar larvae
hemocytes are recruited by direct capture from the hemolymph [20]. Pupal
macrophages are able to crawl [199]. In embryos, H2O2 is generated at sites of
wounding which induces a calcium flash in epithelial cells that directs macrophages
to the wound (reviewed in [200]). Recently, Evans et al. published that the ITAM
motif of Draper in combination with Src42A and Shark is required for recognition of
H2O2 and migration of macrophages to wounds [201]. Currently it is not known in
which manner plasmatocytes are recruited to the wasp egg. But while analyzing the
images of wasp-infected larvae at early time points, I often had the impression that in
19
the vicinity of the wasp egg sessile hemocytes had vanished. This may suggests that
plasmatocytes actively migrate towards the wasp egg from the surrounding sessile
compartments. Additional experiments are needed to clarify how plasmatocytes are
recruited to the wasp egg at the beginning of the encapsulation response.
With live imaging of cultured wasp eggs, we were able to show that eaterGFP-high
plasmatocytes differentiate into activated plasmatocytes type II on the wasp egg.
Thus this differentiation process is tightly linked to the wasp egg. In addition,
plasmatocytes type II are rarely seen in circulation and in the sessile compartment.
The signals inducing these changes are so far unknown. Thirty hours after infection,
the count of activated plasmatocytes type I rises over that of eaterGFP-high
plasmatocytes. We are not certain, if these type of activated plasmatocytes
phagocytizes msnCherry-containing material or if these cells truly express
msnCherry. Their increase coincides with the hatching of the L. boulardi wasp larvae.
Lamellocytes develop by demand-adapted hematopoiesis from prohemocytes.
How the lamellocytes are recruited to the wasp egg is not known. However, we are
able to show that blocking the secretion of the small peptide edin from the fat body
reduces eaterGFP-low plasmatocytes. Therefore, the fat body has to receive a signal
that instructs the secretion of edin. The nature of that signal is unknown, however,
possibly dependent on Toll signaling in the fat body (see discussion in Article IV and
[202], [203]). The initial signal how the fat body is alerted to the presence of the wasp
egg is unknown.
Also the lymph glands are alerted to the presence of the wasp egg. After infection
of Drosophila larvae by L. boulardi and L. clavipes, the primary lymph gland lobes
disintegrate. How the lymph glands sense the presence of a wasp egg is not known. A
study by Sinenko et al. suggests that circulating plasmatocytes detect the wasp egg
and signal to the lymph glands. In the medullary zone, the inhibition of Jak/Stat
signaling in prohemocytes is released and ROS levels increase in the posterior
signaling center. This has two effects: On the one hand, together with the inhibition
of Jak/Stat signaling, ROS induces the differentiation of prohemocytes to
lamellocytes in the medullary zone of the lymph gland. On the other hand, increased
ROS levels induce the secretion of spitz, a ligand of Egfr, from the posterior signaling
center into the hemocoel. Spitz then controls the transdifferen-tiation of
plasmatocytes into lamellocytes [192], [204].
It is clear that a concerted action of all blood cell compartments, including those
blood cells on the wasp egg, contribute to the encapsulation response. The origin and
the nature of many of the signals are however still elusive. The development of new
genetic tools is needed to unravel the interaction of different tissues.
20
Article III
“Genetic screen in Drosophila larvae links ird1 function to Toll
signaling in the fat body and hemocyte motility.”
Preface
As Article I of this thesis shows, several genes of distinct signaling pathways
induce lamellocyte formation. In comparison to crystal cells and plasmatocytes that
are present at all developmental stages, lamellocytes are only activated in
immunocompromised animals in a demand-dependent manner. Lamellocytes are
immune effector cells whose hematopoiesis is likely controlled differently from that
of plasmatocytes and crystal cells. We have been interested in lamellocytes because of
their role in encapsulation and melanotic nodule formation. A genetic screen seemed
a reasonable approach to discover genes involved in lamellocyte hematopoiesis. My
contribution to this project was designing the screen, developing the screening
strategy, and constructing genetic tools. Martin Schmid and Jesper Kronhamn
contributed to laying out some parts of the screening strategy. And Martin conducted
the screen and did most of the work on ird1. I will therefore focus my discussion on
the design of the screen and the screening strategy.
Summary of main results
1. The Toll10b gain-of-function mutation can be used as a sensitized background
for a genetic modifier screen.
2. Ird1 rescues the pattern of sessile bands and the loss of the primary lymph
gland lobes, but enhances all other Toll10b phenotypes.
Discussion
1. Genetic modifier screens have been successfully used to unravel genetic
interactions and to discover new genes within signaling pathways (reviewed in [132]).
The key to success is a sensitized genetic background that can be modified by
mutations in genes that genetically interact with the gene providing the sensitized
genetic background. In the work described in Article I, we show that the directed
expression of a number of genes induced lamellocytes. Lamellocyte activation is
usually accompanied by a change in the pattern of sessile hemocytes and melanotic
nodule formation. Thus our criterion was that any gene whose overexpression
induced melanotic nodule formation might provide a suitable sensitized genetic
background for screening. I initially designed genetic tools for three sensitized genetic
backgrounds based on UAS-Alk, UAS-Pvr.λ, and the Toll10b (Tl10b) mutation [205],
[206]. All three genes have in common that they are receptors. Their overactivation
induces gain-of-function genotypes, which are phenotypically characterized by the
presence of lamellocytes, melanotic nodules, changes in the orderliness of the pattern
of sessile hemocytes, and a mild to strong increase in plasmatocyte counts.
Hierarchically, a receptor is the first gene in a signaling cascade within a cell, and
mutations that decrease the functions of genes signaling downstream of the receptor
are theoretically able to enhance or suppress the increased signaling activity of a
genetically activated receptor. This is advantageous for screening because the yield of
modifiers should be high. However, the outcome of the screen depends on the nature
of the gain-of-function allele used (for discussion see [132]).
Our screening strategy was to conduct an F1 deficiency screen with DrosDel
deficiencies [134], [135], map the loci to single gene level with in vivo RNAi lines from
21
the VDRC (Vienna Drosophila Resource Center) [207], and confirm the newly
discovered genes with mutations.
Experiments we conducted earlier pointed out the importance of keeping the
genetic background constant. This is due to the very nature of blood cells. Their
numbers vary during larval development and between individuals of the same age
and the same fly line. Because of this variability, a high number of individuals need to
be screened to obtain interpretable results. To minimize the workload and make
results more reliable that is to reduce the amount of false positives, keeping the
genetic background constant and controlled is of paramount concern. The DrosDel
deficiency kit was created in an isogenized genetic background, which tremendously
helped in reducing the risks of false positives.
The advantage of RNAi lines over mutations is that there exists at least one RNAi
line for each gene in the fly. Moreover, the original GD lines of the VDRC RNAi
library were generated in the genetic background of the DrosDel deficiency kit. In
contrast, mutations come with diverse genetic backgrounds. The disadvantage of
using RNAi is that deficiencies are functional null mutations and thus affect every
tissue. Gene knockdown by RNAi is dependent on the GAL4-UAS system and
therefore only affect those tissues the driver is active in. This implicates that
whenever using RNAi lines to map loci determined on the basis of deficiencies, genes
will be false positives, if their effects on the sensitized genetic background is
dependent on a tissue the GAL4 driver is not expressed in.
