KOSAKONIA & CHROMOBACTERIUM VS. MALARIA &
DENGUE – INSIGHTS INTO ANTIPATHOGENIC STRATEGIES
OF MOSQUITO MIDGUT BACTERIA
by Raúl G. Saraiva
A dissertation submitted to Johns Hopkins University in conformity with the requirements for the degree of Doctor of Philosophy.
Baltimore, Maryland March, 2018
© Raúl G. Saraiva 2018 All rights reserved
ABSTRACT
Malaria and dengue are arguably today’s most relevant tropical diseases vectored by Anopheles and Aedes spp. mosquitoes, respectively. Both pathogens enter the vector through a blood meal from an infected individual and inhabit the mosquito’s midgut even if briefly, encountering a plethora of antipathogenic factors. To those, there is an unequivocal contribution of the mosquito gut microbiota, as discussed herein.
In this work, we explore the role of individual members of the mosquito gut microbiota in limiting the vector’s ability to acquire and transmit Plasmodium spp. and the dengue virus. A
Kosakonia isolate from an anopheline in Zambia is shown to interfere with Plasmodium development by inducing the shutdown of the parasite’s antioxidant response. A
Chromobacterium species isolated from Aedes aegypti in Panama secretes an aminopeptidase that can ablate infectivity by the dengue virus, while also secreting the histone deacetylase romidepsin that can prevent P. falciparum maturation in Anopheles gambiae.
These findings constitute the first mechanistic descriptions on how certain members of the mosquito microbiota may exercise a pivotal role in the control of disease transmission, generating key knowledge towards the understanding of the dynamics of both malaria and dengue. At the same time, strategies can be devised to employ our conclusions towards the development of novel antimalarial and anti-DENV regimens, or towards the deployment of microbiota-shifting approaches by which the enrichment in pathogen-protective bacteria within field mosquito populations is promoted.
ii ACKNOWLEDGEMENTS
I would like to express my sincere gratitude for everyone who guide me, helped me and supported me throughout this project:
Dr. George Dimopoulos for taking me as an advisee and understanding something fruitful would come out of our complementary expertises.
Drs. Jürgen Bosch, Marcelo Jacobs-Lorena, Cynthia Sears, Photini Sinnis and Craig Townsend for all their contributions as members of my advising committee; the collaboration of Drs. Bosch and Townsend was instrumental for the success of this study.
Present and past members of the Dimopoulos group for the guidance and camaraderie, especially those that are co-authors in resulting publications for their hardwork and dedication.
The MMI department for its people and for the financial support to my journey, together with
Boehringer Ingelheim Fonds, Fulbright and Emergent Biosolutions.
The Fulbright Association, PAPS and JHSPH Student Assembly for accepting me as a volunteer and superbly complementing my student experience.
My friends here, back home and across the world for making this a richer adventure than I could ever expect.
My parents and family for all the love.
iii TABLE OF CONTENTS
CHAPTER 1. Introduction ...... 1
1.1. Malaria parasites encounter a bottleneck in the Anopheles midgut ...... 2
1.1.1. The peritrophic matrix and epithelial barriers ...... 4
1.1.2. Immunosurveillance ...... 7
1.1.3. A complement-like defense system ...... 8
1.1.4. The melanization defense system ...... 10
1.1.5. Oxidative and nitrosative stresses in anti-Plasmodium defense ...... 11
1.1.6. Other defense effectors ...... 13
1.1.7. The influence of midgut microbiota on Plasmodium infection ...... 15
1.2. Dengue infection is limited by factors in the mosquito gut ...... 17
1.2.1. Antiviral defense pathways ...... 18
1.2.2. Apoptosis of host cells as a potential antiviral mechanism ...... 21
1.2.3. Role of midgut bacteria in the anti-DENV response ...... 21
1.3. Bacterial natural products in the fight against mosquito-borne disease ...... 22
1.3.1. Antimalarials of bacterial origin ...... 22
1.3.2. Bacteria-produced metabolites as dengue antivirals ...... 26
1.3.3. A role for metabolites produced by the mosquito gut microbiota? ...... 27
1.4. Aims of this work ...... 29
iv Authors’ contributions...... 30
CHAPTER 2. Functional genomic analyzes of Kosakonia, Anopheles and Plasmodium reciprocal interactions that impact vector competence ...... 31
2.1. Summary ...... 32
2.2. Background ...... 34
2.3. Methodology ...... 36
2.3.1. Mosquito rearing and antibiotic treatment ...... 36
2.3.2. Bacteria cocktail preparation ...... 36
2.3.3. Introduction of bacteria through sugar or blood meals ...... 37
2.3.4. Anopheles gambiae midgut selection of Kco_Z ...... 37
2.3.5. Colonization experiments and DNA extraction...... 37
2.3.6. Absolute qPCR quantification ...... 38
2.3.7. Relative qPCR quantification ...... 39
2.3.8. Longevity, fecundity, and fertility assays ...... 40
2.3.9. Plasmodium infection assays ...... 41
2.3.10. Kco_Z genome sequencing and transcriptome analysis ...... 41
2.3.11. In vitro parasite culture and co-culture with bacteria ...... 42
2.4. Results ...... 44
2.4.1. Colonization of the Anopheles gambiae midgut by Kco_Z ...... 44
v 2.4.2. Inhibition of Plasmodium sporogonic development following sugar-meal introduction
of Kco_Z ...... 45
2.4.3. Kco_Z influence on mosquito fitness parameters ...... 46
2.4.4. Kco_Z genome analysis ...... 48
2.4.5. Transcriptomic survey of Kco_Z genes that mediate selection in the mosquito midgut
environment ...... 48
2.4.6. Transcriptomic survey for Kco_Z-encoded Plasmodium inhibition factors ...... 51
2.4.7. Influence of Kco_Z on Plasmodium antioxidant gene expression ...... 53
2.5. Discussion ...... 56
Authors’ contributions...... 58
CHAPTER 3. Chromobacterium spp. mediate their antimalarial activity through secretion of the HDAC inhibitor romidepsin...... 59
3.1. Summary ...... 60
3.2. Background ...... 61
3.3. Methodology ...... 63
3.3.1. Bacterial cultures and n-butanol extraction ...... 63
3.3.2. Plasmodium falciparum strains and cultivation ...... 63
3.3.3. Fast Performance Liquid Chromatography ...... 63
3.3.4. In vitro antimalarial assays ...... 64
3.3.5. Mass spectrometry ...... 65
vi 3.3.6. Genome curation and comparisons...... 65
3.3.7. Romidepsin biosynthetic gene cluster analysis ...... 66
3.3.8. Anopheles gambiae rearing and Plasmodium infection assays ...... 66
3.3.9. Chromobacterium spp. phylogenetic analysis ...... 67
3.3.10. P. falciparum nuclear extracts and HDAC inhibition measurements ...... 67
3.4. Results ...... 69
3.4.1. Anti-Plasmodium activity of Chromobacterium sp. Panama fractionates with
romidepsin ...... 69
3.4.2. Antimalarial Chromobacterium spp. are closely related genetically and encode for
romidepsin production ...... 72
3.4.3. Romidepsin has potent anti-Plasmodium activity ...... 74
3.4.4. Romidepsin production is required for Chromobacterium spp. anti-Plasmodium
properties ...... 75
3.4.5. Romidepsin is active against drug resistant isolates and inhibits P. falciparum HDAC
activity ...... 77
3.5. Discussion ...... 80
Authors’ contributions...... 83
CHAPTER 4. Aminopeptidase secreted by Chromobacterium sp. Panama inhibits dengue virus infection by degrading its E protein ...... 84
4.1. Summary ...... 85
4.2. Methodology ...... 86
vii 4.2.1. Cell culture maintenance, virus propagation and bacterial growth ...... 86
4.2.2. Dengue virus titration ...... 86
4.2.3. Solvent extractions of bacterial supernatants ...... 87
4.2.4. Protein extract preparation ...... 87
4.2.5. Pre- and post-virus attachment neutralization assays ...... 88
4.2.6. Transmission electron microscopy ...... 88
4.2.7. Protein gels and Western blotting ...... 88
4.2.8. Immunofluorescence microscopy ...... 89
4.2.9. Hydrophobic interaction chromatography ...... 89
4.2.10. Proteomics and protein sequence analysis ...... 90
4.2.11. Protease inhibiton ...... 90
4.3. Results ...... 92
4.3.1. C. sp. Panama culture supernatant shows potent anti-DENV activity ...... 92
4.3.2. Anti-DENV activity is associated with C. sp. Panama-produced proteinous factor(s)
...... 92
4.3.3. C. sp. Panama supernatant proteins inhibit DENV attachment to host cells ...... 93
4.3.4. C. sp. Panama interferes with DENV attachment by mediating degradation of the E
protein ...... 94
4.3.5. Secreted proteases are present in active anti-DENV protein fraction ...... 95
4.3.6. Aminopeptidase secreted by C. sp. Panama mediates its anti-DENV activity ...... 96
viii 4.4. Discussion ...... 100
4.5. Authors’ contributions ...... 102
CHAPTER 5. Concluding Remarks ...... 103
TABLES ...... 109
FIGURES ...... 122
ABBREVIATIONS ...... 155
REFERENCES ...... 158
ix LIST OF TABLES
Table 1.1. Examples of compounds with antimalarial activity isolated from bacteria ...... 110
Table 2.1. Colonization-related candidate genes that are differentially regulated between the
parental (P1) and passaged (P3) Kco_Z strains ...... 115
Table 2.2. ROS-producing candidate genes that are differentially regulated between Kco_Z
grown in ookinete medium conducive to ROS production, and LB medium ...... 116
Table 3.1. Bacterial strains in this study ...... 117
Table 3.2. Plasmodium falciparum strains in this study ...... 118
Table 3.3. Sources of genome sequences of the Chromobacterium species/strains analyzed in this
study ...... 119
Table 3.4. Nucleotide alignment results (megaBLAST) querying putative biosynthetic gene
clusters detected in the C. sp. Panama genome by antiSMASH against the genomes of the
remaining Chromobacterium spp. used in this study ...... 120
Table 4.1. Proteins recovered from active anti-DENV fraction of C. sp. Panama ...... 121
x LIST OF FIGURES
Figure 1.1. Plasmodium interactions with the mosquito gut...... 123
Figure 1.2. Antiviral immunity in the mosquito gut...... 124
Figure 2.1. Colonization of the Anopheles gambiae midgut by Kco_Z and impact on Plasmodium
falciparum sporogonic development...... 126
Figure 2.2. Sugar-meal introduction of Kco_Z has no impact on Anopheles gambiae fitness ... 128
Figure 2.3. Kco_Z genome ...... 129
Figure 2.4. Differential gene expression in Kco_Z on different culture media and upon serial
passage in the Anopheles gambiae midgut ...... 130
Figure 2.5. Exposure to Kco_Z limits Plasmodium’s antioxidant response ...... 132
Figure 2.S1. Impact of blood feeding Kco_Z on mosquito fitness ...... 134
Figure 3.1. Bioassay-guided fractionation of Chromobacterium sp. Panama culture supernatant
points to romidepsin as the main antimalarial compound ...... 135
Figure 3.2. Antimalarial activity of chromobacteria is restricted to the romidepsin-producing
subcluster comprised of C. sp. Panama and C. haemolyticum ...... 138
Figure 3.3. Romidepsin strongly inhibits Plasmodium ...... 140
Figure 3.4. Romidepsin production is necessary and sufficient for the antimalarial activity of
Chromobacterium spp...... 141
Figure 3.5. Romidepsin surpasses common drug resistance mechanisms and interferes with P.
falciparum HDAC activity ...... 143
Figure 3.S1. UPLC chromatograms of Fraction F ...... 145
Figure 3.S2. Predicted NRPS organization and function of the Sch 20561/2 biosynthetic cluster
...... 146
xi Figure 3.S3. Tentative structural assignments of C. sp Panama-isolated Sch 20561/2 ...... 147
Figure 3.S4. Survival curves of Anopheles gambiae (females only, males only and both) upon
ingestion of Chromobacterium sp. 968 wildtype or ∆depA at approximately 106 CFU/mL .. 148
Figure 4.1. Anti-DENV activity of Chromobacterium sp. Panama is mediated by secreted
proteinous factors ...... 149
Figure 4.2. Protein extracts of culture supernatants of C. sp. Panama inhibit DENV entry by
mediating degradation of the E protein ...... 151
Figure 4.3. Purification of the anti-DENV proteins of C. sp. Panama ...... 153
Figure 4.4. Aminopeptidase secreted by C. sp. Panama induces DENV E protein degradation,
thereby inhibiting viral entry ...... 154
xii
CHAPTER 1. Introduction
1 Mosquitoes are the deadliest animals in the world mediating the death of approximately 725,000-
1,000,000 people per year [1]. Rather than playing the role of cruel assassins, mosquitoes tend to act as seemingly innocent carriers of numerous clinically relevant pathogens that exploit these insects’ feeding habits in order to complete their life cycles. Sometimes this cohabitation of mosquito and pathogen actually comes at a cost to the invertebrate host. Significant efforts are being made to understand the biology of mosquitoes and to develop potential means of interfering with their vector competence to stop the spread of devastating diseases such as malaria and dengue – the most prevalent vector-transmitted illnesses, affecting annually an estimated combined 500 million people worldwide [2], [3].
All pathogens transmitted by mosquitoes are acquired as part of a blood meal and, at some point during their life cycle, they must reside and invade the mosquito gut in order to migrate to their developmental site or their site for transmission to the human host. In the digestive tract, both parasites and viruses will encounter a resident microbiota believed to be acquired stochastically from the larval stages and as the mosquito feeds on nectar sources throughout its adult life [4].
This tripartite interaction between mosquitoes, their symbionts and the pathogens they transmit is believed to be crucial to determine the ability of the vector to harbor a productive infection [4].
In this chapter, we explore malaria, dengue and the role mosquitoes and their gut microbiota can play in controlling disease transmission. Also discussed is the contribution of bacterial metabolites in the fight against tropical disease, and how natural product development remains a critical component of current drug discovery.
1.1. Malaria parasites encounter a bottleneck in the Anopheles midgut
2 About 40 anopheline species can act as vectors for five human Plasmodium parasites –
Plasmodium falciparum (P. falciparum), P. vivax, P. ovale, P. malariae, and P. knowlesi.
According to the latest reports, 3.3 billion people are at risk for this disease, resulting in an estimated 212,000,000 cases of infection and 429,000 deaths per year, a vast majority of which occurring as a result of P. falciparum infection of children under the age of 5 in sub-Saharan
Africa [2].
Malaria transmission, however, is far from a trivial process. When feeding on blood of an infected human, the female mosquito takes up male and female Plasmodium gametocytes that mature into micro- and macrogametes, respectively, resulting in fertilization and the establishment of zygotes. The zygotes then transform into motile ookinetes, which invade and traverse the mosquito midgut epithelium about a day after blood uptake in order to reach the basal side of the epithelium, where they develop into oocysts. Approximately 14 days after the ingestion of the blood meal, these structures release thousands of sporozoites that migrate to the mosquito salivary glands and render the mosquito infectious. The timing of these various stages vary slightly depending on environmental factors and the specific Plasmodium-Anopheles pairing [5].
Thus, during a short but crucial period, the parasites inhabit the lumen of the mosquito midgut, and must invade its lining epithelium in order to establish a productive infection. Perhaps not surprisingly, this invasion of the epithelium proves to be a challenge and constitutes a major bottleneck for Plasmodium development, such that among the thousands of ookinetes, only approximately a half dozen will successfully mature to oocysts [6], [7]. This drastic reduction is believed to be the result of the presence of physical barriers and active immune responses
3 established by the mosquito, in combination with potential detrimental interactions with the residing midgut microbiota [4].
It is important to note that not all anopheline species can harbor and transmit all Plasmodium species; this selectivity provides a tool for identifying immune responses of interest. The studies detailed here were primarily conducted using the African vector An. gambiae and the Asian vector An. stephensi, infected with either P. falciparum or one of two rodent models, P. berghei or P. yoelli. The establishment of these systems has resulted from a combination of their public health relevance and their ease of manipulation in laboratory settings. Given the variability between and also within each mosquito and parasite species, experimental findings tend to require further confirmation in a number of mosquito-parasite combinations before they can be generalized [8]–[10].