Originally, I had planned to screen for the modification of two phenotypes: the
pattern of sessile cells in larvae and the melanotic nodule phenotype in larvae and
imagines. In fact, screening for modification of melanotic nodules in imagines was my
preferred strategy. To be able to visualize sessile blood cells in larvae and to be able to
express RNAi lines in blood cells, I crossed HmlΔ-GAL4, UAS-GFP (HmlΔ>GFP) [155]
into the fly line containing the Tl10b mutation. This generated
w;HmlΔ>GFP;Tl10b/T1;3(OR60)/TM6,Tb. The Tl10b mutation is dominant female
sterile [205]. Therefore, the Tl10b mutation is kept only in males to preserve the
mutation and propagate it from generation to generation. For this reason, the Tl10b
mutant fly line essentially consists of two individual fly stocks: w/Y; Tl10b/TM6,Tb
(males) and w; T1;3(OR60)/TM6,Tb (females). TM6 is the balancer chromosome
(Third Multiple 6) the Toll mutation is kept over. T1;3(OR60) is a translocation
between chromosome one and three, which causes lethality in males. Crossing
together these males and females produces only one viable type of male: w/Y;
Tl10b/TM6,Tb and three viable types of females of which only one is fertile: w;
T1;3(OR60)/TM6,Tb. In order to cross any construct into the Toll10b stock, it has to
be done separately for each sex. HmlΔ-GAL4 (HmlΔ>GFP;Tl10b) drives GFP
expression only in plasmatocytes without ectopic expression in salivary glands or the
gut that hampered the use of the He driver for this screen. However, the viability of
this newly created HmlΔ>GFP;Tl10b was lower than the original Tl10b stock. It was
especially low when crossed to DrosDel deletions. I screened DrosDel deletions for
the entire X-chromsome for enhancers or suppressors of nodule formation in the
adult fly. It turned out that pupal lethality was too high. I was never able to recover
more than ten or twenty flies per cross. This number of flies was insufficient to base a
reliable screen on. Thus we abandoned this approach and turned to larvae, as larval
viability was not as severely affected.
DrosDel deficiencies are kept over balancer chromosomes with genetic markers
that affect only structures visible in flies. Therefore, I crossed green balancer
chromosomes (balancer chromosomes that express GFP also visible in larvae) of all
chromosomes into the isogenized genetic background of the DrosDel kit. The GFP
expression pattern of these green balancers differs from HmlΔ>GFP. All deficiencies
were rebalanced over green balancers and used to distinguish deficiency
chromosomes from balancer chromosomes in the F1 generation. In addition, most of
22
the fly stocks used in this project whose genetic backgrounds differed from that of
DroDel were backcrossed six times into the genetic background of the DrosDel
deficiency kit.
Martin tested that mutations of MyD88, an adaptor protein within the Toll
signaling cascade, suppressed the disruption of sessile cells, which verified our
approach. Martin, Jesper, and I developed the hemocyte mobilization index based on
a publication by the Li lab [208]. Because chromosome 3R had a high coverage rate
with a relatively low number of individual deficiencies, we decided to screen this
chromosome arm.
In the end, Martin had screened 88 deficiencies, uncovered six suppressor
regions, and discovered ird1. Ironically, mutations of ird1 also have an activated
immune phenotype. However, the pattern of sessile cells and he loss of the primary
lymph gland lobe is rescued.
23
Article IV
“Control of Drosophila blood cell activation via Toll signaling in
the fat body.”
Preface
In addition to designing and contributing to the Toll screen, I conducted a
modifier screen of my own specifically tailored to discover novel genes involved in the
activation of lamellocytes. Notably in this screen, deficiencies that uncover genes in
the Toll signaling pathway were overrepresented. According to my strategy to use in
vivo RNAi to map these loci to the single gene level, I knocked down genes of the Toll
signaling cassette in hemocytes. However, RNAi-dependent depletion of Toll
signaling in hemocytes did not suppress the phenotypes I was screening for, whereas
mutants of the Toll signaling pathway were suppressors. Additionally, hemocytespecific expression of the RTKs Alk and Pvr, as well as other genes that induce
lamellocyte activation, induced the activation of Toll signaling in the fat body
measured by Drosomycin-GFP activity. Taken together, these findings suggested that
there must be a signal generated in hemocytes that activates Toll signaling in the fat
body, which would induce the expression of Toll-dependent genes that in turn would
control hemocyte behavior. At the same time, Martin discovered in a joint project
with the Rämet group that hemocyte-specific knock-down of MyD88 was not
sufficient to rescue the melanotic nodule phenotype nor the aberrant pattern of
sessile cells of the Tl10b mutation. Thus we joined forces to get to the bottom of the
tissue specificity of Toll signaling.
Summary of main results
1. It is sufficient to express an activated form of Toll in hemocytes, but Toll
signaling is sufficient and necessary in the fat body to direct hemocyte behavior. All
aberrant phenotypes of the Toll10b mutant can be suppressed by knocking down Toll
signaling in the fat body.
2. The immune response to L. boulardi is correlated with Toll activation in the
fatbody.
Discussion
1. The fact that we were able to suppress every phenotype associated with the
gain-of-function mutation Tl10b (melanotic nodules, lamellocyte activation, loss of the
pattern of sessile cells, and loss of the primary lymph gland lobes) when we
suppressed Toll signaling in the fat body suggests that the fat body secretes inducible
Toll-dependent gene products into the hemolymph that control hemocyte behavior.
In human immunology, cytokines, small molecules that shape the behavior of blood
cells, have a long history [209], [210]. In humans, Toll signaling activity induces the
production of inflammatory cytokines and interferons in many tissues (reviewed in
[211]). Therefore, it seems likely that some of the Toll-dependent genes secreted by
the fat body of Drosophila might be cytokines. However, very little is known about
cytokines secreted by the fat body that affect hemocyte behavior. Attractive
candidates for Toll-dependent cytokines are antimicrobial peptides. But until now to
the best of my knowledge, no one has shown that antimicrobial peptides instruct the
behavior of hemocytes. We recently published that the infection-inducible peptide
edin (elevated during infection) is secreted from fat body and is required for the
release of sessile hemocytes after wasp infection [96]. It is not known so far by which
mechanism edin instructs hemocyte release. Edin was discovered in an S2 cell screen
24
where it was inducible in a Relish-depended manner in response to heat killed E. coli
[212]. Moreover, edin was induced in wntD mutants during L. monocytogenes
infection [203]. Notably, wntD is a negative feedback inhibitor under the control of
Dorsal/Toll signaling [202]. This suggests that edin expression might be induced by
Toll signaling in the fat body. The Drosophila TNF homolog eiger (egr) is expressed
in the fat body after infection with S. typhimurium and mediates melanization,
antimicrobial peptide secretion, and survival of the Drosophila host [213].
Toll10b is a gain-of-function mutant of Toll that does not require ligand activation
[206]. But under normal conditions Toll needs a ligand for its activation. Spätzle
(spz) is the only known ligand for the Toll receptor in immunity (reviewed in [214]).
Spz is secreted by hemocytes into the hemolymph where it is proteolytically cleaved
and then activates Toll signaling in the fat body [36]. More interactions are known
where hemocytes send signals to the fat body, however, not all signals induce Toll
signaling. The lysosomal protein psidin (phagocyte signaling impaired) is needed for
phagosome maturation in plasmatocytes, but also sends an unknown cytokine to the
fat body to produce Defensin in response to bacterial infection [215]. Synergistically,
nitric oxide and CanA1 (Calcineurin A1) trigger Relish signaling in the fat body in
response to feeding of gram-negative bacteria [216]. Septic injury induces the
expression of upd3 in hemocytes. Upd3 is secreted from hemocytes and triggers
Jak/Stat signaling in the fat body [217].
More complex bidirectional signaling between hemocytes and fat body was
described in a tumor model of Drosophila epithelial cells. Polarity loss in tumors
triggers the secretion of Pvf1 which induces hemocyte proliferation via its receptor
Pvr. Hemocyte-dependent secretion of egr causes tumor cell death and hemocytedependent spz activates the Toll signaling pathway in the fat body resulting in a
reduction of tumor burden. The signal the fat body sends to control tumor growth is
not yet known [218]. Yet another example for more complex signaling is known for
Growth-blocking peptide (Gbp). Gbp is constitutively present in the hemolymph as
an inactive form that is activation by proteolysis. Upon various stresses, Gbp activates
Metchnikowin expression in an imd- and Jnk-signaling-dependent manner in the fat
body [219]. In S2 cells, in addition to activating Metchnikowin expression in an imdand Junk-dependent manner, Gbp stimulates a Ca2+ influx by PLC activation.
Elevated Ca2+ levels induce the expression and secretion of Pvf, which in turn
activates Erk-signaling within the same cell by binding to its receptor Pvr. Then, Erksignaling promotes cellular immunity while simultaneously inhibiting antimicrobial
gene expression [220].