1.1.1. The peritrophic matrix and epithelial barriers
As ookinetes mature into oocysts, they encounter two important physical barriers. The first is known as the peritrophic matrix, and the second is the midgut epithelium itself. It should be noted, however, that the parasites can be targeted before they even differentiate into ookinetes, while they inhabit the lumen of the Anopheles midgut. Recent reviews have highlighted the roles of human-derived factors and the resident mosquito microbiota in this process [4], [11]. As regards mosquito-secreted digestive enzymes, the general belief is that the early mosquito stages of the malaria parasite are generally protected, since lytic enzyme production peaks at ~24 h after a blood meal [12], [13], and these enzymes diffuse relatively slowly through the blood bolus; this temporal and spatial separation results in the survival of the majority of the parasites [14], [15].
4 The peritrophic matrix (PM) is a thick acellular sheath composed mainly of chitin fibrils and glycoproteins [16], [17] that develops after blood-meal ingestion and envelops the blood bolus, separating it from the midgut epithelium. Thus, the PM has direct contact with several gut- derived stimuli that range from bacterial commensals to pathogens and from products of digestion to toxins. To preserve homeostasis, this matrix establishes a selective barrier, permeable to nutrients but forming a first line of defense against detrimental agents such as the
Plasmodium parasite [18], [19]. Unlike what happens with digestive enzymes, there is both a temporal and spatial correlation between PM maturation and ookinete invasion: the PM is believed to achieve its peak thickness of 1 to 20 μm approximately 24 h after a blood meal [14],
[20], precisely the timeframe for ookinete traversal.
It has been well established that in order to successfully cross the PM, the parasite secretes micronemal chitinases that enzymatically degrade this structure, allowing for parasite invasion; genetic or chemical disruption of these chitinases severely affects the ability of Plasmodium to invade the mosquito tissue [21]–[25]. However, these parasite-encoded chitinases seem to only partially explain the success of ookinete PM invasion. In fact, it has been proposed that a human- derived chitinase, chitotriosidase, also aids in the process [26], [27]. Mosquitoes also produce a chitinase that is thought to be involved in the regulation of PM secretion, and this enzyme may paradoxically assist with parasite invasion [28], [29]. A comprehensive follow-up study to the proteomic characterization of the PM in An. gambiae is still needed and would be important for understanding the potential role of non-chitin components in PM traversal by Plasmodium.
However, there is increasing evidence for the importance of proteinaceous components of the
PM, such as fibrinogen-related protein-1 (FREP1) [30], [31], as mediators of parasite invasion.
5 After traversing the PM, the ookinetes encounter the midgut epithelium, which they must cross before lodging themselves next to its basement membrane and developing into oocysts. The evidence collected thus far indicates that the ookinetes exhibit gliding motility along the surface of the epithelium before they initiate midgut invasion [32], [33]. The authors of a recent study have postulated that there are multiple receptors expressed by the epithelial cells that can modulate this invasion process, and they have identified an enolase-binding protein (EBP) as one of them [34].
Once a topic of debate, the overwhelming amount of evidence that now exists seems to indicate that ookinetes take an intracellular rather than intercellular route while crossing the epithelium
[18], [35]. In fact, several studies have shown that ookinetes usually invade multiple midgut epithelial cells before reaching the basement membrane, and each of these cells undergoes apoptosis upon invasion (Figure 1.1) [32], [35], [36]. Two models have arisen to explain this phenomenon. The first, called the “time-bomb” theory, postulates that, after ookinete invasion, midgut epithelial cells protrude into the midgut lumen, express high levels of NOS, acquire abnormal nuclear morphology, fragment their DNA, and radically reorganize their actin cytoskeleton, which ultimately leads to cell death and extrusion of the dying cell from the epithelium [36]. Hence, the “time-bomb”: ookinetes would have a limited window of time to emerge unharmed from these cells, so they employ lateral movement as a strategy to migrate from cell to cell before they ultimately reach the basal side of the epithelium [36].
Another model builds upon these findings but proposes a “cellular treadmill” instead. Rather than supporting lateral movement of ookinetes, it proposes that the parasite always moves from the luminal to the basal side of the epithelial cell. If cell extrusion happens, the ookinete encounters the apical side of the replacing epithelial cell as it exits the dying cell and invades it,
6 resetting the invasion process. That way, a “treadmill” is established, with the parasite needing to traverse multiple epithelial cells before it finally reaches its destination [35].
1.1.2. Immunosurveillance
Both before and after traversing the epithelium, the ookinetes are believed to encounter both free and membrane-bound pattern recognition receptors (PRRs) that can recognize pathogen- associated molecular patterns (PAMPs) and either promote downstream signaling-mediated immune activation, or directly induce endocytic processes. Even at the early stages in the blood bolus, the parasites appear to be susceptible to targeting by mosquito-produced factors. In a study by Dong et al. (2011), immune deficiency (IMD) pathway activation was found to significantly limit ookinete numbers prior to invasion of the midgut epithelium, suggesting that many of the effector molecules also play an antiparasitic role within the midgut lumen. In addition to the
IMD pathway, other major immune signaling pathways that modulate pathogen responses in the mosquito after PRR activation are the Toll, Jun-N-terminal kinase (JNK), and Janus kinase/signal transducers and activators of transcription (JAK-STAT) pathways [37], [38].
Among the membrane-bound PRRs, one of the most relevant for anti-Plasmodium defense is the peptidoglycan recognition protein LC (PGRP-LC), a receptor for the IMD signaling pathway
[39]–[41]. Whereas the IMD pathway has emerged as the most effective pathway against P. falciparum, a role has also been proposed for both the Toll and JAK-STAT pathways. For the three pathways, infection with the parasite results in a signaling cascade whose mediators have been comprehensively discussed in the recent past in the context of malaria transmission [42],
[43]. Protein-level negative regulators of these pathways have similarly been extensively described, but recently a role for microRNAs in sequence-specific post-transcriptional regulation has also been uncovered [44]. aga-miR-305 in An. gambiae was found to be upregulated upon P.
7 falciparum infection and to target immune genes associated with the IMD pathway, in what is thought to be a mechanism to prevent over-allocation of resources to the host defense response.
Activation of these innate immune pathways in the mosquito culminates in the expression of important PRRs such as FBN9, TEP1, APL1, LRIM1, and LRRD7, as well as other effector genes some of which also responsible for parasite killing.
Dong et al. (2006) identified several putative PRRs involved in the responses of both P. berghei and P. falciparum in An. gambiae. Among these PRRs were AgMDL1, TEP1, FBN8, FBN9,
FBN39, and LRRD7; AgMDL1 and FBN39 appeared to be specific for P. falciparum [45]. It is important to note that the fibrinogen-related protein family (FBN or FREP) is arguably the largest PRR gene family in An. gambiae, with as many as 59 putative members. FBN9 has been identified as one of the most potent anti-Plasmodium FBN proteins, acting against both the rodent and human malaria parasites, with co-localization studies strongly suggesting that its defensive activity involves direct interaction with the pathogen [46]. More recently, the Down syndrome cell adhesion molecule (AgDscam) has also been shown to be protective against challenge with either P. berghei or P. falciparum. The AgDscam gene is a Toll and IMD pathway-regulated hypervariable PRR with the potential to generate over 30,000 alternative splicing forms, which are responsible for different pathogen interaction and defense specificities
[47]–[49].
1.1.3. A complement-like defense system
The ookinetes that eventually lodge themselves on the basal surface of the midgut epithelium become exposed to the mosquito’s circulatory system, the hemocoel. Of particular relevance are the hemolymph-circulating proteins that are considered to be evolutionary precursors of
8 components of the vertebrate complement system and produce a complement-like response in
Anopheles.
The first molecule identified in this context was the IMD pathway-regulated thioester protein 1
(TEP1), with its remarkable structural and functional similarities to the mammalian complement factor C3, including the ability to act as an opsonin and to promote phagocytosis of bacteria [50],
[51]. In the context of Plasmodium infection, TEP1 is believed to recognize and bind to the ookinete surface and tag the parasite for lysis and/or melanization (Figure 1.1) [50].
Interestingly, allelic variants in the TEP1 gene explain the differential susceptibility to P. berghei infection of distinct An. gambiae strains [50], [52].
TEP1, however, does not act alone. Two leucine-rich repeat (LRR) domain-containing proteins,
LRIM1 and APL1C, are believed to act as a dimer to chaperone mature TEP1, binding and stabilizing it in the hemolymph (Figure 1.1) [53], [54]. In fact, APL1C is encoded in a region of chromosome 2L that includes three paralogs (APL1A, APL1B, APL1C), APL1A having at least three allelic variants identified thus far in An. gambiae mosquitoes from Gambia, Cameroon, and
Mali [55]–[57]. Of particular relevance is the fact that different members of the APL1A-C locus appear to confer protection against different parasites [58], [59]. Initial studies showed that
APL1C was relevant to combating P. berghei or P. yoelli and that APL1A was the most active paralog against P. falciparum [58], but the same pattern does not seem to hold true if another strain of An. gambiae is considered [59], probably because of the high allelic diversity seen at this locus [43]. Recent evidence suggests that all APL1A-C can dimerize with LRIM1 [60], supporting the hypothesis that TEP1, LRIM1, and APL1A-C act in synergy against Plasmodium.
LRRD7 (or APL2), another member of the LRR domain-containing protein family, has been shown to have anti-Plasmodium activity [48], but its role in this complement-like system has not
9 yet been established. Other members of the TEP family [61] and the non-catalytic serine protease
SPCLIP1 [62] have been postulated to participate in this system, but a comprehensive cascade of events explaining the mechanism of deposition of these proteins onto the parasite’s surface has yet to be proposed. Similarly, it is still not clear how this protein complex achieves specificity in recognizing foreign pathogens or how it induces lysis of the ookinete, although early evidence suggests that these processes may occur via still-unidentified pore-forming protein complexes analogous to the vertebrate complement membrane attack complex [63].
1.1.4. The melanization defense system
As mentioned above, TEP1 has been identified as a marker for subsequent melanization of the invading ookinetes (Figure 1.1). In this process, melanin is deposited as a polymer surrounding the parasite and preventing gas diffusion, thus depriving the Plasmodium ookinete of oxygen.
Melanization is triggered by proteolytic activation of the prophenoloxidase (PPO) enzyme through a cascade of Clip-domain serine proteases (CLIP); the resulting catalytically active phenoloxidase oxidizes phenolic compounds such as tyrosine and DOPA into quinones, which are further converted to melanins [64], [65]. If a parasite is melanized, it remains in the mosquito’s midgut throughout the life of the insect [66].
The formation of a filamentous actin ring surrounding the dying or dead ookinete has also been described and associated with TEP1 deposition, and it is thought to be required for melanization to succeed [67]. In addition to TEP1, the Frizzled 2 (Fz2) and cell division cycle 42 (Cdc42) genes have been implicated in the formation of this actin zone [67], together with WASP, a positive regulator of actin polymerization [9], [63], [68]. Recent studies have also described this ookinete actin “hood” as also being linked to ookinete killing through lysis [69].
10 This process of melanization is thought to act mainly as a wound-healing response, isolating moribund parasites from the surrounding tissues, and therefore being primarily relevant to dead or damaged ookinetes [63], [64]. However, study of the C-type lectins CTL4 and CTLMA2 has shown that exacerbated melanization alone can limit the development of P. berghei in certain strains of An. gambiae [70], [71]. Similarly, studies focused on mosquito-derived serine protease inhibitors (serpins, SRPN) have also identified SRPN2 [72] and SRPN6 [73] as facilitators of midgut invasion by the Plasmodium parasites through inhibition of melanization, pointing to this encapsulation process as a cause, and not just a consequence, of ookinete death. While the melanization-dependent anti-Plasmodium defense system appears to be very effective, it is however rarely observed to be acting in natural field transmission conditions.
1.1.5. Oxidative and nitrosative stresses in anti-Plasmodium defense
In parallel to their involvement in melanotic encapsulation of Plasmodium, quinones also generate high local levels of reactive oxygen species (ROS) that are toxic to the parasite [64],
[74], [75]. ROS and reactive nitrogen species (RNS), the latter generated from the interaction of
− superoxide (O2 ) with NO, can also be produced as byproducts of mitochondrial respiration [76],
[77]. A blood meal alone induces ROS/RNS production, and Plasmodium traversal of the midgut epithelium is also believed to exacerbate ROS/RNS production. Different strains of An. gambiae have been shown to sustain different levels of Plasmodium infection related to the levels of
ROS/RNS they produce [74], [76], [78]. NO has also been postulated to be relevant for the killing of young oocysts [79]–[81].
Importantly, the parasite is not the only organism sensible to the oxidative and nitrosative stresses generated: mosquito cells are also susceptible to damage in such an environment, with notable effects on the insect’s longevity and fertility [82], [83]. In response to Plasmodium
11 infection, several detoxifying enzymes such as catalase and superoxide dismutase are upregulated in the mosquito to keep global levels of ROS/RNS in check [78], [84], and the oxidation resistance 1 (OXR1) gene in An. gambiae is involved in the regulation of the expression of these enzymes [85]. However, at the midgut level in infected mosquitoes, there is evidence of a specific reduction in catalase expression, which would increase the local amount of free radicals and result in a reduction in the parasite population [78]. ROS/RNS have also been shown to be important for activating further systemic innate immune responses [86], [87].
ROS/RNS production has been shown to be at least partially a function of genes such as the adenine nucleotide translocator (ANT) [76], the mitochondrial carrier 1 (AgMC1) [77], and the key signaling molecule Akt [88]. Induction of ROS/RNS has been mapped to blood-associated factors such as insulin-like growth factor 1 (IGF1) [89] and parasite-associated molecules such as a prenyl pyrophosphate (specifically, HMBPP) [90]. Interestingly, studies of P. vivax in An. aquasalis have shown no evidence of parasite-mediated induction of ROS, even though P. vivax was susceptible to artificial manipulation of ROS levels in this mosquito, which is one of its natural vectors [91].
Even before the ookinete reaches the basal side of the epithelium, nitrosative stress is believed to play a role in immunity against Plasmodium. Among the several alterations experienced by the invaded midgut epithelial cell, induction of nitric oxide synthase (NOS) is believed to be of particular relevance for controlling further development of the parasite. These findings were first described in P. berghei-infected An. stephensi [36], [80] and An. gambiae [92], but similar reports have surfaced regarding the more relevant P. vivax–An. aquasalis system [93].
The most recent studies propose that, upon invasion of rodent malaria parasites, expression of
NOS is upregulated in the targeted epithelial cells, leading to the intracellular pulse-release of
12 nitric oxide (NO). This effect is accompanied by the upregulation of the expression of two other enzymes, heme peroxidase 2 (HPX2) and NADPH oxidase 5 (NOX5), that ultimately enhance
NO toxicity [94]. The JNK pathway has since been implicated in regulating the expression of these enzymes [38]. This two-step process is called nitration and is believed to induce modifications in the surface of ookinetes that render them more likely to be recognized by TEP1, thus enhancing parasite clearance (Figure 1.1) [94].
Curiously, the same phenomenon has not been demonstrated in P. falciparum-infected mosquitoes, with reports pointing to a non-significant induction of NOS in the An. gambiae midgut after P. falciparum infection [95]. Since then, it has been proposed that the P. falciparum
Pfs47 gene is a key mediator of immune evasion, allowing the ookinete to escape nitration by disrupting JNK signaling [96], [97].
1.1.6. Other defense effectors
Many studies have postulated the role of other effector molecules in combating Plasmodium at the midgut level. More often than not, these effectors have been shown to interfere with the parasite load in the mosquito gut, but there is no clear idea yet as to their function and/or how they integrate with the better-understood mechanisms described above.
1.1.6.1. Antimicrobial peptides
Anopheles mosquitoes produce AMPs in response to challenge by a pathogen. The three main classes of AMPs are defensins, cecropins, and gambicin, the latter being the only one specific to mosquitoes [98], [99]. Within the midgut, these AMPs are predominantly expressed anteriorly
[98], [100], [101]. Defensins have been shown to have no antiparasitic activity against P. berghei
[102], even though they are induced upon infection with this parasite [101]. As for cecropins and
13 gambicin, not only are they both induced by P. berghei infection, but they also seem to have some degree of anti-P. berghei activity [98], [100], [103]. Much work has been carried out on understanding the regulation of AMP expression, but very little is known about the in vivo mechanisms of action of AMPs [99].
1.1.6.2. Lipid transporters
Reports have surfaced that point to a role in Plasmodium development for components of the mosquito lipid-transporting system, such as apolipoprotein D (APOD), apolipophorins, and their respective precursors [45], [68], [104]. A retinoid and fatty-acid binding glycoprotein, RFABG, a known precursor of apolipophorins I and II, is induced by P. berghei invasion and seems to facilitate its development [68]. Apolipophorin-III (ApoLp-III) itself has since been shown to behave similarly [104]. Silencing of APOD strongly reduces both P. falciparum and P. berghei development in An. gambiae, but APOD is only induced in the mosquito midgut after invasion by ookinetes of P. falciparum, and not P. berghei [45]. Several theories have emerged to explain these findings. Enhanced lipid trafficking is required for membrane protrusion and epithelial-cell motility, and thus these factors may be required for parasite traversal of the midgut epithelium, as discussed above. In addition, Plasmodium lacks certain pathways for de novo lipid biosynthesis and may employ the mosquito’s lipid machinery in an effort to scavenge lipids.
There are also reports of a direct interplay of lipophorin (Lp) with known components of the mosquito’s innate immune system: Lp appears to decrease the binding efficiency of TEP1 to
Plasmodium ookinetes, thus reducing TEP1-mediated parasite killing [105].
1.1.6.3. Cytoplasmic actin
14 In a recent report, the ubiquitous cytoplasmic actin has been proposed to be a novel extracellular immune factor in An. gambiae [106]. Silencing of the actin 5c gene results in increased P. falciparum infection at the midgut level via a mechanism that is distinct from the postulated antibacterial effect of the same gene [106]. A relationship has been suggested between this phenomenon and the formation of actin zones and ookinete hoods, as discussed above, but further studies are required to elucidate the regulation of actin externalization and its role as a
Plasmodium antagonist.
1.1.7. The influence of midgut microbiota on Plasmodium infection
Recent discoveries have stimulated a growing interest in the interaction between the malaria parasite and the mosquito midgut microbiota. Analysis of the Anopheles innate immune responses against Plasmodium has shown that mosquito’s antiparasitic responses are largely overlapping with those mounted to control bacterial challenges. Conversely, immune responses mounted by the mosquito against invading or endogenous midgut bacteria are mediated by genes associated with anti-Plasmodium immune response [39], [107]–[111] and, in most cases, lead to the elimination of a large number of parasites. Hence, the gut microbiota is able to modulate the vectorial capacity of the Anopheles mosquito through the innate immune system. Bacterial population expansion of the midgut following a blood meal coincides with Plasmodium infection of the epithelium, and an immune response mounted by the mosquito vector targets both types of invaders. Interactions taking place in the midgut during Plasmodium infection involve components of different taxa, including the host mosquito vector, vertebrate blood factors, and the parasitic or viral pathogen, as well as other symbiotic microbes. It has been shown that a potential over-activation of the mosquito’s innate immune response following a blood meal is controlled by a complex formed by the immune peroxidase (ImPer) and dual oxidase (DUOX)
15 enzymes, located between the epithelium and the peritrophic matrix of the mosquito midgut
[112]. This protecting network allows for the proliferation of Plasmodia and bacteria within the midgut, by minimizing their interaction with microbial immune elicitors that would overstimulate antimicrobial and antiparasitic immune responses.
The activation of the Anopheles immune system in response to bacterial challenge is to a significant degree regulated by the IMD pathway [39], [107], as in Drosophila [113]. In An. gambiae, the IMD pathway-regulated transcription factor REL2 has been shown to mediate this pathway’s immune response after the activation of PGRP-LC by the midgut bacteria, thereby modulating Plasmodium infection [39]. Transcription analysis using a microarray screen has revealed an overlap in Plasmodium- and bacteria-stimulated gene regulation [45]. In this screen, important anti-Plasmodium factors such as the PRRs TEP1, FBN9, and LRRD7, among others, have been found to control resistance to bacterial infection [45]. Another study, in which the susceptibility to P. falciparum was compared between septic and aseptic (antibiotic-treated) An. gambiae mosquitoes, the bacteria-depleted group was found to be significantly more susceptible to malaria infection [107]. Alternatively, co-feeding mosquitoes with bacteria and parasites resulted in a decreased number of P. falciparum oocysts in the midgut [107].
The fact that early activation of the IMD pathway limits ookinete maturation [114] may partially explain these findings, since recognition of midgut bacteria by PGRP-LC may result in the release of antimalarial effectors into the midgut lumen, limiting the parasite’s development before it invades the mosquito tissue. In addition, a recent study has shown that Plasmodium infection elicits an immune priming mechanism that enhances immunity to subsequent infection with the parasite and that this memory-like mechanism is dependent on the mosquito microbiota
[115], [116]. Given the very low incidence of consecutive exposures of mosquitoes to
16 Plasmodium in the field, and a likely negligible selective pressure from the parasite on the mosquito, it is quite possible that exposure of the mosquito to microbiota may be priming its immune system against the malaria parasite.
A further understanding of parasite-mosquito midgut microbiota interactions has recently been achieved through stable infection of An. stephensi with Wolbachia, a common maternally inherited mosquito bacterial endosymbiont [117]. It has been shown in transient infections that
Wolbachia can inhibit the development of the malaria parasite in Anopheles mosquitoes, possibly by stimulating a mosquito antiparasitic immune response [118], [119]. In the referred study
[117], it was shown that the Wolbachia wAlbB strain can form a stable symbiosis with An. stephensi, invade laboratory populations, and confer elevated resistance to P. falciparum infection, thus opening the path for the use of this interaction as a potential approach to permanently reduce the vectorial capacity of the major malaria vectors in the field.
1.2. Dengue infection is limited by factors in the mosquito gut
Mosquitoes transmit numerous arthropod-borne viruses (arboviruses) that threaten humans medically and economically. Dengue virus (DENV) is perhaps the most prominent example, as it symptomatically infects 100 million people annually, and 3.6 billion people are at risk of infection [3], [120]. DENV belongs to the Flaviviridae family and is transmitted by Aedes mosquitoes; other flaviviruses include yellow fever (YFV), West Nile (WNV) and Zika (ZIKV) viruses.
After DENV reaches the lumen of the mosquito midgut within uptake of an infected blood meal, it interacts with receptors on the midgut epithelial cells. For successful infection, the virus must penetrate these cells, replicate, and disseminate to the respective target organs, namely the
17 salivary glands, thus rendering the mosquito infectious. In this process, PRRs activate antiviral pathways such as the Toll, IMD, and JAK-STAT pathways (Figure 1.2).
Compared to the breadth of knowledge that has been accumulated over the years concerning mosquito anti-Plasmodium responses, information is still limited regarding similar mechanisms in anti-DENV defense. However, significant progress has been made over the last decade in identifying the immune signaling pathways that most significantly contribute to dampening viral loads in mosquitoes, as discussed below. When appropriate, responses to other flaviviruses are discussed in an attempt to distinguish between virus-specific or pan-viral effects.
1.2.1. Antiviral defense pathways
1.2.1.1. The Toll pathway
The Toll pathway is initiated when the cytokine Spätzle is activated by PRRs, which activate the
Toll transmembrane receptor. This activation initiates the degradation of Cactus, which is an inhibitor of the Nf-κb-like transcription factor Rel1, through the adaptor protein MyD88.
Degradation of Cactus enables the translocation of Rel1 to the nucleus and the subsequent transcription of effector genes [121]–[124].
The Toll pathway plays a key role in anti-DENV mechanisms in the Ae. aegypti midgut.
Infection with DENV in Ae. aegypti activates the main components of the Toll pathway, such as
Spätzle, Toll, and Rel1A [125], [126]. This anti-DENV function of the Toll pathway has been further supported by the silencing of negative and positive regulators of this pathway. Silencing of Cactus decreases DENV titers in the midgut, whereas MyD88 silencing significantly increases viral titers [125]. WNV and YFV are structurally similar to DENV. However, the role of the Toll pathway against WNV and YFV in mosquitoes is still unclear. The Spätzle-like cytokine SPZ5
18 has been found to be significantly (2.5-fold) downregulated in Ae. aegypti one day after YFV infection. YFV infection also causes downregulation of the Toll-like receptor TOLL1B, by 2.3- fold and 2.6-fold at 1 and 7 days after infection, respectively. However, in this last study the virus was inoculated by intrathoracic injection, which may not allow the induction of natural midgut immune responses. The same study showed that WNV did not significantly regulate the components of the Toll pathway [127].
1.2.1.2. The IMD pathway
As discussed above, the IMD pathway is well known to play a crucial role in insect defense against bacteria and P. falciparum in mosquitoes. In culicine mosquitoes such as Aedes, the IMD pathway has recently been shown to induce antiviral mechanisms against various arboviruses
[128]–[130]. A previous study has demonstrated that the activation of the IMD pathway by silencing Caspar, an inhibitor of transcription factor Rel2, does not negatively affect DENV in the midguts of DENV-susceptible Ae. aegypti mosquitoes [125]. However, a recent study has indicated that a cecropin-like antibacterial gene mediated by IMD is upregulated during DENV infection in the salivary glands of mosquitoes, implying that the IMD pathway is associated with
DENV infection [129]. A more recent study has shown that depletion of IMD transcripts in a
DENV-refractory strain significantly increases viral titers in mosquito midguts [130]. Taken together, these results suggest that the IMD pathway plays an anti-DENV role in mosquitoes, but in susceptible strains it may be operating at maximum capacity; hence, laboratory inhibition of the negative regulator cannot increase the viral titers further.
1.2.1.3. The JAK-STAT pathway
19 It is well known that the JAK-STAT pathway plays an important antiviral role in mammals
[131], [132]. Through the use of the Drosophila system, it has been shown that this pathway also has an antiviral role in invertebrates [133]. An antiviral role for the JAK-STAT pathway in mosquitoes has also been suggested by several studies. It has been shown that the expression of
JAK-STAT pathway-mediated DENV effector genes (DVRF) leads to a decrease in viral titers in the mosquito midgut, indicating an anti-DENV function for this pathway in Ae. aegypti [134].
Interestingly, one of the DVRFs contains several antifreeze and allergen domains that are not found in An. gambiae orthologs, suggesting that this gene may be a culicine mosquito-borne virus-specific antiviral gene.
The JAK-STAT pathway can also restrict WNV in Culex mosquitoes via Vago in an IMD pathway-dependent manner. Vago was first identified as an upregulated gene in virus-infected
Drosophila. The mechanism of action of Vago in mosquitoes has been characterized in Hsu cells
(Culex quinquefasciatus) [135], [136]. Transcription of Vago is dependent on Dicer2 (a component of the RNAi pathway) and Rel2 (IMD pathway). Vago is secreted and activates the
JAK-STAT pathway. Interestingly, this pathway is independent of the RNAi pathway and Dome
(a receptor for the JAK-STAT pathway) [136], indicating a new type of signaling pathway functioning through potential pathway crosstalk. The antiviral role of Vago has been tested in adult Culex pipiens f. quinquefasciatus, but the virus was delivered by intrathoracic injection in this study, which may have disguised the direct and immediate midgut responses [135], [136].
1.2.1.4. The RNAi pathway
RNAi plays an important antiviral role in mosquito defenses, although it is not part of a classical antiviral pathway. Silencing of RNAi components such as Dicer2, R2D2, and Ago2 in adult Ae. aegypti significantly decreases orally delivered DENV in midguts at 7 days after infection. These
20 experiments showed that RNAi plays an antiviral role, whereas DENV remains in the midguts of
Ae. aegypti [137].
1.2.2. Apoptosis of host cells as a potential antiviral mechanism
The role of apoptosis in mosquito antiviral systems remains unclear. Apoptotic cell death has been considered to be an immune defense mechanism that can limit viral replication and spread in vertebrates [138]. Apoptosis during WNV infection has been observed in the midguts of a
Culex pipiens-refractory strain, showing that apoptosis may be a causative factor in limiting viral dissemination [139]. A pro-apoptotic gene, Michelob_x, is rapidly induced during DENV infection only in an Ae. aegypti DENV-refractory strain and not in a susceptible strain [140].
Apoptosis occurring in refractory mosquitoes may be a virus-limiting factor in mosquito midguts. However, there is no direct evidence pointing to an antiviral role for this process. On the other hand, viruses may exploit apoptosis as a host factor that facilitates viral dissemination in mosquitoes.
1.2.3. Role of midgut bacteria in the anti-DENV response
As is true for the malaria parasite, interactions between DENV, the vector mosquito Ae. aegypti, and the midgut microbiota have also been reported to play a likely role in modulating the spread of DENV. Some bacteria have been shown to impair DENV infection in the mosquito midgut
[125], [141], [142], and antibiotic-treated mosquitoes can support a higher DENV titer than septic mosquitoes [125]. This difference is thought to be related to the lower expression level of immune-related AMPs observed in aseptic Ae. aegypti mosquitoes, as a result of a lesser microbial exposure, and therefore a diminished immune activation [125]. In parallel, it has been demonstrated that reducing gut bacteria by feeding antibiotics to adult Ae. aegypti mosquitoes
21 affects the digestion of the blood proteins, thus decreasing mosquito fecundity [143]. It has also been shown that the anti-bacterial Toll and JAK-STAT pathways control DENV infection in the mosquito [125], [134].
1.3. Bacterial natural products in the fight against mosquito-borne disease
Secondary metabolites of microbial origin have long been acknowledged as medically relevant, but their full potential remains largely unexploited. Of the countless natural compounds discovered thus far, only 5-10% have been isolated from microorganisms. At the same time, while whole genome sequencing has demonstrated that bacteria and fungi often encode for natural products, only a few genera have yet been mined for new compounds. This subchapter explores the contributions of bacterial natural products in tackling malaria and dengue. It highlights how molecules isolated from microorganisms ranging from marine cyanobacteria to mosquito endosymbionts can be explored for their antimicrobial effects. Pursuit of this mostly untapped source of chemical entities will hopefully result in new interventions against these tropical diseases, very much needed to combat the increase in the incidence of resistance.
1.3.1. Antimalarials of bacterial origin
Natural product research has been crucial in providing advances towards antimalarial therapies.
Some of the most relevant drugs against this disease, quinine and artemisinin, were first discovered and extracted from plants [144]. Together with their synthetically optimized derivatives, quinine and artemisinin are both still integral component of antimalarial regimens
[145].
In the realm of natural products, there have also been important contributions of compounds of microbial origin in malaria prevention and treatment. Most notably, tetracycline – originally
22 isolated from Streptomyces spp. – is to this day recommended by the WHO for both prophylaxis and treatment in certain groups even if doxycycline, a semisynthetic derivative, is favored [145]; tetracyclines act on the parasite by inhibiting expression of apicoplast genes [146]. Similarly, clindamycin also acts as a protein synthesis inhibitor and is recommended in combination with artesunate (an artemisinin) or quinine against both severe and uncomplicated malaria [145].
Clindamycin is a chlorinated derived of lincomycin, an antibiotic discovered from Streptomyces lincolnensis [147].
On the other hand, deferoxamine is a siderophore from Streptomyces pilosus that reached clinical trials as an antimalarial; the requirement for an intravenous administration, no evidence of protection from death in children and overall increased likelihood of adverse effects has led to a recommendation against the use of iron chelators in the treatment of malaria [148].
Fosmidomycin, however, is still being pursued as a potential new drug against malaria, with mixed but generally good results in clinical trials thus far [149]; fosmidomycin is an inhibitor of isoprenoid synthesis via the mevalonate pathway and was isolated from Streptomyces lavendulae
[150].
While these bacterially-produced compounds are or were being actively pursued as part of antimalarial therapies, many more have been described in the literature that show great potential for activity. A selection of those are summarized in Table 1.1; the list is not supposed to be exhaustive, but illustrative of the variety of bacterial metabolites that have been shown to possess anti-Plasmodium effects.