In conclusion, these examples illustrate that cooperation and communication
between tissues are important and necessary in facilitating immune signaling. As
sentinels, hemocytes recognize dangers and alert the fat body and most likely other
tissues by sending signals in the form of cytokines. In response, the fat body and
other immune responsive tissues orchestrate the humoral and cellular immune
response. The Tl10b mutant might be of help in deciphering the instructive signals the
fat body secretes to control hemocyte behavior.
2. The analysis of mutants of the Toll pathway infected by wasps [175] and gene
profiling of D. melanogaster after wasp infection [176], [177], [221] illustrate the
central role of Toll signaling in the melanotic encapsulation response. The females of
some wasp species inject venoms into fly larvae that aim to negatively regulate Toll
signaling [197], [198], [222]. We monitored the activity of the Toll signaling pathway
with a Drs-GFP reporter [223]. Interestingly, the majority of larvae that had
successfully killed the parasite did not express Drs-GFP (Fig. 5 D in paper 4). The
analysis of Drs-GFP expression was done 48 h after infection. Possibly, the activity of
Toll signaling had abated by that time point. Alternatively, other signaling pathways
are able to fill the gap of attenuated Toll signaling. Surprisingly, knocking down Toll
signaling components in hemocytes lead to an enhanced encapsulation response early
25
during infection. This result is puzzling because the L. boulardi venoms are aiming to
block Toll activity. An explanation might be that Toll signaling in hemocytes has a
dampening effect to balance out the immune response.
26
Acknowledgements
Thank you!
Tack!
Kiitos!
Danke!
A long time has gone by and a huge project has finally come to its end. Many
people have contributed to my work, and I owe a big ”thank you” to many people.
First, I would like to thank you, Dan, for the opportunity of being able to work in
your labs in Umeå and Tampere. And the patience you had with me and my many
projects during the years. Soon you will also my ”Doktorvater”, not only Martin´s.
Then, I would like to thank my committee, Åsa and Ruth, for some valuable
advice during my PhD thesis.
I would like to thank all the present (Sajna and Hairu) and past members (JensOla, Jesper, Karin, Svenja, Ingrid, Mazen Michael, MagdaLena, Pia, Calle,
AnnaKarin, Karin Ekström, and Anni) of the Hultmark lab in Umeå and the danlab in
Tampere. Laura, I am so glad you joined the lab in Tampere. Together we have not
only compiled an insane amount of data, but have also done some insanely good
research. I hope we will be able to work together in the future. Martin, thank you for
the collaboration on our Toll projects. All the best for your fly-bee project. I would
also like to say thank you to Tiina and Tuula for the great help you have been in the
lab.
To all the people at Molecular Biology in Umeå that I still know. It is always nice
coming back and feeling welcome.
During my time in Tampere, I had the pleasure of supervising a number of very
talented students. Thank you, Otto, Siina, Sandra, Johannes, Volker, Ali, and Eveliina
for your input into the projects.
I would like to say thank you to Mika and the people in his lab, especially Kaisa,
Leena-Maija, Susanna, Henna, and Mataleena, for the hospitality I received, when I
first came to Tampere. Leena-Maija and Susanna, working with you was nice, and we
managed to get some good stuff together. Henna, not so much lab work togther, but
the more fun. It was great, spending some long nights together with ”skratanfall”.
Eric, I am so glad you never gave up in trying to convince us to using flow
cytometry. Alberto, thank you for all the science and carreer related discussions.
Bettina, Ana, Tiina, and Päivi, thank you for being good friends here in Tampere.
Anna and Dirk, Esther and Arne, Örjan and Annika, andZlatko thank you for all the
time we spent together and the fun we had outside of work.
Finally, I would like to thank my family for their love and support.
27
References
1. Kaufmann SH (2008) Immunology's foundation: the 100-year anniversary
of the Nobel Prize to Paul Ehrlich and Elie Metchnikoff. Nat
Immunol 9: 705-712.
2. Janeway CA, Jr. (1989) Approaching the asymptote? Evolution and
revolution in immunology. Cold Spring Harb Symp Quant Biol 54 Pt
1: 1-13.
3. Medzhitov R (2009) Approaching the asymptote: 20 years later. Immunity
30: 766-775.
4. Doulatov S, Notta F, Laurenti E, Dick JE (2012) Hematopoiesis: a human
perspective. Cell Stem Cell 10: 120-136.
5. Wynn TA, Chawla A, Pollard JW (2013) Macrophage biology in
development, homeostasis and disease. Nature 496: 445-455.
6. Hultmark D (2003) Drosophila immunity: paths and patterns. Curr Opin
Immunol 15: 12-19.
7. Thaiss CA, Levy M, Suez J, Elinav E (2014) The interplay between the
innate immune system and the microbiota. Curr Opin Immunol 26:
41-48.
8. Ryu JH, Kim SH, Lee HY, Bai JY, Nam YD, et al. (2008) Innate immune
homeostasis by the homeobox gene caudal and commensal-gut
mutualism in Drosophila. Science 319: 777-782.
9.
(http://www.globalchange.umich.edu/globalchange2/current/lectures/biod
iversity/biodiversity.html).
10. Travis J (2009) Origins. On the origin of the immune system. Science
324: 580-582.
11. Imler JL (2014) Overview of Drosophila immunity: a historical
perspective. Dev Comp Immunol 42: 3-15.
12. Boman HG, Hultmark D (1987) Cell-free immunity in insects. Annu Rev
Microbiol 41: 103-126.
13. Boman HG, Nilsson I, Rasmuson B (1972) Inducible antibacterial defence
system in Drosophila. Nature 237: 232-235.
14. Sun SC, Faye I (1992) Cecropia immunoresponsive factor, an insect
immunoresponsive factor with DNA-binding properties similar to
nuclear-factor kappa B. Eur J Biochem 204: 885-892.
15. Engstrom Y, Kadalayil L, Sun SC, Samakovlis C, Hultmark D, et al. (1993)
kappa B-like motifs regulate the induction of immune genes in
Drosophila. J Mol Biol 232: 327-333.
16. Rosetto M, Engstrom Y, Baldari CT, Telford JL, Hultmark D (1995)
Signals from the IL-1 receptor homolog, Toll, can activate an
immune response in a Drosophila hemocyte cell line. Biochem
Biophys Res Commun 209: 111-116.
17. Lemaitre B, Nicolas E, Michaut L, Reichhart JM, Hoffmann JA (1996)
The dorsoventral regulatory gene cassette spatzle/Toll/cactus
controls the potent antifungal response in Drosophila adults. Cell
86: 973-983.
28
18. Theopold U, Schmidt O, Soderhall K, Dushay MS (2004) Coagulation in
arthropods: defence, wound closure and healing. Trends Immunol
25: 289-294.
19. Theopold U, Krautz R, Dushay MS (2014) The Drosophila clotting system
and its messages for mammals. Dev Comp Immunol 42: 42-46.
20. Galko MJ, Krasnow MA (2004) Cellular and genetic analysis of wound
healing in Drosophila larvae. PLoS Biol 2: E239.
21. Stramer BM, Dionne MS (2014) Unraveling tissue repair immune
responses in flies. Semin Immunol 26: 310-314.
22. Eleftherianos I, Revenis C (2011) Role and importance of phenoloxidase
in insect hemostasis. J Innate Immun 3: 28-33.
23. Nappi AJ, Christensen BM (2005) Melanogenesis and associated
cytotoxic reactions: applications to insect innate immunity. Insect
Biochem Mol Biol 35: 443-459.
24. Carton Y, Nappi AJ (1997) Drosophila cellular immunity against
parasitoids. Parasitol Today 13: 218-227.
25. Keebaugh ES, Schlenke TA (2014) Insights from natural host-parasite
interactions: the Drosophila model. Dev Comp Immunol 42: 111-123.
26. Jiravanichpaisal P, Lee BL, Soderhall K (2006) Cell-mediated immunity
in arthropods: hematopoiesis, coagulation, melanization and
opsonization. Immunobiology 211: 213-236.
27. Evans CJ, Hartenstein V, Banerjee U (2003) Thicker than blood:
conserved mechanisms in Drosophila and vertebrate hematopoiesis.
Dev Cell 5: 673-690.
28. Wood W, Jacinto A (2007) Drosophila melanogaster embryonic
haemocytes: masters of multitasking. Nat Rev Mol Cell Biol 8: 542551.