Of the compounds listed, many showed impressive selectivity indexes (SI, i.e. ratio between activity against mammalian cell line and against P. falciparum; a SI greater than 10 is usually considered of interest in initial screenings as it indicates activity is not due to general in vitro
23 toxicity [151]–[153]). That was the case of salinipostin A (SI > 1000), echinomycin (SI > 916), micrococcin (SI > 857), gramicidin D (SI = 789) and nostocarboline (SI = 623). Salinipostin A was isolated from a marine Actinobacteria of the Salinospora genus and showed an IC50 of 50 nM against P. falciparum W2, and activity was shown to be restricted to early vs. late asexual stages [154]. Salinipostins are stereoisomers of cyclophostins and cyclipostins and these compounds have been described for their ability to inhibit microbial lipases [155], [156]; no testing has been done, however, to evaluate if salinipostin A acts in a similar fashion against
Plasmodium and attempts at establishing their mode of action were inconclusive [154].
Echinomycin was also isolated from Actinobacteria (Streptomyces echinatus, among others
[157]) and tested against asexual P. falciparum CSC-1 to produce an IC50 of 3.70±0.01 nM. This drug is known to act as a transcription inhibitor and thus inhibit protein synthesis, and the same has been shown in P. falciparum [158]; side effects have been reported in phase I clinical trials
[159] which may preclude the recommendation of its use in the current formulation.
Both Actinobacteria (Micrococcus varians) and Firmicutes (Bacillus pumilus and
Staphylococcus equorum) are able to produce micrococcin [160] and this thiopeptide exhibited an IC50 of 35 nM against P. falciparum 3D7 by inhibiting protein synthesis in the parasite [161].
Thiostrepton, a related compound, presents a much higher IC50 against Plasmodium and, consequently, a lower selectivity index as seen in Table 1.1. Micrococcin has recently been proposed as a viable candidate for therapy against both mycobacteria [162] and Clostridium difficile [163].
Gramicidin D is a heterogeneous mixture of six antibiotic compounds produced by Bacillus brevis that act as channel-forming ionophores shown to have an IC50 of 1.9 pM against P. falciparum (Nigerian strain) [164]. This impressively low value has been justified with a
24 potential preference of this drug to integrate into the membrane of infected vs. healthy erythrocytes [164]. However, low oral availability of these peptides has precluded further applications other than that of a topical antibiotic [165].
A Nostoc species of Cyanobacteria produces the alkaloid nostocarboline with 194 nM IC50 against P. falciparum K1 [166]. Also showing an impressive selectivity index, this compound’s role as a protease inhibitor has been proposed, and there seems to be tropism of inhibitory activity towards chlorophytes [167], [168]; activity against Plasmodium could then be explained by nostocarboline targeting an apicoplast process, given the algal origin of this relict plastid.
As seen in Table 1.1, these are just a few select examples of an incredible variety of compounds described from bacteria of different phyla. To be noted, a majority of them stem from
Actinobacteria – where Streptomyces spp. are dominant – or Cyanobacteria. The biosynthetic potential of bacteria from these phyla has long been known and explored, and so it is not surprising to see a bias on compounds from these organisms, as they are more actively mined for secondary metabolites than others.
Also of note are the strategies some authors are employing to improve the activity profile of the compounds listed. Often, they resort to total or semi-synthesis to recreate the product of interest and introduce variations in certain radicals, in hopes of improving parameters such as selectivity and bioavailability. More creatively, some authors have employed mutasynthesis, or mutational biosynthesis, where chemical synthesis is coupled with molecular biology to generate derivatives of natural products of interest [169]. This is rather apparent in the case of pactamycin (Table 1.1,
IC50 = 14 nM, SI = 7): through the study of its natural biosynthetic pathway in Streptomyces pactum and directed genetic manipulation, Mahmud and colleagues were able to generate
25 pactamycin analogues that preserved their antimalarial activity but were significantly less toxic to mammalian cells [170], [171].
1.3.2. Bacteria-produced metabolites as dengue antivirals
Contrary to malaria, there is currently no approved therapy against DENV. Many approaches have been used in search for dengue antivirals, namely screening for activity against specific dengue enzymes or structure-based computational discovery, with limited success to date. In this context, the potential of also looking for active natural products has not been overlooked [172], but evidence for the role of bacterial metabolites is still scarce.
Of the limited examples available, bafilomycin arguably provides the one that is best characterized. As can been seen in Table 1.1, bafilomycin is a fermentation product of
Streptomyces spp. and has a significant antimalarial activity, somewhat compromised by its also rather significant cytotoxicity against mammalian cells. That is perhaps because bafilomycin acts as an inhibitor of the host’s vacuolar type H(+)-ATPase [173]; in doing so, however, it prevents dengue virus entry by preventing the acidification of endocytic vesicles [174]. These initial studies in vitro have been followed with in vivo evaluation of the antiviral activity of bafilomycin when fed or injected to Aedes aegypti mosquitoes; in either case, DENV titers were significantly reduced when the drug was administered prior to infection, reinforcing the notion that bafilomycin interferes with viral entry [175].
By means of a high-throughput screen, additional compounds of bacterial origin were identified for their anti-DENV effects [176]. Those included antimycin A from Streptomyces spp. with an
IC50 of 80±20 nM and a selectivity index of 58-102, whose activity has since been validated against additional arboviruses and the hepatitis C flavivirus also showing promising selectivity
26 indexes [177]. Also in the same screen, acetylspiramycin (from Streptomyces ambofaciens [178]) and colistin (from Paenibacillus polymyxa [179]) were assigned as anti-DENV, even though only the former was validated in a dose-response evaluation with an IC50 of 0.91±0.15 µM and no detected cytotoxic effect [176].
Recently, diketopiperazines isolated from Microbulbifer variabilis were evaluated for their anti-
DENV activity and the most promising one showed an IC50 of 11.2 µM and a selectivity index greater than 8.9 [180]. This compound, cyclo-(4-trans-hydroxy-L-proline-L-phenylalanine), had been reported before [181] and its analogue explored for antibacterial and antifungal activity
[182]–[184]. A mechanism of action for these activities has yet to be established, and the same is true for its antiviral effect.
1.3.3. A role for metabolites produced by the mosquito gut microbiota?
The adult mosquito is believed to acquire a gut microbiota primarily through ingesting bacteria in the water from the breeding site upon emergence, as well as through nectar feeds during its lifespan [185], [186]. These bacteria experiences rapid proliferation following a blood meal
[185], and the gut microbiota is usually dominated by one or two bacterial species, as has been consistently reported from metagenomic studies [187]–[189]. High-throughput sequencing of gut microbiota of different mosquito species has shown a consistent predominance of Proteobacteria,
Bacteroidetes and Actinobacteria in the gut of adult mosquitoes, dominated by the Acinetobacter,
Aeromonas, Asaia, Elizabethkingia, Enterobacter, Pantoea, Pseudomonas and Serratia genera
[188], [190]–[194].
As discussed above, the gut microbiota of the mosquito can modulate disease transmission by eliciting antibacterial immune responses from mosquito cells that overlap with their antiparasitic
27 and antiviral pathways. Another contribution may relate to resource competition, by which gut bacteria compete with the human pathogens for nutrients and other factors that are required for their development in the vector [4]. It is not unfeasible to conceive, however, that on top of this general contribution of the mosquito microbiota in priming the mosquito’s antipathogenic immune response or otherwise outcompeting these pathogens for resources, certain individual members of the mosquito’s gut bacterial community further aid in that effect by secreting metabolites that directly impact parasites, viruses or mosquito host factors required for their invasion/entry and replication.
Because the adult mosquito’s gut microbiota is believed to be stochastically acquired from the insect’s environment as it feeds on nectar sources, a true symbiotic relationship between these bacteria and their mosquito host has yet to be postulated. However, that bacteria of rather limited taxa are regularly found within mosquitoes of distinct species and geographical origins creates the notion that there may indeed exist a symbiotic association other than a commensal one between certain members of the gut microbiota and their mosquito host, with both benefiting from the relationship. At the same time, mosquito factors that govern gut microbiota composition are just now beginning to be understood, and further work is needed in this area to conclude on mutualistic relationships in this context [195].
In other insects, however, there have been reports of potential symbiotic associations [196]. One of the most striking examples is that of the relationship between Serratia marcescens and the trypanosomal vector Rhodnius spp., where the bacteria appears to independently aid in hemolysis of the insect’s blood meal and secrete prodiogisin, a secondary metabolite active against trypanosomes [197], [198]. This is arguably to the triatomine’s advantage, and one can speculate that the bacterium also benefits from colonizing such a nutrient-rich environment.
28 Evidence that the same may be true for mosquitoes has recently arisen. In a study by Cirimotich et al., an Enterobacter species (now Kosakonia; see Chapter 2) was isolated from Anopheles arabiensis in Zambia and was shown to interfere with the malaria parasite maturation within the mosquito vector without obvious fitness costs to the insect [199]. The effects on Plasmodium are independent of mosquito factors and seem to stem from the bacteria stimulating a highly oxidative environment within the mosquito gut [199]. Pseudomonas and Pantoea species have also been shown to limit Plasmodium development when they colonize the gut of Anopheles gambiae without significant costs to the vector [111].
Nonetheless, the antipathogenic activity of mosquito gut bacteria may not always be in the host’s best interest. S. marcescens association with An. gambiae has been shown to be deleterious to both the mosquito and P. falciparum development [111]. Similarly, a Chromobacterium species isolated from Aedes aegypti in Panama has been proven to limit the life-span of laboratory-reared
Ae. aegypti and An. gambiae, as well as directly interfere with Plasmodium growth and DENV replication [142]. These findings open yet another avenue to explore in the control of transmission of both malaria and dengue: can bacteria be exploited towards diminishing vectorial capacity of mosquitoes by simultaneously reducing their pathogen load and their survival?
1.4. Aims of this work
In the context of all that is discussed above, the present body of work aims at establishing molecular mechanisms by which individual members of the mosquito microbiota are able to interfere with pathogens within the vector. For that purpose, three tripartite systems are explored: the An. gambiae – Kosakonia cowanii Zambiae – Plasmodium (Chapter 2), the An. gambiae –
Chromobacterium sp. Panama – P. falciparum (Chapter 3) and the Ae. aegypti –
Chromobacterium sp. Panama – DENV (Chapter 4).
29 Authors’ contributions
Seokyoung Kang, Maria Luísa Simões, Yesseinia I. Angleró-Rodríguez and George Dimopoulos contributed to subchapters 1.1 and 1.2. The candidate coordinated all the efforts and was responsible for all final edits, as well as being the primary contributor towards the writing of the manuscript.
A version of subchapters 1.1 and 1.2 has been published as:
R. G. Saraiva, S. Kang, M. L. Simões, Y. I. Angleró-Rodríguez, and G. Dimopoulos, “Mosquito gut antiparasitic and antiviral immunity,” Dev Comp Immunol., vol. 64, p. 53, Nov. 2016.
Subchapter 1.3 is being considered for submission as:
R. G. Saraiva and G. Dimopoulos, “Bacterial natural products in the fight against mosquito- borne diseases”.
30
CHAPTER 2. Functional genomic analyzes of Kosakonia,
Anopheles and Plasmodium reciprocal interactions that impact
vector competence
31 2.1. Summary
The Kosakonia cowanii Zambiae (Kco_Z) bacterium, which is naturally harbored in the midgut of field mosquitoes, can inhibit the development of Plasmodium parasites prior to their invasion of the midgut epithelium through a mechanism that involves oxidative stress. Here, a multifaceted approach is used to study the tripartite interactions between the mosquito, Kco_Z and Plasmodium, towards the molecular dissection of the inhibitory mechanism, and to address the feasibility of using sugar-baited exposure of mosquitoes to the Kco_Z bacterium for interruption of malaria transmission.
The ability of Kco_Z to colonize Anopheles gambiae midguts harboring microbiota derived from wild mosquitoes was determined by qPCR targeting the 16S rRNA gene. Upon introduction of
Kco_Z via nectar feeding, the permissiveness of colonized mosquitoes to Plasmodium falciparum infection was determined, as well as the impact of Kco_Z on mosquito fitness parameters, such as longevity, number of eggs laid and number of larvae hatched. The genome of
Kco_Z was sequenced, and transcriptome analyses were performed to identify bacterial genes that are important for colonization of the mosquito midgut, as well as for reactive oxygen species
(ROS)-production. A gene expression analysis of members of the oxidative defense pathway of
Plasmodium berghei was also conducted to assess the parasite’s oxidative defense response to
Kco_Z exposure.
Kco_Z persisted for up to 4 days in the An. gambiae midgut after introduction via nectar feeding, and was able to significantly inhibit Plasmodium sporogonic development. Introduction of this bacterium did not adversely affect mosquito fitness. Candidate genes involved in the selection of a better fit Kco_Z to the mosquito midgut environment and in its ability to condition oxidative
32 status of its surroundings were identified, and parasite expression data indicated that Kco_Z is able to induce a partial and temporary shutdown of the ookinete’s antioxidant response.
Kco_Z is capable of inhibiting sporogonic development of Plasmodium in the presence of the mosquito’s native microbiota without affecting mosquito fitness. Several candidate Kco_Z genes are likely mediating midgut colonization and ROS production, and inhibition of Plasmodium development appears to involve a shutdown of the parasite’s oxidative defense system. A better understanding of the complex reciprocal tripartite interactions can facilitate the development and optimization of a Kco_Z-based malaria control strategy.
33 2.2. Background
Screening isolates of the midgut microbiota of Zambian field mosquitoes for Plasmodium- blocking activity has allowed the identification of a Kosakonia cowanii bacterium, Kco_Z, which is naturally harbored in the mosquito midgut tissue and can inhibit the development of
Plasmodium parasites prior to their invasion of the mosquito midgut epithelium, independent of the mosquito’s immune system [108]. Kco_Z was previously known as Esp_Z, or Enterobacter sp. Zambiae, as speciation assignment was done before the Enterobacter genus was split in 2013
[200] and the full genome for this isolate was available (cf. Figure 2.3). Kco_Z nearly eliminates the development of malaria parasites to the human-infective sporozoite stage, thereby showing potential as a transmission-blocking agent.
Even at low concentrations, Kco_Z, is effective at inhibiting parasites in the mosquito intestine through a mechanism involving reactive oxygen species (ROS) [108]. Plasmodium parasites are susceptible to changes in the redox balance of their environment, and they defend themselves against ROS via an upregulation of antioxidant genes [201]. Two antioxidant systems with partially overlapping functions have been described in Plasmodium parasites, the thioredoxin and glutathione systems (reviewed in [202]). These systems are required to neutralize the highly oxidative environments encountered in both human erythrocytes [202] and the mosquito midgut
[78]. The thioredoxin system is believed to be the main antioxidant system, since P. falciparum parasites lack a glutathione-dependent peroxidase [203], [204], suggesting that thioredoxin reduction of peroxiredoxin antioxidant enzymes plays a prominent role in protecting
Plasmodium from toxic ROS [201]. Peroxiredoxin antioxidant enzymes such as peroxiredoxin-1
(Tpx-1) and 1-cys-peroxiredoxin (1-Cys-prx) act as electron acceptors from thioredoxin-1 (Trx-
1), maintaining them in a reduced state [205]. Plasmodium falciparum Tpx-1 knockout lines
34 show an increased sensitivity to both ROS and reactive nitrogen species, but the deletion mutation is not lethal, suggesting redundancy in the parasite’s antioxidant system [206]. A glutathione peroxidase-like thioredoxin peroxidase (TPxGl) has also been shown to be part of the thioredoxin system in Plasmodium and is reduced by Trx-1 [207]. Inhibitors of the thioredoxin pathway have been investigated as potential antimalarial compounds, rendering the parasite more susceptible to oxidative stress [208].
Recently, reintroduction of several bacteria through the nectar meal, a vital mosquito energy source, has been demonstrated to inhibit Plasmodium sporogonic development, and methods for exposing mosquitoes to various agents through artificial nectars have been developed [111],
[142], [209]. Manipulation of the composition of the mosquito microbiota may offer a strategy to reduce the prevalence of Plasmodium-infected mosquitoes and, therefore, malaria transmission.