29. Russo J, Brehelin M, Carton Y (2001) Haemocyte changes in resistant
and susceptible strains of D. melanogaster caused by virulent and
avirulent strains of the parasitic wasp Leptopilina boulardi. J Insect
Physiol 47: 167-172.
30. Rizki MTM (1957) Alterations in the haemocyte population of Drosophila
melanogaster. Journal of Morphology 100: 437-458.
31. Lanot R, Zachary D, Holder F, Meister M (2001) Postembryonic
hematopoiesis in Drosophila. Dev Biol 230: 243-257.
32. Shrestha RBG, E. (1982) Ultrastructure and Cytochemistry of the Cell
Types in the Larval Hematopoietic Organs and Hemolymph of
Drosophila Melanogaster. Development, Growth and Differentiation
24: 65-82.
33. Babcock DT, Brock AR, Fish GS, Wang Y, Perrin L, et al. (2008)
Circulating blood cells function as a surveillance system for damaged
tissue in Drosophila larvae. Proc Natl Acad Sci U S A 105: 1001710022.
34. Dimarcq JL, Imler JL, Lanot R, Ezekowitz RA, Hoffmann JA, et al. (1997)
Treatment of l(2)mbn Drosophila tumorous blood cells with the
steroid hormone ecdysone amplifies the inducibility of antimicrobial
peptide gene expression. Insect Biochem Mol Biol 27: 877-886.
29
35. Pastor-Pareja JC, Wu M, Xu T (2008) An innate immune response of
blood cells to tumors and tissue damage in Drosophila. Dis Model
Mech 1: 144-154; discussion 153.
36. Shia AK, Glittenberg M, Thompson G, Weber AN, Reichhart JM, et al.
(2009) Toll-dependent antimicrobial responses in Drosophila larval
fat body require Spatzle secreted by haemocytes. J Cell Sci 122:
4505-4515.
37. Cordero JB, Macagno JP, Stefanatos RK, Strathdee KE, Cagan RL, et al.
(2010) Oncogenic Ras diverts a host TNF tumor suppressor activity
into tumor promoter. Dev Cell 18: 999-1011.
38. Woodcock KJ, Kierdorf K, Pouchelon CA, Vivancos V, Dionne MS, et al.
(2015) Macrophage-derived upd3 cytokine causes impaired glucose
homeostasis and reduced lifespan in Drosophila fed a lipid-rich diet.
Immunity 42: 133-144.
39. Ayyaz A, Li H, Jasper H (2015) Haemocytes control stem cell activity in
the Drosophila intestine. Nat Cell Biol 17: 736-748.
40. Lunstrum GP, Bachinger HP, Fessler LI, Duncan KG, Nelson RE, et al.
(1988) Drosophila basement membrane procollagen IV. I. Protein
characterization and distribution. J Biol Chem 263: 18318-18327.
41. Kusche-Gullberg M, Garrison K, MacKrell AJ, Fessler LI, Fessler JH
(1992) Laminin A chain: expression during Drosophila development
and genomic sequence. EMBO J 11: 4519-4527.
42. Grigorian M, Liu T, Banerjee U, Hartenstein V (2013) The proteoglycan
Trol controls the architecture of the extracellular matrix and
balances proliferation and differentiation of blood progenitors in the
Drosophila lymph gland. Dev Biol 384: 301-312.
43. Leitao AB, Sucena E (2015) Drosophila sessile hemocyte clusters are true
hematopoietic tissues that regulate larval blood cell differentiation.
Elife 4.
44. Honti V, Csordas G, Markus R, Kurucz E, Jankovics F, et al. (2010) Cell
lineage tracing reveals the plasticity of the hemocyte lineages and of
the hematopoietic compartments in Drosophila melanogaster. Mol
Immunol 47: 1997-2004.
45. Avet-Rochex A, Boyer K, Polesello C, Gobert V, Osman D, et al. (2010) An
in vivo RNA interference screen identifies gene networks controlling
Drosophila melanogaster blood cell homeostasis. BMC Dev Biol 10:
65.
46. Stofanko M, Kwon SY, Badenhorst P (2010) Lineage tracing of
lamellocytes demonstrates Drosophila macrophage plasticity. PLoS
One 5: e14051.
47. Pham LN, Dionne MS, Shirasu-Hiza M, Schneider DS (2007) A specific
primed immune response in Drosophila is dependent on phagocytes.
PLoS Pathog 3: e26.
48. Kiger JA, Jr., Natzle JE, Green MM (2001) Hemocytes are essential for
wing maturation in Drosophila melanogaster. Proc Natl Acad Sci U S
A 98: 10190-10195.
49. Karpac J, Younger A, Jasper H (2011) Dynamic coordination of innate
immune signaling and insulin signaling regulates systemic responses
to localized DNA damage. Dev Cell 20: 841-854.
30
50. Kelsey EM, Luo X, Bruckner K, Jasper H (2012) Schnurri regulates
hemocyte function to promote tissue recovery after DNA damage. J
Cell Sci 125: 1393-1400.
51. Charroux B, Royet J (2009) Elimination of plasmatocytes by targeted
apoptosis reveals their role in multiple aspects of the Drosophila
immune response. Proc Natl Acad Sci U S A 106: 9797-9802.
52. Binggeli O, Neyen C, Poidevin M, Lemaitre B (2014) Prophenoloxidase
activation is required for survival to microbial infections in
Drosophila. PLoS Pathog 10: e1004067.
53. Bidla G, Dushay MS, Theopold U (2007) Crystal cell rupture after injury
in Drosophila requires the JNK pathway, small GTPases and the
TNF homolog Eiger. J Cell Sci 120: 1209-1215.
54. Chen Y, Liu F, Yang B, Lu A, Wang S, et al. (2012) Specific amino acids
affecting Drosophila melanogaster prophenoloxidase activity in
vitro. Dev Comp Immunol 38: 88-97.
55. Nam HJ, Jang IH, You H, Lee KA, Lee WJ (2012) Genetic evidence of a
redox-dependent systemic wound response via Hayan proteasephenoloxidase system in Drosophila. EMBO J 31: 1253-1265.
56. Rizki TM, Rizki RM (1992) Lamellocyte differentiation in Drosophila
larvae parasitized by Leptopilina. Dev Comp Immunol 16: 103-110.
57. Badenhorst P (2014) What can we learn from Flies: Epigenetic
Mechanisms Regulating Blood Cell Development in Drosophila. In:
C. Bonifer PNC, editor. Transcriptional and Epigenetic Mechanisms
Regulating Normal and Aberrant Blood Cell Development,
Epigenetics and Human Health. Berlin Heidelberg 2014: SpringerVerlag.
58. Makhijani K, Bruckner K (2012) Of blood cells and the nervous system:
hematopoiesis in the Drosophila larva. Fly (Austin) 6: 254-260.
59. Ghosh S, Singh A, Mandal S, Mandal L (2015) Active hematopoietic hubs
in Drosophila adults generate hemocytes and contribute to immune
response. Dev Cell 33: 478-488.
60. Abrams JM, Lux A, Steller H, Krieger M (1992) Macrophages in
Drosophila embryos and L2 cells exhibit scavenger receptormediated endocytosis. Proc Natl Acad Sci U S A 89: 10375-10379.
61. Tepass U, Fessler LI, Aziz A, Hartenstein V (1994) Embryonic origin of
hemocytes and their relationship to cell death in Drosophila.
Development 120: 1829-1837.
62. Holz A, Bossinger B, Strasser T, Janning W, Klapper R (2003) The two
origins of hemocytes in Drosophila. Development 130: 4955-4962.
63. Markus R, Laurinyecz B, Kurucz E, Honti V, Bajusz I, et al. (2009) Sessile
hemocytes as a hematopoietic compartment in Drosophila
melanogaster. Proc Natl Acad Sci U S A 106: 4805-4809.
64. Makhijani K, Alexander B, Tanaka T, Rulifson E, Bruckner K (2011) The
peripheral nervous system supports blood cell homing and survival
in the Drosophila larva. Development 138: 5379-5391.
65. Crozatier M, Meister M (2007) Drosophila haematopoiesis. Cell
Microbiol 9: 1117-1126.
31
66. Grigorian M, Mandal L, Hartenstein V (2011) Hematopoiesis at the onset
of metamorphosis: terminal differentiation and dissociation of the
Drosophila lymph gland. Dev Genes Evol 221: 121-131.
67. Jung SH, Evans CJ, Uemura C, Banerjee U (2005) The Drosophila lymph
gland as a developmental model of hematopoiesis. Development 132:
2521-2533.
68. Bretscher AJ, Honti V, Binggeli O, Burri O, Poidevin M, et al. (2015) The
Nimrod transmembrane receptor Eater is required for hemocyte
attachment to the sessile compartment in Drosophila melanogaster.
Biol Open 4: 355-363.
69. Goto A, Kadowaki T, Kitagawa Y (2003) Drosophila hemolectin gene is
expressed in embryonic and larval hemocytes and its knock down
causes bleeding defects. Dev Biol 264: 582-591.
70. Zettervall CJ, Anderl I, Williams MJ, Palmer R, Kurucz E, et al. (2004) A
directed screen for genes involved in Drosophila blood cell
activation. Proc Natl Acad Sci U S A 101: 14192-14197.
71. Anderl I, Hultmark D (2015) New ways to make a blood cell. Elife 4.
72. Kurucz E, Vaczi B, Markus R, Laurinyecz B, Vilmos P, et al. (2007)
Definition of Drosophila hemocyte subsets by cell-type specific
antigens. Acta Biol Hung 58 Suppl: 95-111.
73. Welman A, Serrels A, Brunton VG, Ditzel M, Frame MC (2010) Two-color
photoactivatable probe for selective tracking of proteins and cells. J
Biol Chem 285: 11607-11616.
74. Narita R, Yamashita H, Goto A, Imai H, Ichihara S, et al. (2004)
Syndecan-dependent binding of Drosophila hemocytes to laminin
alpha3/5 chain LG4-5 modules: potential role in sessile hemocyte
islets formation. FEBS Lett 576: 127-132.
75. Honti V, Csordas G, Kurucz E, Markus R, Ando I (2014) The cellmediated immunity of Drosophila melanogaster: hemocyte lineages,
immune compartments, microanatomy and regulation. Dev Comp
Immunol 42: 47-56.
76. Sorrentino RP, Tokusumi T, Schulz RA (2007) The Friend of GATA
protein U-shaped functions as a hematopoietic tumor suppressor in
Drosophila. Dev Biol 311: 311-323.
77. Munier AI, Doucet D, Perrodou E, Zachary D, Meister M, et al. (2002)
PVF2, a PDGF/VEGF-like growth factor, induces hemocyte
proliferation in Drosophila larvae. EMBO Rep 3: 1195-1200.
78. Duvic B, Hoffmann JA, Meister M, Royet J (2002) Notch signaling
controls
lineage
specification
during
Drosophila
larval
hematopoiesis. Curr Biol 12: 1923-1927.
79. Rizki TMR, R. M. (1980) Properties of the larval hemocytes of Drosophila
melanogaster. Experientia 36: 1223-1226.
80. Sorrentino RP, Carton Y, Govind S (2002) Cellular immune response to
parasite infection in the Drosophila lymph gland is developmentally
regulated. Dev Biol 243: 65-80.
81. Crozatier M, Ubeda JM, Vincent A, Meister M (2004) Cellular immune
response to parasitization in Drosophila requires the EBF orthologue
collier. PLoS Biol 2: E196.
32
82. Gateff E (1994) Tumor-suppressor genes, hematopoietic malignancies
and other hematopoietic disorders of Drosophila melanogaster. Ann
N Y Acad Sci 712: 260-279.
83. Qiu P, Pan PC, Govind S (1998) A role for the Drosophila Toll/Cactus
pathway in larval hematopoiesis. Development 125: 1909-1920.
84. Harrison DA, Binari R, Nahreini TS, Gilman M, Perrimon N (1995)
Activation of a Drosophila Janus kinase (JAK) causes hematopoietic
neoplasia and developmental defects. EMBO J 14: 2857-2865.
85. Stofanko M, Kwon SY, Badenhorst P (2008) A misexpression screen to
identify regulators of Drosophila larval hemocyte development.
Genetics 180: 253-267.
86. Minakhina S, Steward R (2006) Melanotic mutants in Drosophila:
pathways and phenotypes. Genetics 174: 253-263.
87. Mackenzie DK, Bussiere LF, Tinsley MC (2011) Senescence of the cellular
immune response in Drosophila melanogaster. Exp Gerontol 46:
853-859.
88. Horn L, Leips J, Starz-Gaiano M (2014) Phagocytic ability declines with
age in adult Drosophila hemocytes. Aging Cell 13: 719-728.
89. Carton Y BM, van Alphen JJM, van Lenteren JC (1986) The Drosophila
parasitic wasps. In: Edited by: Ashburner M CH, Thomson JN.
Sunderland, MA: Sinauer Associates, editor. The Genetics and
Biology of Drosophila pp. 347-394.
90. Allemand R, Lemaitre, C., Frey, F., Bouletreau, M., Vavre, F.,
Nordlander, G., van Alphen, J., Carton, Y. (2002) Phylogeny of six
African Leptopilina species (Hymenoptera: Cynipoidea, Figitidae),
parasitoids of Drosophila, with description of three new species. Ann
Soc entomol Fr (ns) 38: 14.
91. Stand MR (2008) The insect cellular immune response. Insect Science 15:
1-14.
92. Lee ML, Kalamarz, M. E., Paddibhatla, I., Small, C., Rajwani, R., Govind,
S. (2009) Virulence Factors and Strategies of Leptopilina spp.:
Selective Responses in Drosophila Hosts. Advances in Parasitology:
Elsevier Ltd. pp. 123-145.
93. Russo JD, S; Frey, F; Carton, Y, Brehelin, M (1996) Insect immunity:
early events in the encapsulation process of parasitoid (Leptopilina
boulardi) eggs in resistant and susceptible strains of Drosophila.
Parasitology 112: 8.
94. Howell L, Sampson CJ, Xavier MJ, Bolukbasi E, Heck MM, et al. (2012)
A directed miniscreen for genes involved in the Drosophila antiparasitoid immune response. Immunogenetics 64: 155-161.
95. Mortimer NT, Goecks J, Kacsoh BZ, Mobley JA, Bowersock GJ, et al.
(2013) Parasitoid wasp venom SERCA regulates Drosophila calcium
levels and inhibits cellular immunity. Proc Natl Acad Sci U S A 110:
9427-9432.
96. Vanha-Aho LM, Anderl I, Vesala L, Hultmark D, Valanne S, et al. (2015)
Edin Expression in the Fat Body Is Required in the Defense Against
Parasitic Wasps in Drosophila melanogaster. PLoS Pathog 11:
e1004895.
33
97. Williams MJ, Ando I, Hultmark D (2005) Drosophila melanogaster Rac2
is necessary for a proper cellular immune response. Genes Cells 10:
813-823.
98. Irving P, Ubeda JM, Doucet D, Troxler L, Lagueux M, et al. (2005) New
insights into Drosophila larval haemocyte functions through
genome-wide analysis. Cell Microbiol 7: 335-350.
99. Williams MJ (2009) The Drosophila cell adhesion molecule Neuroglian
regulates
Lissencephaly-1
localisation
in
circulating
immunosurveillance cells. BMC Immunol 10: 17.
100. Sampson CJ, Valanne S, Fauvarque MO, Hultmark D, Ramet M, et al.
(2012) The RhoGEF Zizimin-related acts in the Drosophila cellular
immune response via the Rho GTPases Rac2 and Cdc42. Dev Comp
Immunol 38: 160-168.
101. Xavier MJ, Williams MJ (2011) The Rho-family GTPase Rac1 regulates
integrin localization in Drosophila immunosurveillance cells. PLoS
One 6: e19504.
102. Williams MJ, Wiklund ML, Wikman S, Hultmark D (2006) Rac1
signalling in the Drosophila larval cellular immune response. J Cell
Sci 119: 2015-2024.