A multifaceted approach was used to test the feasibility of using sugar-baited exposure of mosquitoes to the Kco_Z bacterium for interruption of malaria transmission, by studying the reciprocal interactions between the bacterium and the mosquito and its microbiota, together with
Kco_Z’s impact on the parasite under laboratory conditions. The Kco_Z genes that are likely to be conducive to effective colonization of the mosquito midgut and ROS–mediated parasite inhibition were specifically assessed, along with the effect of Kco_Z exposure on the parasite’s oxidative defense system, mosquito fitness, and the ability of Kco_Z to compete with the endogenous bacteria of the mosquito midgut microbiota.
35 2.3. Methodology
2.3.1. Mosquito rearing and antibiotic treatment
Anopheles gambiae Keele strain mosquitoes were maintained under laboratory conditions at 27
°C and 80% humidity with a 14 h day/10 h night cycle. Larvae were reared on cat food pellets and ground fish food supplement. Adult mosquitoes were maintained on 10% sucrose and fed on mouse blood (mice were anesthetized with ketamine) for egg production. For clearance of the endogenous bacteria of laboratory mosquitoes, adult female mosquitoes were maintained on antibiotics immediately after eclosion (75 μg/mL gentamicin sulfate [Quality Biological,
Gaithersburg, MD, USA] and 100 μg/mL penicillin-streptomycin [Invitrogen, Carlsbad, CA,
USA] in a 10% sucrose solution ad libitum for three to four days). The sucrose-antibiotic solution was changed every 48 h. To minimize the impact of any residual antibiotics, the mosquitoes were allowed to feed ad libitum for 24 h on a sterile 10% sucrose solution following antibiotic treatment.
2.3.2. Bacteria cocktail preparation
The bacterial cocktail was prepared from bacterial isolates previously identified from wild An. arabiensis mosquitoes obtained through landing catches in Zambia [108]. Twelve species representing both Gram-negative and Gram-positive bacteria were selected, and mixed in equal proportions. Single bacterial colonies from LB plates were selected and used to inoculate overnight cultures, then used to seed fresh cultures and grown to an OD600 of 1.0. Species names and GenBank sequence accession numbers were as follows: Knoellia sp. JF690939.1;
Acinetobacter sp. JF690925.1; Bacillus sp. JF690926.1; Pseudomonas sp. JF690929.1;
Exiguobacterium sp. JF690932.1; Kocuria sp. JF690933.1; Pantoea sp. JF690934.1;
36 Pseudomonas sp. JF690935.1; Staphylococcus sp. JF690936.1; Arthrobacter sp. JF690937.1;
Comamonas sp. JF690938.1; Bacillus sp. JF690930.1.
2.3.3. Introduction of bacteria through sugar or blood meals
Bacteria were grown in either LB or ookinete medium overnight at 30 °C, used to seed fresh cultures, and then diluted to an OD600 of 1.0, pelleted by centrifugation (10 min, 3,000 rpm in a tabletop centrifuge), washed in 1x PBS, and finally resuspended in PBS. For sugar-meal introduction, bacteria were diluted in 3% sucrose to the concentration indicated in the text and provided to mosquitoes on moistened cotton strips. For blood-meal introduction, mosquitoes were allowed to membrane-feed on blood containing bacteria (10% bacterial solution, 40% blood, 50% human serum) at the concentration indicated in the text. Non-fed mosquitoes were removed within 24 h after blood-feeding.
2.3.4. Anopheles gambiae midgut selection of Kco_Z
The parental Kco_Z strain was provided to An. gambiae mosquitoes via blood meal at 109
CFU/ml (P1). Engorged mosquitoes were maintained under standard insectary conditions, and 10 mosquitoes from this cohort were sampled daily for the presence of Kco_Z in the midgut. The
Kco_Z bacteria that were found in the midgut for the longest time were then used to challenge a second mosquito population, and the process was repeated to isolate the “most fit” Kco_Z bacteria for midgut colonization (P2). This process of serial passage was then repeated a third time to isolate the midgut-selected Kco_Z bacteria (P3).
2.3.5. Colonization experiments and DNA extraction
To establish the ability of Kco_Z to colonize midguts harboring a resident microbiota, 70-80 female mosquitoes were treated with antibiotics as described above. Cocktail bacteria (106
37 CFU/ml) were introduced ad libitum for 48 h via sugar meal; the mosquitoes were then starved for 6-8 h, and Kco_Z (106 CFU/ml) was introduced by either blood-feeding or sugar meal. Every
24 h, midguts were dissected from 10 individual mosquitoes and transferred to a centrifuge tube containing 100 µl PBS, then stored at -80 °C for DNA extraction; three independent replicates were performed. Mosquitoes were surface-sterilized by washing them in 100% ethanol for 2 min and then rinsing them for 1 min in sterile 1x PBS. DNA extractions were carried out as previously described [210] with minor modifications: in brief, dissected midguts were homogenized in 90 µl PBS, followed by the addition of 90 µl lysozyme (40 mg/ml) and incubation at 37 °C for 1 h; 300 µl of extraction buffer (1% SDS; 50 mM Tris-HCl, pH 8.0; 25 mM NaCl; 25 mM EDTA, pH 8.0) was added to samples and incubated at 65 °C for 10 min.
Following the addition of 200 µl of 3 M K acetate (pH 7.2), samples were incubated on ice for 1 h and then centrifuged at 14,000 rpm in a tabletop centrifuge for 10 min and the supernatants removed. The pellets were then treated with RNase (40 mg/ml) at room temperature for 30 min.
The supernatants were extracted twice with phenol/chloroform/isoamyl alcohol and precipitated with 2.5 volumes of 100% ethanol and 1:10 3M Na acetate at -20 °C overnight; the pellets were resuspended in 10 µl of water.
2.3.6. Absolute qPCR quantification
All mosquito tissues were stored at -80 °C following dissection, and DNA was extracted as described. Absolute qRT-PCR determination was used to determine total bacteria and Kco_Z gene copy numbers. Degenerate primers targeting the 16S rRNA sequence of each bacterium were designed from previously sequenced 16S rRNA gene fragments, following ClustalW nucleotide alignment of the sequences [108]. Primers targeting Kco_Z gene sequences were designed from Kco_Z genomic scaffolds. Primers were checked for potential cross-hybridization
38 through BLASTn searches against the nr gene database, retrieving no significant hits (E-value =
0.1). Kco_Z primers were tested for specificity for the bacterium through PCR amplification from DNA of An. gambiae midguts fed the bacterial cocktail species. PCR fragments used for standard curves were cloned using the pGEM-T Easy vector (Promega). Standard curves for 16S rRNA and Kco_Z had efficiencies between 80 and 100% and R2 > 0.99. The relative proportion of Kco_Z was calculated by dividing the total number of Kco_Z -specific copies by the number of amplified 16S rRNA copies across three biological replicates. Amplification and detection of bacterial DNA was performed using the QuantiTect SYBR Green PCR Kit (Qiagen, Valencia,
CA, USA) and ABI Detection System ABI Prism 7000, using two technical replicates. The PCR reaction was performed in a total volume of 20 µl, which consisted of 2x Sybr Green Master Mix
(Applied Biosystems, Foster City, California, USA), 75 nM of each primer, and 50 ng of template DNA. The cycling parameters were as follows: initial denaturation at 95 °C for 2 min, followed by 40 cycles of 95 °C for 15 sec, 56 °C for 30 sec, and 60 °C for 30 sec. Following amplification, a melting curve analysis was performed from 60 to 95 °C, collecting fluorescence data every 0.5 °C.
2.3.7. Relative qPCR quantification
To conduct relative real-time qPCR assays, RNA was extracted using TRIzol® (Life
Technologies, Carlsbad, CA) according to the manufacturer’s guidelines and treated with Turbo
DNase; first-strand cDNA was produced using Superscript III reverse transcriptase (Invitrogen). cDNA templates were normalized to the Plasmodium berghei 18S rRNA gene as previously described [201], and fold changes in gene expression levels were determined using the standard
[211]. Assays were performed using Sybr Green Master Mix with the following cycling
39 parameters: initial denaturation at 95 °C for 2 min, followed by 40 cycles of 95 °C for 15 sec, 56
°C for 30 sec, and 72 °C for 30 sec.
2.3.8. Longevity, fecundity, and fertility assays
To assess the impact of bacterial introduction on mosquito longevity, bacteria were introduced into aseptic female mosquitoes, and the impact of Kco_Z was compared to that of the introduction of the bacterial cocktail or a PBS-only control. For each treatment, approximately
60 female mosquitoes were rendered free of their microbiota through antibiotic treatment, and then either Kco_Z, the bacterial cocktail, or PBS was introduced by either a blood or sugar meal.
Each cohort was then provided with a naïve blood meal at 4 days post-introduction. Non-fed mosquitoes were removed and excluded from the analysis. Mosquito mortality was monitored daily; monitoring continued until all the mosquitoes had perished, and survival percentages were calculated across three biological replicates. Kaplan-Meier survival analysis was carried out using GraphPad Prism 5 software, and p-values were determined by log-rank test (Mantel-Cox) and corrected for multiple comparisons by the Bonferroni method.
For the fecundity and fertility experiments, 30-60 female mosquitoes were treated with antibiotics as described, and three treatment groups were established with Kco_Z, the bacterial cocktail, and PBS being introduced by both sugar and blood meals. Following blood-meal introduction, blood-fed mosquitoes were removed to individual vials (12 ml) lined with moistened filter paper and then incubated under normal rearing conditions. Following sugar-meal introduction, mosquitoes were provided a naïve blood meal 48 h later, and the fed mosquitoes were removed to individual vials. Eggs oviposited on the filter paper were counted daily using a light microscope for 2 days. Females that did not produce eggs by day 2 were re-examined on day 3. After counting, eggs were submerged in water under standard larval rearing conditions,
40 and the hatch rate (fertility) was determined by counting first instar larvae, which were then removed daily. Three biological replicates were performed for the fecundity and fertility assays and significance was determined using the Kruskal-Wallis test.
2.3.9. Plasmodium infection assays
Mosquitoes were fed on NF54 P. falciparum gametocyte cultures (0.01% gametocytaemia; provided by the Johns Hopkins Malaria Institute Parasitology Core Facility) through artificial membranes at 37 °C. The adult mosquitoes were starved for 8-12 h prior to feeding to ensure engorgement, and unfed mosquitoes were removed from the cohort within 24 h. To determine oocyst numbers, the mosquitoes were incubated for a further 7 days at 27 °C, and midguts were dissected out in PBS, stained with 0.2% mercurochrome, and examined using a light-contrast microscope (Olympus). To establish whether Kco_Z is capable of inhibiting Plasmodium development when provided via sugar meal, 50-70 female septic mosquitoes were reared as described above and then provided a sugar meal containing Kco_Z at varying concentrations for
3 to 4 days; they were then provided a P. falciparum-infected blood meal the following day.
Oocyst numbers were determined as described above and compared to a cohort fed only PBS containing 3% sucrose.
2.3.10. Kco_Z genome sequencing and transcriptome analysis
Total bacterial DNA was extracted from a Kco_Z liquid culture. A 3 kb insert paired-end shotgun genomic library was prepared from the extracted DNA and sequenced using the 454 GS FLX
Titanium sequencing platform at the Genome Resource Center at the Institute for Genome
Sciences (IGS) of the University of Maryland according to the manufacturer’s protocol.
Sequencing reads were then assembled using Celera Assembler, and ORFs were predicted and
41 annotated using the IGS Annotation Engine implemented within the CLoVR-Microbe pipeline
[212]. Total bacterial RNA from bacterial liquid cultures grown in either LB or ookinete medium was extracted using TRIzol according to the manufacturer's protocol, treated with Turbo DNase
(5U, Life Technologies), cleaned using an RNeasy Mini Kit (Qiagen, Valencia, CA, USA), and the quality determined by an Agilent Bioanalyzer 2100. A custom 8x44K Agilent microarray was designed based on the 454-generated Kco_Z genome sequence. Cy-3- or Cy-5-labeled cRNA probes was synthesized from 300 ng of total RNA per replicate using the LabelIT®
µArray Dual Labeling Kit (Mirus Bio, Madison, WI, USA). Labelled RNA was hybridized overnight at 65 °C, and arrays were scanned with an Agilent scanner. Transcript abundance data were processed and analyzed as previously described [130], [213]. In brief, LOWESS normalized background-subtracted median fluorescent values were used to determine Cy5/Cy3 ratios from replicate assays and were then subjected to t-tests at a significance level of p<0.05 using MIDAS, GEPAS, and TMEV software [214], [215]. The rate for self-self hybridizations was used to calculate a cut-off value for the significance of gene regulation on these microarrays of Log2FC = 0.78 or 1.72-fold regulation [216].
2.3.11. In vitro parasite culture and co-culture with bacteria
For the rodent malaria parasite production, cultures were carried out as previously described
[108] with certain modifications. In brief, P. berghei GFP parasites were injected into donor
Swiss Webster mice and monitored daily for parasitaemia for 3 days post-infection. Once parasitaemia reached >15%, infected blood was collected by heart puncture and transferred to phenylhydrazine-treated mice. Mice were treated with pyrimethamine 72 h later and monitored daily for parasitemia and exflagellation. Parasitized mice with ≥20 exflagellation events per 20x microscope field were used for in vitro experiments. Parasitized blood was collected by heart
42 puncture and diluted 1:10 in ookinete medium (10% fetal bovine serum, 50 μg/ml hypoxanthine,
2 mg/ml NaHCO3, 1 μM xanthurenic acid in RPMI 1640 medium, pH 8.3), 4% RBC lysate, and experiment-specific constituents in a 500 μl total volume. To assess the induction of P. berghei antioxidant genes, bacteria (final concentration 106 CFU/ml) were added to the ookinete culture at 0 h. Culture samples were directly transferred to Tri-Reagent (Ambion) at various times, from
1 to 10 h after setup, for total RNA extraction. First-strand cDNA was then prepared for qPCR analysis. Development of ookinetes was confirmed by Giemsa staining at 24 h.
43 2.4. Results
2.4.1. Colonization of the Anopheles gambiae midgut by Kco_Z
Kco_Z has been detected within the midguts of field-caught anopheline malaria vectors [108], but its capacity to persist within the mosquito following artificial introduction is not known. The extent to which Kco_Z can colonize the mosquito midgut, the site of Plasmodium pre-oocyst development, is likely to contribute to the magnitude of the Plasmodium inhibition. The persistence of Kco_Z in the mosquito midgut when co-present with a simulated natural microbiota was determined, as the midgut microflora of field-caught mosquitoes has been shown to differ markedly from its laboratory-reared counterparts [188], [217]. For this purpose, we utilized a cocktail of bacteria isolated from the same southern Zambian An. arabiensis population from which Kco_Z was identified [108]. To determine the relative abundance of Kco_Z after introduction over time, the ratio of Kco_Z to total bacterial 16S rRNA was calculated through quantitative real-time PCR. Two approaches were taken to introduce Kco_Z into mosquitoes already harboring a resident microbiota: either through a blood meal as shown before for this bacterium [108] or through a sugar meal to inquire on the viability of introducing Kco_Z to anophelines via nectar feeding. Kco_Z bacteria were able to persist within the midguts of cocktail-colonized mosquitoes following blood-meal introduction for up to 72 h. Kco_Z constituted an average 32.8, 48.5, and 30.3% of the total bacterial population at 24, 48, and 72 h post-introduction, respectively. Kco_Z bacteria were detected up to 120 h post-introduction but reflected <1% of the total microbiota (Figure 2.1A, top). Following the introduction of Kco_Z through sugar feeding into mosquitoes already harboring a cocktail microbiota, Kco_Z bacteria were detected for four days, peaking at an initial 11.1% of the total microbiota following introduction and still detectable (3.5%) at day 4 (Figure 2.1A, bottom).
44 Although the bacterial cocktail consisted of strains isolated from field mosquitoes, the microflora of individual mosquitoes are highly variable [188]. Therefore, the persistence of Kco_Z for four days in the presence of natural microflora is encouraging. It also indicates that, when possible, a bacterium destined for field-based release must be investigated in the context of a natural microbiota. In addition, acquisition of a sugar meal provides energy reserves for flight [218] and promotes mosquito longevity [209], [219], suggesting that multiple sugar meals may be taken during the mosquitoes’ lifespan. Note that when introduced through the sugar meal, Kco_Z tended to constitute a lower proportion of the total microbiota when compared to ingestion by blood-feeding (Figure 2.1A). In the field, it is estimated that Anopheles mosquitoes feed on blood every two or three days [220], [221]; knowing there’s a rapid expansion of the gut microbiota upon blood intake [110], [187], it is reasonable to assume that Kco_Z, even when introduced by nectar feeding, could persist in the mosquito gut beyond what was observed in a laboratory setting. Therefore, when considering a field-based application of bacteria, sugar-meal introduction would be best evaluated in the context of the specificities of the anophelines and their feeding behavior in any given ecological niche. In summary, we have shown that Kco_Z can be introduced through both bacteria-enriched blood and sugar sources and persist for multiple days in competition with a natural microbiota.
2.4.2. Inhibition of Plasmodium sporogonic development following sugar-meal introduction
of Kco_Z
Having determined that Kco_Z can persist within the mosquito midgut for up to four days following sugar-meal introduction, the ability of Kco_Z to inhibit Plasmodium development when provided to mosquito cohorts through a sugar meal was investigated next. Previous experiments have shown Kco_Z is highly effective at inhibiting Plasmodium development at the
45 ookinete stage when provided in a blood meal [108]. Use of attractive toxic sugar baits has proven successful in reducing mosquito populations [222], indicating that bacteria, in this context, could be disseminated in mosquito populations through sugar feeding. To simulate an approximation of how Kco_Z bacteria would be disseminated through a natural population, mosquitoes, containing their endogenous microflora, were provided Kco_Z suspended in 3% sucrose and allowed to feed for three to four days. Plasmodium infection assays revealed that the impact of Kco_Z upon parasite development when provided through a sugar meal averaged a 45-
65% inhibition across the different concentrations tested (Figure 2.1B). Of note, however, is the inter-replicate variability that was observed, especially at the lowest tested concentration of 106
CFU/mL; inhibition varied from 22.2% in one replicate (from May 2014) to 90.5% in another
(February 2014). Seasonal variations in the mosquitoes’ resident microbiota may provide an explanation for these results. In particular, the midguts of mosquitoes fed in May 2014 were dominated by a different bacterium as observed by the colonies that resulted of plating ground midguts in LB agar. It is also likely that individual mosquitoes actually fed on the Kco_Z sucrose solution at different times prior to the infected blood meal, suggesting that they would harbor different Kco_Z concentrations and explaining how these results might differ from the ones obtained when Kco_Z was introduced directly with the blood meal [108]. Within 24 h after mosquito ingestion of Plasmodium gametocytes, the number of developing ookinetes is substantially lower than the initial gametocyte number [223], and this bottleneck occurs as the result of numerous factors [11]. Therefore, if detectable numbers of Kco_Z are present 24 h after an infectious blood meal, they are likely to further constrict this bottleneck and reduce
Plasmodium transmission.
2.4.3. Kco_Z influence on mosquito fitness parameters
46 It has previously been shown that the reintroduction of naturally occurring bacteria can affect mosquito fitness [111]. Recently, a Chromobacterium species has been identified that displays both anti-parasitic and entomopathogenic properties and may therefore be used to reduce both disease transmission and vector populations [142]. To provide insight into the impact of Kco_Z bacteria on An. gambiae physiology, its effects on mosquito longevity, fecundity, and fertility were assayed following introduction via the two methods: a blood meal and continuous sugar feeding. To assay longevity, Kco_Z, the bacterial cocktail, or a bacteria-free PBS control were introduced into aseptic mosquitoes and mortality was monitored daily. Following introduction via sugar-feeding, there was no significant difference between the mosquitoes harboring no bacteria (aseptic) and those continuously fed Kco_Z (p-value = 0.3852), or those harboring the bacterial cocktail and those fed on Kco_Z (p > 0.9999), all having a median survival of 16 days
(Figure 2.2A). Conversely, when introduced through the blood meal, Kco_Z had a significant negative impact on longevity when compared to the bacterial cocktail (p = 0.0004) and to the aseptic group (p = 0.0308) (Figure 2.S1A). The presence of Kco_Z, introduced by either method, did not significantly affect egg production (via sugar meal, Figure 2.2B, p = 0.0596; via blood meal, Figure 2.S1B, p = 0.0507) nor mosquito fertility measured by hatched larvae (via sugar meal, Figure 2.2C, p = 0.0741; via blood meal, Figure 2.S1C, p = 0.1000).
There was no impact of Kco_Z on mosquito survival when provided in a sugar meal (Figure
2.2A), whereas blood-feeding significantly negatively affected mosquito longevity (Figure 2.S1).
This result is perhaps expected, since acquisition of the native microflora likely occurs through sugar meals [110], [187] and not through a blood meal, which can promote a dramatic expansion of the midgut microbiota. Taken together, these data suggest that in a field-based approach,
Kco_Z introduced through sugar-baited traps would not adversely affect mosquito fitness and,
47 therefore, the presence of Kco_Z would not be selected against when artificially reintroduced into field mosquitoes.
2.4.4. Kco_Z genome analysis
To gain insight into the genetic basis of the Kco_Z-mediated inhibition of Plasmodium and develop tools for bacterial functional genomics, the Kco_Z genome was sequenced via a whole- genome shotgun sequencing approach, as described in Materials and Methods. A total of 864,207
Titanium sequencing reads were obtained, corresponding to an approximately 38x genome coverage. After assembly and ORF prediction, a total of 39 scaffolds were obtained, two of which were identified as plasmid-related through identity with known plasmid sequences in
GenBank. PCR and Sanger-based sequencing analysis confirmed the presence of two closed plasmids. The estimated size of the chromosomal Kco_Z genome is then 5.2 Mbp, with a GC- content averaging 55.8% (Figure 2.3A). The Kco_Z genome contains 4,919 putative protein- coding genes that were subjected to SEED functional analysis (Figure 2.3B) and used to construct microarrays for whole-genome expression analysis.
2.4.5. Transcriptomic survey of Kco_Z genes that mediate selection in the mosquito midgut
environment
To enhance the ability of Kco_Z to colonize the Anopheles midgut, the bacterium was serially passaged through mosquitoes and thereby selected for prolonged persistence in the mosquito midgut, as described previously [224]. The persistence of the parental Kco_Z strain was able to increase from four to eight days by selecting for bacteria able to survive in the mosquito gut for longer periods (Figure 2.4A). A total of 41 genes were differentially regulated between the
48 parental (P1) and selected (P3) Kco_Z strains, 20 genes with increased expression in the passaged strain and 21 in the parental (Figure 2.4B).
Of particular interest were a type III secretion apparatus protein, a sugar transport component, a glutathione S-transferase, and an oxidoreductase (Table 2.1). Type III secretion systems have been shown to modulate eukaryotic host immune cell signaling, allowing colonization of the host
[225], [226]. Kco_Z_3940 (Log2FC = 1.07) showed the highest similarity to Pantoea sp. Type
III secretion system export protein SpaR/YscT/HrcT, and a Pantoea Type III secretion has been shown to be important for bacterial persistence in an insect vector, the flea beetle Chaetocnema pulicaria [227]. A sugar ABC transporter component was also found in increased abundance in the passaged strain (Log2FC = 0.83). A component of the sugar transport system has previously been identified as a key virulence factor in bacterial colonization [228], and during Bacillus cereus colonization of the honeycomb moth, a sugar sensing and transport operon is upregulated
[229]. Because sugars and glucosidases are present in the mosquito midgut, it is possible that
Kco_Z adapts to its host environment through increased expression of sugar transport genes to allow the utilization of available nutrients. Following ingestion of a blood meal, the mosquito midgut environment is rich in heme, which can modulate the oxidative state of the midgut environment [230]. An increased abundance of GST (Log2FC = 0.88) and oxidoreductase
(Log2FC = 0.84) transcripts in the selected bacterium was observed, suggesting that the passaged strain is able to elevate its response to the harsh ROS rich environment in order to maintain redox homeostasis and persist within the mosquito midgut. Sugar-fed Aedes aegypti mosquitoes constitutively express ROS in the midgut, so it is possible that the selected bacterium elevates antioxidant genes in both the sugar- and blood-fed environments [231].
49 Amongst the genes with increased expression in the parental Kco_Z strain (Table 2.1), YadA transcripts (Log2FC = 1.0) are of particular note. YadA is a bacterial adhesin able to mediate binding to epithelial cells [232], [233] and may putatively align Kco_Z in close proximity to the mosquito midgut epithelium and result in elicitation of immune response [112]. Decreased YadA transcription in the passaged strain may therefore have the potential to prevent co-localization with the midgut epithelium and allow colonization without detrimental stimulation of an antibacterial immune response. UxuR, also found to have increased abundance in the parental strain (Log2FC = 0.83), is a transcription factor that negatively regulates the expression of UidA, a key enzyme of the hexuronate metabolic pathway in Escherichia coli [234], [235]. Decreased expression of uxuR in the passaged strain is therefore in line with the previous hypothesis that sugar transport may be involved in the selection process: a lower abundance of the uxuR transcriptional repressor would allow greater utilization of sugar in the mosquito midgut as a nutrient source. sulA (Log2FC = 0.90) is regulated by the SOS response, an inducible DNA repair mechanism, and halts cell division to prevent proliferation of DNA damage. The bacterial
ROS response can be activated by exposure to antimicrobials and changes in oxidative conditions [236], conditions that prevail in the mosquito midgut. This hostile environment may cause upregulation of sulA in the parental strain and, conversely, lower expression in the passaged strain, since it is tolerant to the environment.
This microarray-based transcriptome analysis has revealed that elevated expression of genes involved in response to the external environment likely contributes to the ability of the selected
Kco_Z strain to sense the local environment, and it is critical to the bacterium’s ability to more effectively colonize the mosquito midgut. In addition, decreases in the abundance of transcripts
50 involved in the ROS damage response and repression of sugar metabolism suggest that the selected strain is more suited to the hostile midgut environment.
2.4.6. Transcriptomic survey for Kco_Z-encoded Plasmodium inhibition factors
Previous experiments herein have shown that Kco_Z inhibits Plasmodium ookinete formation via a mechanism that involves bacterial production of an oxidative environment. This mechanism is likely controlled either directly or indirectly by a genetic component of the bacteria. Earlier studies also showed that Kco_Z produces factors that inhibit parasite development in vitro when cultured in an ookinete medium, but not when cultured under conditions of basic replication in
LB broth [108]. To identify putative Kco_Z genes involved in the ROS-mediated killing of
Plasmodium parasites, the same whole-genome microarray approach to compare gene expression between Kco_Z grown in ookinete and LB media was employed, since only the ookinete medium induces ROS production [108]. A total of 98 and 61 genes enriched in ookinete medium- and LB medium-grown Kco_Z were identified, respectively (Figure 2.4C). Sequences within the gene ontology groups that included oxidation-reduction process, transport, and respiratory electron transport chain were identified.
Genes of particular interest with increased abundance under ookinete medium conditions are summarized in Table 2.2. Upregulation of a transcript containing molybdenum-pterin binding domains was observed (Log2FC = 1.17) and proteins with such domains are important co-factors for the acquisition of molybdenum. This trace element functions as a catalyst for molybdenum- dependent two-electron reduction-oxidation (redox) reactions [237], indicating that upregulation of this transcript in the ookinete medium-cultured bacteria may lead to the production of elevated
ROS levels. GTP cyclohydrolase II (Log2FC = 1.38) catalyzes the first step of the biosynthesis pathway for riboflavin [238], an important precursor to bacterial redox sensors such as flavin
51 mononucleotide (FMN) and flavin adenine dinucleotide (FAD) [239]. Elevated expression of this redox sensor precursor may suggest that ookinete medium-derived Kco_Z bacteria are responding to the presence of elevated ROS levels. We also observed increased abundance of a putative NADPH nitroreductase (Log2FC = 1.36). Nitroreductases donate electrons to nitrogen- containing aromatic compounds generating ROS in the process [240], and this process has been suggested to mediate the trypanocidal activity of Nifurtimox and Benznidazole [241].
Interestingly, nitroreductases require either FMN or FAD as a co-factor for the generation of free radicals [242]: upregulation of both an NADPH nitroreductase and GTP cyclohydrolase II suggests that ookinete medium-cultured Kco_Z are generating ROS at least partially through the reduction of flavins.
In addition, a cytochrome b561, upregulated in the ookinete medium-grown Kco_Z (Log2FC =
1.11), has been shown to allow for regeneration of the reduced, ROS-detoxifying form of ascorbic acid [243]. It has previously been shown that supplementation with ascorbic acid or reduced glutathione and the resulting decreased redox environment limit the ability of Kco_Z to inhibit Plasmodium development [108]. Upregulation of a reducer of ascorbic acid suggests that the ookinete medium-derived Kco_Z bacteria are in an increased ROS environment and require an increased abundance of ascorbic acid in its reduced, active state. It becomes apparent that
Kco_Z, when grown in an environment conducive to ROS production, elevates transcripts associated with the generation of free radicals, and a combination of pathways are likely to contribute to the generation of anti-Plasmodium reactive species.
Also interesting is the finding of an upregulated 2,3-dihydro-2,3-dihydroxybenzoate dehydrogenase, a known enzyme involved in the biosynthesis of siderophores [244]. This enzyme was identified in parallel through Prediction Informatics for Secondary Metabolomes
52 (PRISM) [245] as being encoded within a gene cluster of roughly 54 kb which contains biosynthetic open reading frames indicative of nonribosomal peptide synthases. Genomic clusters moderately homologous to the one found in Kco_Z (86-87% coverage, 23-42% identity) have been associated to the production of bacillibactin and enterobactin, two well characterized nonribosomal peptides that act as siderophores [246]; BLAST [247] on the cluster’s sequence indicates Enterobacter sacchari SP1 as containing the most closely related version at 70% coverage and 84% identity. Taken together, the information seems to support that, while growing in Plasmodium-limiting conditions, Kco_Z elevates the expression of a siderophore that, albeit similar to other nonribosomal iron chelators, appears to be novel.
The interplay between the homeostasis of iron and oxidative stress has long been studied, and it is well known how iron is crucial for the generation of ROS [248], [249]. It comes at little surprise, then, to understand that Kco_Z putatively increases its siderophore-mediated iron uptake as it produces abundant ROS. Parallel consequences in Plasmodium, a parasite particularly sensitive to iron levels and for which many iron chelating drugs have been described
[250], [251], remain to be explored.
2.4.7. Influence of Kco_Z on Plasmodium antioxidant gene expression
We have previously shown that Kco_Z mediates Plasmodium killing through a ROS-dependent mechanism, as supplementation with antioxidants, such as ascorbic acid or reduced glutathione, limit the ability of Kco_Z to inhibit Plasmodium development [108]. In addition, since genes conducive to an elevated ROS environment are upregulated within this bacterium (Table 2.2), this leads to the hypothesis that the production of ROS may affect the expression of key
Plasmodium antioxidant genes. To test this possibility, in vitro ookinete cultures of P. berghei were exposed to Kco_Z, a control anti-Plasmodium bacteria that does not produce ROS
53 (Pseudomonas putida, Ppu) [108], or to a PBS control, then measured the gene expression of five parasite antioxidant genes over time: thioredoxin reductase (trxr), thioredoxin (trx), thioredoxin 1 (trx1), peroxiredoxin-1 (tpx-1), and 1-Cys peroxiredoxin (1-cys-prx). Expression of tpx-1 and 1-cys prx have previously been shown to peak at 12 and 24 h, respectively, after initiation of the ookinetes culture, and both increase in abundance in the presence of a superoxide producer; no impact on trx expression was detected [201]. Plasmodium berghei tpx-1 knockout lines have been shown to have reduced gametocyte numbers [252] and to exhibit increased numbers of immature oocysts and, consequently, a reduction in the number of sporozoites [253].