103. Mortimer NT, Kacsoh BZ, Keebaugh ES, Schlenke TA (2012) Mgat1dependent N-glycosylation of membrane components primes
Drosophila melanogaster blood cells for the cellular encapsulation
response. PLoS Pathog 8: e1002819.
104. Kurucz E, Zettervall CJ, Sinka R, Vilmos P, Pivarcsi A, et al. (2003)
Hemese, a hemocyte-specific transmembrane protein, affects the
cellular immune response in Drosophila. Proc Natl Acad Sci U S A
100: 2622-2627.
105. Rizki RMR, T. M. (1990) Encapsulation of parasitoid eggs in
phenoloxidase-deficient mutants of Drosophila melanogaster. J
Insect Physiol 36: 7.
106. Eslin P, Prevost, G. (1996) Variation in Drosophila Concentration of
Haemocytes Associated with Different Ability to Encapsulate
Asobara tabida Larval Parasitoid. J Insect Physiol 42: 6.
107. Eslin P, Prevost G (1998) Hemocyte load and immune resistance to
Asobara tabida are correlated in species of the Drosophila
melanogaster subgroup. J Insect Physiol 44: 807-816.
108. Dupas S, Morand S, Eslin P (2004) Evolution of hemocyte
concentration in the melanogaster subgroup species. C R Biol 327:
139-147.
109. Kacsoh BZ, Schlenke TA (2012) High hemocyte load is associated with
increased resistance against parasitoids in Drosophila suzukii, a
relative of D. melanogaster. PLoS One 7: e34721.
110. Gerritsma S, Haan A, Zande L, Wertheim B (2013) Natural variation in
differentiated hemocytes is related to parasitoid resistance in
Drosophila melanogaster. J Insect Physiol 59: 148-158.
111. Eslin P, Doury G (2006) The fly Drosophila subobscura: a natural case of
innate immunity deficiency. Dev Comp Immunol 30: 977-983.
112. Rubin GM, Lewis EB (2000) A brief history of Drosophila's
contributions to genome research. Science 287: 2216-2218.
34
113. Morgan TH (1917) The Theory of the Gene. The American Naturalist 51:
513-544.
114. Nusslein-Volhard C, Wieschaus E (1980) Mutations affecting segment
number and polarity in Drosophila. Nature 287: 795-801.
115. Spradling AC, Rubin GM (1982) Transposition of cloned P elements into
Drosophila germ line chromosomes. Science 218: 341-347.
116. Rubin GM, Spradling AC (1982) Genetic transformation of Drosophila
with transposable element vectors. Science 218: 348-353.
117. Hiesinger PR, Bellen HJ (2004) Flying in the face of total disruption.
Nat Genet 36: 211-212.
118. Bellen HJ, Levis RW, Liao G, He Y, Carlson JW, et al. (2004) The BDGP
gene disruption project: single transposon insertions associated with
40% of Drosophila genes. Genetics 167: 761-781.
119. Venken KJ, Bellen HJ (2007) Transgenesis upgrades for Drosophila
melanogaster. Development 134: 3571-3584.
120. Golic KG, Golic MM (1996) Engineering the Drosophila genome:
chromosome rearrangements by design. Genetics 144: 1693-1711.
121. Theodosiou NA, Xu T (1998) Use of FLP/FRT system to study
Drosophila development. Methods 14: 355-365.
122. Griffin R, Binari R, Perrimon N (2014) Genetic odyssey to generate
marked clones in Drosophila mosaics. Proc Natl Acad Sci U S A 111:
4756-4763.
123. Brand AH, Perrimon N (1993) Targeted gene expression as a means of
altering cell fates and generating dominant phenotypes.
Development 118: 401-415.
124. McGuire SE, Le PT, Osborn AJ, Matsumoto K, Davis RL (2003)
Spatiotemporal rescue of memory dysfunction in Drosophila.
Science 302: 1765-1768.
125. McGuire SE, Roman G, Davis RL (2004) Gene expression systems in
Drosophila: a synthesis of time and space. Trends Genet 20: 384391.
126. McGuire SE, Mao Z, Davis RL (2004) Spatiotemporal gene expression
targeting with the TARGET and gene-switch systems in Drosophila.
Sci STKE 2004: pl6.
127. Lee T, Luo L (1999) Mosaic analysis with a repressible cell marker for
studies of gene function in neuronal morphogenesis. Neuron 22:
451-461.
128. Lee T, Luo L (2001) Mosaic analysis with a repressible cell marker
(MARCM) for Drosophila neural development. Trends Neurosci 24:
251-254.
129. Venken KJ, He Y, Hoskins RA, Bellen HJ (2006) P[acman]: a BAC
transgenic platform for targeted insertion of large DNA fragments in
D. melanogaster. Science 314: 1747-1751.
130. Bassett AR, Tibbit C, Ponting CP, Liu JL (2013) Highly efficient targeted
mutagenesis of Drosophila with the CRISPR/Cas9 system. Cell Rep
4: 220-228.
131. Bassett AR, Liu JL (2014) CRISPR/Cas9 and genome editing in
Drosophila. J Genet Genomics 41: 7-19.
35
132. St Johnston D (2002) The art and design of genetic screens: Drosophila
melanogaster. Nat Rev Genet 3: 176-188.
133. Roote J, Russell S (2012) Toward a complete Drosophila deficiency kit.
Genome Biol 13: 149.
134. Ryder E, Blows F, Ashburner M, Bautista-Llacer R, Coulson D, et al.
(2004) The DrosDel collection: a set of P-element insertions for
generating custom chromosomal aberrations in Drosophila
melanogaster. Genetics 167: 797-813.
135. Ryder E, Ashburner M, Bautista-Llacer R, Drummond J, Webster J, et
al. (2007) The DrosDel deletion collection: a Drosophila
genomewide chromosomal deficiency resource. Genetics 177: 615629.
136. Parks AL, Cook KR, Belvin M, Dompe NA, Fawcett R, et al. (2004)
Systematic generation of high-resolution deletion coverage of the
Drosophila melanogaster genome. Nat Genet 36: 288-292.
137. Golic MM, Rong YS, Petersen RB, Lindquist SL, Golic KG (1997) FLPmediated DNA mobilization to specific target sites in Drosophila
chromosomes. Nucleic Acids Res 25: 3665-3671.
138. Thibault ST, Singer MA, Miyazaki WY, Milash B, Dompe NA, et al.
(2004) A complementary transposon tool kit for Drosophila
melanogaster using P and piggyBac. Nat Genet 36: 283-287.
139. Cook RK, Christensen SJ, Deal JA, Coburn RA, Deal ME, et al. (2012)
The generation of chromosomal deletions to provide extensive
coverage and subdivision of the Drosophila melanogaster genome.
Genome Biol 13: R21.
140. Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, et al. (1998)
Potent and specific genetic interference by double-stranded RNA in
Caenorhabditis elegans. Nature 391: 806-811.
141. Perrimon N, Ni JQ, Perkins L (2010) In vivo RNAi: today and tomorrow.
Cold Spring Harb Perspect Biol 2: a003640.
142. Yamamoto-Hino M, Goto S (2013) In Vivo RNAi-Based Screens: Studies
in Model Organisms. Genes (Basel) 4: 646-665.
143. Bier E (2005) Drosophila, the golden bug, emerges as a tool for human
genetics. Nat Rev Genet 6: 9-23.
144. Wu LP, Choe KM, Lu Y, Anderson KV (2001) Drosophila immunity:
genes on the third chromosome required for the response to
bacterial infection. Genetics 159: 189-199.
145. Mondal BC, Shim J, Evans CJ, Banerjee U (2014) Pvr expression
regulators in equilibrium signal control and maintenance of
Drosophila blood progenitors. Elife 3: e03626.
146. Tan KL, Goh SC, Minakhina S (2012) Genetic screen for regulators of
lymph gland homeostasis and hemocyte maturation in Drosophila.
G3 (Bethesda) 2: 393-405.
147. Tokusumi Y, Tokusumi T, Shoue DA, Schulz RA (2012) Gene regulatory
networks controlling hematopoietic progenitor niche cell production
and differentiation in the Drosophila lymph gland. PLoS One 7:
e41604.
36
148. Rorth P (1996) A modular misexpression screen in Drosophila detecting
tissue-specific phenotypes. Proc Natl Acad Sci U S A 93: 1241812422.
149. Ramond E, Meister M, Lemaitre B (2015) From Embryo to Adult:
Hematopoiesis along the Drosophila Life Cycle. Dev Cell 33: 367368.
150. Rodriguez A, Zhou Z, Tang ML, Meller S, Chen J, et al. (1996)
Identification of immune system and response genes, and novel
mutations causing melanotic tumor formation in Drosophila
melanogaster. Genetics 143: 929-940.
151. Braun A, Lemaitre B, Lanot R, Zachary D, Meister M (1997) Drosophila
immunity: analysis of larval hemocytes by P-element-mediated
enhancer trap. Genetics 147: 623-634.
152. Kurucz E, Markus R, Zsamboki J, Folkl-Medzihradszky K, Darula Z, et
al. (2007) Nimrod, a putative phagocytosis receptor with EGF
repeats in Drosophila plasmatocytes. Curr Biol 17: 649-654.
153. Asha H, Nagy I, Kovacs G, Stetson D, Ando I, et al. (2003) Analysis of
Ras-induced overproliferation in Drosophila hemocytes. Genetics
163: 203-215.
154. Rus F, Kurucz E, Markus R, Sinenko SA, Laurinyecz B, et al. (2006)
Expression pattern of Filamin-240 in Drosophila blood cells. Gene
Expr Patterns 6: 928-934.
155. Sinenko SA, Mathey-Prevot B (2004) Increased expression of
Drosophila tetraspanin, Tsp68C, suppresses the abnormal
proliferation of ytr-deficient and Ras/Raf-activated hemocytes.
Oncogene 23: 9120-9128.
156. Stramer B, Wood W, Galko MJ, Redd MJ, Jacinto A, et al. (2005) Live
imaging of wound inflammation in Drosophila embryos reveals key
roles for small GTPases during in vivo cell migration. J Cell Biol 168:
567-573.
157. Tokusumi T, Shoue DA, Tokusumi Y, Stoller JR, Schulz RA (2009) New
hemocyte-specific enhancer-reporter transgenes for the analysis of
hematopoiesis in Drosophila. Genesis 47: 771-774.
158. Bruckner K, Kockel L, Duchek P, Luque CM, Rorth P, et al. (2004) The
PDGF/VEGF receptor controls blood cell survival in Drosophila. Dev
Cell 7: 73-84.
159. Olofsson B, Page DT (2005) Condensation of the central nervous system
in embryonic Drosophila is inhibited by blocking hemocyte
migration or neural activity. Dev Biol 279: 233-243.
160. Kocks C, Cho JH, Nehme N, Ulvila J, Pearson AM, et al. (2005) Eater, a
transmembrane protein mediating phagocytosis of bacterial
pathogens in Drosophila. Cell 123: 335-346.
161. Lebestky T, Chang T, Hartenstein V, Banerjee U (2000) Specification of
Drosophila hematopoietic lineage by conserved transcription factors.
Science 288: 146-149.
162. Nelson RE, Fessler LI, Takagi Y, Blumberg B, Keene DR, et al. (1994)
Peroxidasin: a novel enzyme-matrix protein of Drosophila
development. EMBO J 13: 3438-3447.
37
163. Franc NC, Dimarcq JL, Lagueux M, Hoffmann J, Ezekowitz RA (1996)
Croquemort, a novel Drosophila hemocyte/macrophage receptor
that recognizes apoptotic cells. Immunity 4: 431-443.
164. Rehorn KP, Thelen H, Michelson AM, Reuter R (1996) A molecular
aspect of hematopoiesis and endoderm development common to
vertebrates and Drosophila. Development 122: 4023-4031.
165. Kroeger PT, Jr., Tokusumi T, Schulz RA (2012) Transcriptional
regulation of eater gene expression in Drosophila blood cells.
Genesis 50: 41-49.
166. Tokusumi T, Sorrentino RP, Russell M, Ferrarese R, Govind S, et al.
(2009) Characterization of a lamellocyte transcriptional enhancer
located within the misshapen gene of Drosophila melanogaster.
PLoS One 4: e6429.
167. Gajewski KM, Sorrentino RP, Lee JH, Zhang Q, Russell M, et al. (2007)
Identification of a crystal cell-specific enhancer of the black cells
prophenoloxidase gene in Drosophila. Genesis 45: 200-207.
168. Honti V, Kurucz E, Csordas G, Laurinyecz B, Markus R, et al. (2009) In
vivo detection of lamellocytes in Drosophila melanogaster. Immunol
Lett 126: 83-84.
169. Csordas G, Varga GI, Honti V, Jankovics F, Kurucz E, et al. (2014) In
vivo immunostaining of hemocyte compartments in Drosophila for
live imaging. PLoS One 9: e98191.
170. Evans CJ, Liu T, Banerjee U (2014) Drosophila hematopoiesis: Markers
and methods for molecular genetic analysis. Methods 68: 242-251.
171. Tirouvanziam R, Davidson CJ, Lipsick JS, Herzenberg LA (2004)
Fluorescence-activated cell sorting (FACS) of Drosophila hemocytes
reveals important functional similarities to mammalian leukocytes.
Proc Natl Acad Sci U S A 101: 2912-2917.
172. Clark RI, Woodcock KJ, Geissmann F, Trouillet C, Dionne MS (2011)
Multiple TGF-beta superfamily signals modulate the adult
Drosophila immune response. Curr Biol 21: 1672-1677.
173. Vilmos P, Nagy I, Kurucz E, Hultmark D, Gateff E, et al. (2004) A rapid
rosetting method for separation of hemocyte sub-populations of
Drosophila melanogaster. Dev Comp Immunol 28: 555-563.
174. Jiang H, Edgar BA (2009) EGFR signaling regulates the proliferation of
Drosophila adult midgut progenitors. Development 136: 483-493.
175. Sorrentino RP, Melk JP, Govind S (2004) Genetic analysis of
contributions of dorsal group and JAK-Stat92E pathway genes to
larval hemocyte concentration and the egg encapsulation response in
Drosophila. Genetics 166: 1343-1356.
176. Wertheim B, Kraaijeveld AR, Schuster E, Blanc E, Hopkins M, et al.
(2005) Genome-wide gene expression in response to parasitoid
attack in Drosophila. Genome Biol 6: R94.
177. Schlenke TA, Morales J, Govind S, Clark AG (2007) Contrasting
infection strategies in generalist and specialist wasp parasitoids of
Drosophila melanogaster. PLoS Pathog 3: 1486-1501.
178. Sinenko SA, Mandal L, Martinez-Agosto JA, Banerjee U (2009) Dual
role of wingless signaling in stem-like hematopoietic precursor
maintenance in Drosophila. Dev Cell 16: 756-763.
38
179. Delaney JR, Stoven S, Uvell H, Anderson KV, Engstrom Y, et al. (2006)
Cooperative control of Drosophila immune responses by the JNK
and NF-kappaB signaling pathways. EMBO J 25: 3068-3077.
180. Tanji T, Hu X, Weber AN, Ip YT (2007) Toll and IMD pathways
synergistically activate an innate immune response in Drosophila
melanogaster. Mol Cell Biol 27: 4578-4588.
181. Kim LK, Choi UY, Cho HS, Lee JS, Lee WB, et al. (2007) Downregulation of NF-kappaB target genes by the AP-1 and STAT complex
during the innate immune response in Drosophila. PLoS Biol 5:
e238.
182. Rizki RM, Rizki TM (1984) Selective destruction of a host blood cell type
by a parasitoid wasp. Proc Natl Acad Sci U S A 81: 6154-6158.
183. Rizki RM, Rizki TM (1990) Parasitoid virus-like particles destroy
Drosophila cellular immunity. Proc Natl Acad Sci U S A 87: 83888392.
184. Carton Y BM, van Alphen JJM, van Lenteren JC (1986) The Drosophila
parasitic wasps. In: Ashburner M CL, Thompson JN, editor. The
Genetics and Biology of Drosophila. London: Academic Press. . pp.
347–394.