Pooling the results of two independent experiments (each including three biological replicates), we observed a reduction in transcript abundance of all selected genes 8 h after treatment with
Kco_Z (Figure 2.5). It is important to understand that arrested parasite development becomes apparent shortly after 8 h of co-culture with Kco_Z [108]. For all genes, the transcript abundance did not alter significantly when the ookinete cultures were exposed to a control bacterium
Pseudomonas putida, nor were there significant differences on antioxidant gene expression between exposure to the ROS-producing Kco_Z or this non-ROS inducing Pseudomonas putida.
Of note, also, is the fact that the expression of 1-Cys-peroxiredoxin seemed to follow a trend similar to that of the more canonical thioredoxin system enzymes, even though it has been suggested that in P. berghei 1-cys prx acts primarily to protect against heme-derived endogenous
ROS, rather than exogenous ROS [254]. Taken together, these data indicate the possibility that
Kco_Z interferes with ookinete maturation by partially and temporarily inducing a shutdown of the parasite’s antioxidant response.
Plasmodium falciparum tpx-1 knockout lines display an increased sensitivity to the presence of superoxide producers [206], and expression of P. berghei tpx-1 and 1-cys prx is increased in
54 response to external ROS [201]. These results suggest that the downregulation of thioredoxin-1 and the thioredoxin-dependent peroxiredoxins by Kco_Z renders the parasite more susceptible to the presence of Kco_Z-derived ROS [108]. P. berghei tpx-1 knockout lines demonstrate a reduced sporozoite load when compared to wild-type parasites [253], suggesting that the anti- parasite phenotype of Kco_Z may, in part, be due to a negative effect on the parasite’s antioxidant defenses.
55 2.5. Discussion
The devastating effects of malaria transmission are ever-present in sub-Saharan Africa, and increasing efforts to counteract these effects are either encountering resistance, as in the case of antimalarial drugs and insecticides, or poor efficiencies, in the case of vaccine development. It is, therefore, imperative that alternate strategies be explored, and the microbiota of disease vectors has become an attractive resource for this purpose. Not only do certain members of the vector microbiota nearly eliminate Plasmodium infection [108], [111], but some members have also been shown to effectively kill the mosquito vectors of several pathogens [142]. Here, the potential of the naturally occurring Kosakonia species Kco_Z as part of a malaria control strategy was evaluated.
Proteobacteria are highly abundant in field- and lab- mosquitoes, including Enterobacter species, showing that they can be acquired and persist in the mosquito and its habitat [188],
[190]. In order to influence the natural microbiota, bacteria could be introduced via sugar-baited traps [222] thus ensuring Plasmodium exposure to them. It was demonstrated that Kco_Z can persist for up to four days after introduction and inhibit P. falciparum development when introduced via sugar meal (Figure 2.1). As part of a translation study, it is important to examine the introduction and retention of bacteria in the adult midgut, because of the as-yet unclear data on transmission of bacteria from larvae to adults [185], [255], [256].
The impact on mosquito fitness of an introduced bacterium is an additional consideration, since a potentially negative impact is likely to be selected against in nature. Unlike the intracellular
Wolbachia species, bacteria that are able to manipulate the host’s reproductive system and thereby ensure vertical dissemination through the population [257], [258], the spread of a bacterium must not induce any fitness cost. Microbiota are important for mosquito fitness, with
56 the proteobacteria Asaia sp. influencing larval development and several other species having been shown to affect mosquito fitness [111], [259]. Naturally occurring bacterial isolates have a varying ability to persist and replicate when re-introduced into aseptic An. gambiae midguts
[111], suggesting that there is variation in the adaptation to the mosquito midgut environment.
This study has shed light on candidate molecular components that are important for bacterial colonization and persistence in the mosquito. It is possible that a combinatorial approach could be undertaken, utilizing a cocktail of anti-parasitic bacteria, each having a different anti-parasitic mechanism, for the control of malaria transmission.
In the present study, the nature of the anti-Plasmodium activity of Kco_Z was also further characterized. On the one hand, the observation that Kco_Z upregulates genes associated with the generation of ROS when cultured under conditions shown to inhibit parasite development in vitro was validated. This finding corroborates previous studies in which Kco_Z was shown to induce an oxidative environment when co-cultured with Plasmodium ookinetes, with such an environment being the likely cause of parasite inhibition. On the other hand, a potential complementary mechanism by which the bacteria appears to directly interfere with the parasite’s antioxidant system was uncovered, thereby limiting its ability to respond and adapt to the oxidative environment it promotes. This mechanism is indicated by the downregulation of key genes of the thioredoxin and glutathione antioxidant pathways in P. berghei ookinetes, until the moment when parasite development is arrested. Current efforts are underway to identify putative mediators of this effect in order to further characterize this phenomenon, in what also constitutes an exploration of Kco_Z’s metabolome from a natural product discovery perspective, in the search for new and innovative antimalarials.
57 Authors’ contributions
To this work also contributed Nathan J. Dennison, Chris M. Cirimotich, Godfree Mlambo,
Emmanuel F. Mongodin and George Dimopoulos. The candidate conceived and performed experiments towards the characterization of the anti-Plasmodium activity of this bacterium. In addition, the candidate was responsible for analyzing the entire set of data presented and was the primary contributor towards the writing of the manuscript.
This work has been published as:
N. J. Dennison*, R. G. Saraiva*, C. M. Cirimotich, G. Mlambo, E. F. Mongodin, and G.
Dimopoulos, “Functional genomic analyses of Enterobacter, Anopheles and Plasmodium reciprocal interactions that impact vector competence,” Malar. J., vol. 15, no. 1, p. 425, Dec.
2016.
* Equally contributing authors
58
CHAPTER 3. Chromobacterium spp. mediate their antimalarial
activity through secretion of the HDAC inhibitor romidepsin
59 3.1. Summary
We have previously shown that a Chromobacterium isolate, C. sp. Panama, exerts entomopathogenic activity against both adult and larval stages of malaria and dengue vector mosquitoes, along with in vivo and in vitro anti-Plasmodium and anti-dengue activities. To assess the nature of the Chromobacterium-produced anti-Plasmodium factors, chemical partition was conducted by bioassay-guided fractionation where different fractions were assayed for activity against asexual stages of P. falciparum. The isolated compounds were further partitioned by reversed-phase FPLC followed by size-exclusion chromatography; higher resolution UPLC and ESI/MS data were then collected and revealed that the most active fraction contained a depsipeptide (a cyclic peptide ester), which was identified as romidepsin. A pure sample of this
FDA-approved HDAC inhibitor allowed us to independently verify this finding, and establish that romidepsin also has potent effect against mosquito stages of the parasite’s life cycle.
Genomic comparisons between C. sp. Panama and multiple species within the Chromobacterium genus further demonstrated a correlation between presence of the gene cluster responsible for romidepsin production and effective antimalarial activity. A romidepsin-null Chromobacterium spp. mutant loses its anti-Plasmodium properties and the activity appears to be mediated by direct inhibition of P. falciparum HDAC activity. Taken together, our data indicate that romidepsin mediates the antimalarial activity of Chromobacterium spp. through a unique mode of action compared to currently used antimalarials and can be explored as a lead compound towards a novel therapeutic drug. It also substantiates the use of a chromobacteria-based approach not only towards anopheline mosquito control, but also as a malaria transmission- blocking strategy.
60 3.2. Background
Bacteria of the genus Chromobacterium are Gram-negative betaproteobacteria of the
Neisseriaceae family that occur as flagellated rods or cocci usually in water or soil environments
[260]. Originally only comprised of C. violaceum – a purple-pigmented bacterium due to the production of violacein that has been associated with opportunistic infections in humans – since
2007 there has been an expansion of the genus that now comprises more than 8 fully- characterized species [261]–[266]. Our group has recently characterized the novel
Chromobacterium sp. Panama, notable for inducing lethality in larvae and adult Aedes and
Anopheles mosquitoes, as well as in vitro and in vivo antipathogen activity against the malaria parasite and the dengue virus [142]. This makes this bacterium an interesting candidate to both control mosquito populations and pathogen transmission; manipulation of mosquito gut microbiota has been proven successful through exposing mosquitoes to bacteria-spiked artificial nectars (cf. Chapter 2).
As it concerns its anti-Plasmodium potential, C. sp. Panama was found to render Anopheles gambiae more resistant to malaria parasite infection when laboratory-reared mosquitoes were colonized by the bacterium ahead of feeding in infectious blood [142]. This antimalarial activity was proven to be mediated by bacteria-produced and secreted metabolites as in vitro assays independent of the mosquito system showed potent activity against asexual and sexual (both gametocytes and ookinetes) stages of the parasite [142].
The goal of this present study is to further develop these initial findings and characterize the antimalarial activity of chromobacteria by isolating and characterizing the secreted factor responsible for Plasmodium inhibition. For that purpose, we combine in silico, in vitro and in vivo approaches to compare the antimalarial activity of a multitude of Chromobacterium species
61 to conclude on a common compound, romidepsin, responsible for the anti-Plasmodium activity seen. The spectrum of this activity and the mechanism of action of the drug is analyzed, and potential applications of this discovery are discussed.
62 3.3. Methodology
3.3.1. Bacterial cultures and n-butanol extraction
Unless otherwise noted, Chromobacterium spp. (Table 3.1) were grown for 72 hrs. at 30 ºC in
LB Lennox broth (Sigma, L3022) without agitation, allowing for the formation of a biofilm at the surface of the culture. For n-butanol-based extraction, cultures were then thoroughly mixed
1:1 with H2O-saturated n-butanol and the top organic phase was recovered. Following a short evaporation step under reduced pressure, the mixture was run through a filter (particle retention size of 10 µm) and the filtrate was then evaporated in its entirety and the resulting residue resuspended in methanol. Subsequently, the sample was added to the same volume of 1:1 petroleum ether/diethyl ether under constant agitation, filtered (particle retention size of 1 µm) and the residue obtained resuspended in DMSO and stored at -20 ºC until further use. To ensure comparability, cultures of different chromobacteria were run in parallel and equivalent volumes of culture and solvents applied to the chemical extraction protocol.
3.3.2. Plasmodium falciparum strains and cultivation
P. falciparum strains (Table 3.2) were maintained in continuous culture according to the method described by Trager and Jensen [267]. Briefly, P. falciparum was grown in O+ red blood cells at
2% hematocrit and RPMI 1640 medium supplemented with 10 mM glutamine, 25 mM HEPES,
50 µg/ml hypoxanthine and 10% O+ human serum. In order to ensure a microaerophilic environment, the parasites were maintained in a candle jar at 37 ºC. Use of human erythrocytes to support the growth of P. falciparum was approved by the Internal Review Board of the Johns
Hopkins University Bloomberg School of Public Health.
3.3.3. Fast Performance Liquid Chromatography
63 FPLC was performed using an ÄKTA Explorer system. Reverse-phase FPLC was conducted on the n-butanol extract of C. sp. Panama (resuspended in start buffer) using a RESOURCE RPC 3 mL (GE Healthcare Life Sciences) column under gradient elution between 2% and 85% methanol in water 0.1% TFA at a constant 2 mL/min flow rate. Fraction F was collected, dried under vacuum, and the resulting residue resuspended in 20% DMSO. Size exclusion FPLC of this sample was performed on a Superdex Peptide 10/300 GL (GE Healthcare Life Sciences) column under isocratic elution with 20% DMSO in water at a constant flow rate of 1mL/min.
3.3.4. In vitro antimalarial assays
Antimalarial activity of Chromobacterium spp. bacterial culture extracts against asexual stages of P. falciparum was assessed using a SYBR green I-based fluorescence assay as described earlier [268], [269]. Different concentrations of filtered bacterial culture extracts, their fractions or pure compound in 20% DMSO were dispensed in triplicate wells of 96 well microplates, followed by addition of synchronous ring stage P. falciparum cultures at 1% hematocrit and 1% parasitemia; parasites were synchronized using 5% Sorbitol as described previously [270].
Chloroquine (250 nM) was used as positive control and 1% DMSO (i.e. final DMSO concentration) was used as negative control. After 72 hrs. of incubation in a candle jar at 37 ºC, equal volume of SYBR green-I solution in lysis buffer [Tris (20 mM; pH 7.5), EDTA (5 mM), saponin (0.008%; w/v) and Triton X-100 (0.08%; v/v)] was added to each well and mixed gently and incubated for 1-2 hrs. in the dark at room temperature. Plates were read on a fluorescence plate reader (HTS 7000, Perkin Elmer) with excitation and emission wavelengths of 485 and 535 nm, respectively. Percent inhibition was calculated relative to growth in negative (0% inhibition) and positive controls (100% inhibition). Results are shown as mean of at least 3 replicates ± standard deviation. For each treatment, significance was determined using a one-tailed one
64 sample t-test to determine if it significantly lowered parasite growth compared to control. IC50 values were estimated from a dose-response curve fitted to the normalized data using the least squares method with variable slope.
3.3.5. Mass spectrometry
Active fractions from FPLC purification were further resolved by ultra-performance liquid chromatography (UPLC) and analyzed by high-resolution electrospray mass spectrometry on a
Waters Acquity Xeno-G2 (UPLC-ESI-MS). Samples were dissolved in 20% DMSO and separated using a BEHC18 column (Waters, 130 Å, 1.7 μm, 2.1 x 50 mm) at 0.3 mL/min with a gradient elution of 20% to 80% aqueous acetonitrile + 0.1% formic acid. MS and MS/MS spectra were collected in positive ion mode.
3.3.6. Genome curation and comparisons
Genomes were curated, and mining for secondary metabolite gene clusters was performed as described by Adamek, et al. [271], with modifications. Genome sequences for nine different
Chromobacterium species/strains were obtained from the sources listed in Table 3.3. Those at contig assembly level were run through MeDuSa [272] against the sequence deposited for C. violaceum ATCC 12472 (NCBI RefSeq NC_005085.1), which was used as reference for further assembly. Pairwise genome average nucleotide identity (gANI) was calculated using the
OrthoANIu algorithm [273], and the resulting similarity matrix was used to generate an UPGMA tree using DendroUPGMA [274].
The genomic sequence obtained for C. sp. Panama was then uploaded to antiSMASH v. 4.
(bacterial) [275], [276] and the algorithm was run together with ClusterFinder for probabilistic detection of biosynthetic gene clusters [277]. The uncurated clusters obtained were run through
65 megaBLAST [278] against each of the other eight Chromobacterium genomes and a similarity index was compounded for each query by multiplying query cover with % identity. A similarity index of 90% or above indicates that said C. sp. Panama gene cluster is present in the genome of that particular species; one of 30% or below indicates with confidence that the cluster is not replicated in the other genome.
3.3.7. Romidepsin biosynthetic gene cluster analysis
The romidepsin biosynthetic gene cluster in C. sp. 968 (GenBank: EF210776.1) [279], [280] was used as reference for nucleotide sequence alignment in Geneious v5.4 (global alignment with free end gaps; cost matrix: 65% similarity) against the megaBLAST hit in each of the other
Chromobacterium spp. genomes. Prodigal [281] was run in each of these sequences to determine the boundaries of coding DNA sequences, and megaBLAST [278] was used to assign their identity based on the original annotation in the 968 strain. Average pairwise sequence identity for coding sequences between the reference and other species was determined by averaging the pairwise identity values as returned by Geneious v5.4 for each of the genes in the cluster.
3.3.8. Anopheles gambiae rearing and Plasmodium infection assays
An. gambiae Keele strain mosquitoes were maintained in the laboratory at 27 °C and 80% humidity with a 14 h day/10 h night cycle. Mosquito larvae were reared on cat food pellets and ground fish food supplement; adult mosquitoes were maintained on 10% sucrose and fed on mice anesthetized with ketamine for egg production. Female mosquitoes were infected with P. falciparum NF54 by allowing them to feed on gametocyte cultures (0.02% gametocytemia; provided by the Johns Hopkins Malaria Institute Parasitology Core Facility) through artificial membrane feeders at 37 °C. Adult mosquitoes were starved for at least 4 h prior to feeding to
66 guarantee robust feeding rates, and unfed mosquitoes were removed from the cohort after feeding. Mosquitoes were then incubated for a further 7-8 days at 27 °C and, to determine oocyst counts, midguts were dissected out in PBS, stained with 0.2% mercurochrome and examined using a light-contrast microscope.