185. Herzenberg LA, Parks D, Sahaf B, Perez O, Roederer M, et al. (2002)
The history and future of the fluorescence activated cell sorter and
flow cytometry: a view from Stanford. Clin Chem 48: 1819-1827.
186. De Rosa SC, Brenchley JM, Roederer M (2003) Beyond six colors: a new
era in flow cytometry. Nat Med 9: 112-117.
187. Takizawa H, Boettcher S, Manz MG (2012) Demand-adapted regulation
of early hematopoiesis in infection and inflammation. Blood 119:
2991-3002.
188. Lazzaro BP (2015) Adenosine signaling and the energetic costs of
induced immunity. PLoS Biol 13: e1002136.
189. Andres AJ, Cherbas P (1992) Tissue-specific ecdysone responses:
regulation of the Drosophila genes Eip28/29 and Eip40 during
larval development. Development 116: 865-876.
190. Cubitt AB, Heim R, Adams SR, Boyd AE, Gross LA, et al. (1995)
Understanding, improving and using green fluorescent proteins.
Trends Biochem Sci 20: 448-455.
191. Li X, Zhao X, Fang Y, Jiang X, Duong T, et al. (1998) Generation of
destabilized green fluorescent protein as a transcription reporter. J
Biol Chem 273: 34970-34975.
192. Sinenko SA, Shim J, Banerjee U (2012) Oxidative stress in the
haematopoietic niche regulates the cellular immune response in
Drosophila. EMBO Rep 13: 83-89.
193. Sinenko SA, Hung T, Moroz T, Tran QM, Sidhu S, et al. (2010) Genetic
manipulation of AML1-ETO-induced expansion of hematopoietic
precursors in a Drosophila model. Blood 116: 4612-4620.
194. Rizki TM, Rizki RM, Carton Y (1990) Leptopilina heterotoma and L.
boulardi: strategies to avoid cellular defense responses of Drosophila
melanogaster. Exp Parasitol 70: 466-475.
195. Labrosse C, Eslin P, Doury G, Drezen JM, Poirie M (2005) Haemocyte
changes in D. Melanogaster in response to long gland components of
39
the parasitoid wasp Leptopilina boulardi: a Rho-GAP protein as an
important factor. J Insect Physiol 51: 161-170.
196. Labrosse C, Stasiak K, Lesobre J, Grangeia A, Huguet E, et al. (2005) A
RhoGAP protein as a main immune suppressive factor in the
Leptopilina
boulardi
(Hymenoptera,
Figitidae)-Drosophila
melanogaster interaction. Insect Biochem Mol Biol 35: 93-103.
197. Colinet D, Dubuffet A, Cazes D, Moreau S, Drezen JM, et al. (2009) A
serpin from the parasitoid wasp Leptopilina boulardi targets the
Drosophila phenoloxidase cascade. Dev Comp Immunol 33: 681689.
198. Colinet D, Cazes D, Belghazi M, Gatti JL, Poirie M (2011) Extracellular
superoxide dismutase in insects: characterization, function, and
interspecific variation in parasitoid wasp venom. J Biol Chem 286:
40110-40121.
199. Sampson CJ, Williams MJ (2012) Real-time analysis of Drosophila postembryonic haemocyte behaviour. PLoS One 7: e28783.
200. Evans IR, Wood W (2014) Drosophila blood cell chemotaxis. Curr Opin
Cell Biol 30: 1-8.
201. Evans IR, Rodrigues FS, Armitage EL, Wood W (2015) Draper/CED-1
Mediates an Ancient Damage Response to Control Inflammatory
Blood Cell Migration In Vivo. Curr Biol 25: 1606-1612.
202. Gordon MD, Dionne MS, Schneider DS, Nusse R (2005) WntD is a
feedback inhibitor of Dorsal/NF-kappaB in Drosophila development
and immunity. Nature 437: 746-749.
203. Gordon MD, Ayres JS, Schneider DS, Nusse R (2008) Pathogenesis of
listeria-infected Drosophila wntD mutants is associated with
elevated levels of the novel immunity gene edin. PLoS Pathog 4:
e1000111.
204. Meister M, Ferrandon D (2012) Immune cell transdifferentiation: a
complex crosstalk between circulating immune cells and the
haematopoietic niche. EMBO Rep 13: 3-4.
205. Erdelyi M, Szabad J (1989) Isolation and characterization of dominant
female sterile mutations of Drosophila melanogaster. I. Mutations
on the third chromosome. Genetics 122: 111-127.
206. Schneider DS, Hudson KL, Lin TY, Anderson KV (1991) Dominant and
recessive mutations define functional domains of Toll, a
transmembrane protein required for dorsal-ventral polarity in the
Drosophila embryo. Genes Dev 5: 797-807.
207. Dietzl G, Chen D, Schnorrer F, Su KC, Barinova Y, et al. (2007) A
genome-wide transgenic RNAi library for conditional gene
inactivation in Drosophila. Nature 448: 151-156.
208. Shi S, Calhoun HC, Xia F, Li J, Le L, et al. (2006) JAK signaling globally
counteracts heterochromatic gene silencing. Nat Genet 38: 10711076.
209. Locksley RM, Killeen N, Lenardo MJ (2001) The TNF and TNF receptor
superfamilies: integrating mammalian biology. Cell 104: 487-501.
210. Dinarello CA (2007) Historical insights into cytokines. Eur J Immunol
37 Suppl 1: S34-45.
40
211. Kawai T, Akira S (2011) Toll-like receptors and their crosstalk with other
innate receptors in infection and immunity. Immunity 34: 637-650.
212. Valanne S, Kleino A, Myllymaki H, Vuoristo J, Ramet M (2007) Iap2 is
required for a sustained response in the Drosophila Imd pathway.
Dev Comp Immunol 31: 991-1001.
213. Mabery EM, Schneider DS (2010) The Drosophila TNF ortholog eiger is
required in the fat body for a robust immune response. J Innate
Immun 2: 371-378.
214. Lindsay SA, Wasserman SA (2014) Conventional and non-conventional
Drosophila Toll signaling. Dev Comp Immunol 42: 16-24.
215. Brennan CA, Delaney JR, Schneider DS, Anderson KV (2007) Psidin is
required in Drosophila blood cells for both phagocytic degradation
and immune activation of the fat body. Curr Biol 17: 67-72.
216. Dijkers PF, O'Farrell PH (2007) Drosophila calcineurin promotes
induction of innate immune responses. Curr Biol 17: 2087-2093.
217. Agaisse H, Petersen UM, Boutros M, Mathey-Prevot B, Perrimon N
(2003) Signaling role of hemocytes in Drosophila JAK/STATdependent response to septic injury. Dev Cell 5: 441-450.
218. Parisi F, Stefanatos RK, Strathdee K, Yu Y, Vidal M (2014) Transformed
epithelia trigger non-tissue-autonomous tumor suppressor response
by adipocytes via activation of Toll and Eiger/TNF signaling. Cell
Rep 6: 855-867.
219. Tsuzuki S, Ochiai M, Matsumoto H, Kurata S, Ohnishi A, et al. (2012)
Drosophila growth-blocking peptide-like factor mediates acute
immune reactions during infectious and non-infectious stress. Sci
Rep 2: 210.
220. Tsuzuki S, Matsumoto H, Furihata S, Ryuda M, Tanaka H, et al. (2014)
Switching between humoral and cellular immune responses in
Drosophila is guided by the cytokine GBP. Nat Commun 5: 4628.
221. Wertheim B, Kraaijeveld AR, Hopkins MG, Walther Boer M, Godfray
HC (2011) Functional genomics of the evolution of increased
resistance to parasitism in Drosophila. Mol Ecol 20: 932-949.
222. Gueguen G, Kalamarz ME, Ramroop J, Uribe J, Govind S (2013)
Polydnaviral ankyrin proteins aid parasitic wasp survival by
coordinate and selective inhibition of hematopoietic and immune
NF-kappa B signaling in insect hosts. PLoS Pathog 9: e1003580.
223. Ferrandon D, Jung AC, Criqui M, Lemaitre B, Uttenweiler-Joseph S, et
al. (1998) A drosomycin-GFP reporter transgene reveals a local
immune response in Drosophila that is not dependent on the Toll
pathway. EMBO J 17: 1217-1227.
41