To study the influence of Chromobacterium spp. or romidepsin on P. falciparum infection of An. gambiae, female mosquitoes were provided with roughly 105 CFU/mL bacterial suspensions in
3% sucrose or romidepsin (AOBIOUS, AOB1853) in 0.5% DMSO 3% sucrose at different concentrations, respectively. Oocyst numbers were determined as described above and compared to cohorts fed on vehicle alone; median values of a combination of at least 3 replicates are shown. Significance was determined using the Kruskal-Wallis test by comparing treatment groups to the control and correcting for multiple comparison by the Dunn’s test.
3.3.9. Chromobacterium spp. phylogenetic analysis
16S rRNA genomic sequences from the different chromobacteria (Table 3.3) in addition to C. sp.
968 (GenBank: EF210776.1) and other related bacterial species (Neisseria gonorrhoeae
NCTC13800; Neisseria meningitidis LNP21362; Pseudomonas aeruginosa DSM 50071) were aligned in MEGA6 [282] using ClustalW [283]. Evolutionary relationships between the sequences were inferred by the UPGMA method [284]; evolutionary distances were computed using the Maximum Composite Likelihood method [285] and are shown in the units of the number of base substitutions per site.
3.3.10. P. falciparum nuclear extracts and HDAC inhibition measurements
Nuclear protein extracts of P. falciparum NF54 were obtained as described before [286] following the modifications by Anne Hempel from the Manuel Llinás group (2007). Briefly, P.
67 falciparum NF54 cultures were pelleted (800g, 5 min, low brake) and red blood cell lysis was promoted in a 0.1% PBS/saponin solution; parasites were then pelleted (800g, 10 min, low brake) and lysed in 20 mM HEPES, pH 7.8, 10 mM KCl, 1 mM EDTA, 1 mM DTT, 1 mM
PMSF and 1% Triton X-100. Nuclei were harvested by centrifugation at 2,500g for 5 min and nuclear proteins were extracted for 30 min using 20 mM HEPES, pH 7.8, 800 mM KCl, 1 mM
EDTA, 1 mM DTT, 1 mM PMSF and 1x protease inhibitor cocktail (Sigma, P1860). Debris was removed by centrifugation (13,000g, 30 min) and nuclear protein extract was kept at -80 ºC in
15% glycerol until further use. HDAC inhibition by serial dilutions of romidepsin (AOBIOUS,
AOB1853) in 20% DMSO was assessed using the colorimetric EpiQuik HDAC
Activity/Inhibition Assay Kit (EpiGenTek, P-4002) according to the manufacturer’s protocol and using 10 µg of these P. falciparum nuclear protein extracts as source of Plasmodium HDAC enzymes.
68 3.4. Results
3.4.1. Anti-Plasmodium activity of Chromobacterium sp. Panama fractionates with
romidepsin
We had previously shown the in vitro inhibitory effect of the supernatant of C. sp. Panama cultures against different stages of the malaria parasite [142]. To understand the nature of this antimalarial activity we employed a bioassay-guided fractionation approach by which the presence of active compounds, against asexual stages of P. falciparum, within fractions of the C. sp. Panama supernatant was evaluated following successive rounds of chemical partition and liquid chromatography. Mass spectrometry analysis was then employed to identify compounds within fractions of interest.
First, the supernatant of a 72 h culture grown in LB medium at 30 ºC was subjected to a n- butanol-based extraction as described in the methods section. This chemical processing served to significantly retain and concentrate the desired activity against asexual stages of P. falciparum
NF54 (Figure 3.1A), as well as minimize hemolysis sometimes observed when using the untreated supernatant and that could confound the results. Given the enrichment in activity seen after n-butanol processing, we then hypothesized that the antimalarial factor was more likely to be a relatively lipophilic secondary metabolite as opposed to a protein given their general low solubility in organic solvents [287], [288], informing our next steps.
To identify possible small molecule(s) responsible for the antimalarial activity, the n-butanol extract of C. sp. Panama was subjected to reversed-phase FPLC, where the extract was partitioned using polystyrene/divinyl benzene as stationary phase and a gradient of methanol in water as mobile phase (Figure 3.1B). Fractions were collected, combined according to the peaks
69 obtained in the UV chromatogram (wavelength: 210 nm), dried and resuspended in 20% DMSO and assayed for antimalarial activity against P. falciparum asexual stages as before. Such efforts allowed to identify fraction F as the one that captured the majority of the activity (Figure 3.1C) and, as such, the fraction used for subsequent analysis.
Higher resolution UPLC separation of fraction F revealed, as expected, that the original crude fraction contained multiple components (Figure 3.S1). Three major components as judged by UV absorption (UPLC) and total ion current (MS) were isolated and designated F-I, F-II and F-III.
For F-I, and when sulfur was included as a potential element, the measured mass of 541.2151 gave a predicted elemental composition of C24H37N4O6S2 with high confidence (Figure 3.1D).
Two prominent fragment ions were visible indicating sequential losses of m/z = 117 and 83, corresponding to fragment losses of –C5H11NO2 and –C4H5NO, respectively. The observed parent mass and the unusual predicted presence of two sulfur atoms led us to a tentative assignment of the structure to FR901228, or romidepsin, a metabolite previously described from
C. sp. 968 [289]. This structure could be confirmed by unique features of the fragmentation pattern observed in its mass spectrum. Typically, peptide bonds cleave adjacent to the carbonyl to give stable acylium ion fragments. For romidepsin, however, an unusually favorable - elimination releases the valyl peptide subunit with loss of a proton followed by normal peptide scission to render loss of this amino acid fully intact (Figure 3.1D). Sequential loss then of a dehydrobutyrine (Dhb, from dehydration of a Thr residue) unit was seen further in accord with the structure assignment to romidepsin [290].
While romidepsin has a mixed polyketide and non-ribosomal peptide (NRPS) biosynthetic origin, the principal natural products in F-II and F-III had significantly greater molecular weights
(1357.6859 and 1195.6311, respectively), but proved to be identical NRPS products that differed
70 only by the presence or absence of a glucose modification. Here the interplay of mass spectrometry, genome sequence information and the availability of in silico tools to predict the identity of common amino acid building blocks [291]–[293] and their order in a NRPS product allowed us to suggest a tentative hexapeptide substructure containing a specific site of N- methylation: H2N•••Thr–Tyr–Thr–Gln–Gly–N-Me-Thr–Xxx(Leu/His/Arg)-COOH. Other fragmentary genomic data pointed to another threonine-activating domain, a putative glycosyltransferase and the presence of a specialized loading domain associated with N-terminal acylation by long-chain -hydroxyacids [294]. High-resolution mass spectrometric observation of the higher mass product, F-II, gave a prominent m/z = 1195 fragment consistent with the loss of glucose, followed by a series of fragment ions mirrored precisely in the mass spectrum of F-
III. These common fragments allowed the following residue sequence to be assigned: H2N-Thr–
Tyr–Dbh (from dehydration of Thr)–Gln–Gly–Thr–His-COOH in complete agreement with prediction (Figure 3.S2). The exact masses of F-II and F-III (± glucose) and the unique signature of this shared hexapeptide substructure dictated with high probability that F-II and F-III correspond to the previously investigated antifungal agents Sch 20562 and 20561(Figure 3.S3)
[295], [296].
To further resolve fraction F in its components, orthogonal size-exclusion FPLC (100-7000 g/mol) ran in 20% DMSO in water was employed and showed that this fraction comprises at least ten distinct entities (Figure 3.1E). These were individually collected and bioassayed for antimalarial activity against P. falciparum asexual stages as before. Potent activity was observed in fractions F6 and F7 (Figure 3.1F) and both subfractions gave mass spectrometric data fully consistent with the depsipeptide romidepsin. It is not totally clear from the mass spectrometry charts why two distinct peaks corresponding to romidepsin arose; it is possible the macrobicyclic
71 structure of romidepsin exists in two distinct structural conformations that are differentiated by a sizing resin. In fact, previous studies have observed that the oxidized and reduced forms of romidepsin elute at different retention times in reversed-phase HPLC [297]. Fractions F2 and F3, on the other hand, returned mass signatures consistent with the admixture of lipodepsipeptides
Sch 20561 and Sch 20562; no significant antimalarial activity was detected using these compounds in our in vitro assay against asexual stage Plasmodium (Figure 3.1F).
3.4.2. Antimalarial Chromobacterium spp. are closely related genetically and encode for
romidepsin production
In parallel with our bioassay-guided fractionation efforts, and in order to better understand the scope and nature of the antimalarial activity of species belonging to the Chromobacterium genus, we obtained different bacterial strains (Table 3.1) and evaluated their culture supernatants for inhibitory effects on asexual Plasmodium stages. Supernatants were subjected to n-butanol extraction as described prior to testing and, upon parallel processing of supernatants from 72 h biofilm-forming cultures, tests revealed that antimalarial activity was restricted to C. haemolyticum (MDA0585 and W10 strains) and C. sp. Panama (Figure 3.2A). Other species induced a non-significant variation in growth of Plasmodium parasites when compared to vehicle control (1% DMSO), indicating that only certain species are able to successfully express and secrete the factors responsible for parasite inhibition.
Next, we sought to understand how the strains exerting antimalarial effects compared to those that do not. Having access to genomic information for all of them (Table 3.3), we employed genome average nucleotide identity (gANI) to measure similarities and distances across the entire genomes [298], [299]. An UPGMA tree of these results indicated that C. haemolyticum and C. sp. Panama are closely related and cluster away from the remaining Chromobacterium
72 species tested (Figure 3.2B). This fact further explains why these two species, and these two alone, seem to carry significant antimalarial activity – they are somewhat unique amongst the cluster of Chromobacterium species and their genome identity significantly differs from the remaining species, indicating that they encode for specific proteins and factors not shared by the other members of the cluster.
In order to further understand the nature of the factor responsible for the antimalarial activity seen in C. haemolyticum and C. sp. Panama, the genome of the latter was mined for secondary metabolite biosynthetic gene clusters using antiSMASH [18], [19]. It was hypothesized that the antimalarial factor was a secondary metabolite as opposed to a protein, given the enrichment in activity seen after n-butanol processing as described earlier. The gene clusters retrieved were aligned to the genomic sequences of the remaining Chromobacterium spp. used in this study and a similarity index for each one was compounded (Table 3.4); a biosynthetic gene cluster present in both the genomes of C. haemolyticum and C. sp. Panama and absent from the other chromobacteria was to be considered of special relevance, as that would map with the antimalarial activity profile.
Mining of the genome of C. sp. Panama allowed for the identification of 31 sequences putatively involved in biosynthesis of secondary metabolites. Of those, 5 were revealed to fall within, or proximal to, the similarity index ranges equivalent to presence in C. haemolyticum and absence in the genomes of other Chromobacterium species (Table 3.4). Clusters 27 and 31 were discarded from further analysis given their relatively small size (less than 10 kb). Cluster 01 is assigned to encode a homoserine lactone, a known bacterial signaling molecule related to quorum sensing [300], [301]; lack of evidence implicating homoserine lactones as antipathogenic agents led us to discard this cluster from further study. Cluster 03 was extensively analyzed and
73 shown to encode the lipodepsipeptides Sch 20561 and Sch 20562, which had meanwhile been ruled out from having any relevant anti-Plasmodium effect (cf. Figure 3.1).
Cluster 05, on the other hand, significantly aligned with the biosynthetic gene cluster responsible for the production of romidepsin (Figure 3.2C), a previously described anti-tumor drug isolated from C. sp. 968 [289], [302], [303]. Alignment of the original genetic sequence of this cluster from C. sp. 968 (GenBank: EF210776.1) to those of C. sp. Panama and C. haemolyticum (both
MDA0585 and W10 strains) revealed conservation of the cluster structure with its 12 genes
(depA–depJ, depM and depR) being replicated in each sequence (Figure 3.2C); the pseudogene depN is absent from the new sequences tested [280]. Average pairwise sequence identity for coding sequences between the reference C. sp. 968 and each of the other species was of 89.6% for CSPP, 98.3% for CHAE and 98.0% for CW10.
3.4.3. Romidepsin has potent anti-Plasmodium activity
Taken together, our bioassay-guided fractionation approach and the genomic analysis of active chromobacteria strongly suggest romidepsin as the source of the antimalarial properties of these species. Romidepsin is a relatively well-studied natural product known for its potent HDAC inhibitory activity and an FDA approved cancer therapeutic against T-cell lymphoma marketed as Istodax® [304]–[306]. To follow up on our initial findings, we obtained this compound from a commercial source and proceeded to comprehensively study its effects on asexual and mosquito stages of the parasite.
As our preliminary assays focused on growth inhibition of asexual P. falciparum NF54, we started by replicating those experiments with the pure compound, allowing us to control the effective dosage being administered. As shown in Figure 3.3A, romidepsin alone exhibits
74 significant activity against this stage of the Plasmodium life cycle, with a calculated IC50 of
150.7 nM (95% CI: 114.1-198.9 nM). By falling in the nanomolar range, this IC50 further supports romidepsin as the main antimalarial effector of Chromobacterium spp.: this drug alone seems potent enough to limit parasite growth to the extent seen in previous experiments, making it unlikely other chromobacteria-produced factors are necessary to coadjuvate its activity.
Our initial findings of antimalarial activity of C. sp. Panama were seen against mosquito stages of P. falciparum [142]. To investigate whether romidepsin has activity against oocyst formation in An. gambiae, mosquitoes were fed different concentrations of this compound (vehicle: 0.5%
DMSO, 3% sucrose) for 24 h prior to ingestion of a P. falciparum NF54 gametocyte-containing blood meal. After 7-8 days post infection, the anopheline mosquitoes were dissected, and the number of oocysts in each midgut was counted (Figure 3.3B). Mosquitoes that were allowed to feed on either a 50, 200 or 1,000 µM romidepsin solution were significantly less infected than the control. It is not surprising that these concentration values differ considerably from those seen in vitro against asexual stages of P. falciparum, as these refer to the concentration of romidepsin in the solution mosquitoes fed from. It is to be expected that in a 24 h period the mosquitoes will only ingest nanoliters of the mixture, rendering the effective concentration of romidepsin available upon Plasmodium infection much lower than that of the original source; variations in infection levels seen can be explained by variations in the amount of romidepsin ingested pre-infection.
3.4.4. Romidepsin production is required for Chromobacterium spp. anti-Plasmodium
properties
Having established that chromobacteria with antimalarial properties produce romidepsin, and having demonstrated significant potency of romidepsin to inhibit growth and maturation of P.
75 falciparum in vitro and in vivo, it became essential to understand if romidepsin production was a necessary and sufficient condition for the antimalarial effect seen in supernatants of
Chromobacterium spp. cultures. For this purpose, n-butanol extracts of C. sp. 968 and a derived mutant lacking the depA gene and thus incapable of secreting romidepsin [280] were tested against asexual stages of P. falciparum as before. The inability of producing romidepsin rendered the ∆depA mutant ineffective against limiting the growth of the malaria parasite in vitro
(Figure 3.4A).
The romidepsin-null mutant was also fed to An. gambiae mosquitoes and its lethality did not differ from that of wildtype C. sp. 968 control (Figure 3.S4), indicating that romidepsin does not appear to be responsible for the adulticidal activity previously seen with C. sp. Panama against
Aedes and Anopheles mosquitoes [142]. Upon infection with P. falciparum NF54, mosquitoes that had previously fed on the wildtype or ∆depA bacteria exhibited different infection levels:
Plasmodium maturation was significantly impaired in those that ingested wildtype C. sp. 968, whereas those exposed to the romidepsin-null mutant showed no variation compared to the control (Figure 3.4B). Of note, the reduction in the number of mosquitoes between the control and the experimental groups is a result of the lethality induced by exposure to these chromobacteria independently of the mutation.
These results clearly place C. sp .968 amongst the Chromobacterium species with antimalarial activity. Commonly believed to be a strain of the C. violaceum species [279], [289], that fact would contradict our findings as described earlier (cf. Figure 3.2). However, upon phylogenetic analysis based on 16S rRNA gene sequence comparisons it becomes apparent that this 968 strain clusters with C. haemolyticum and C. sp. Panama and not with C. violaceum (Figure 3.4C). This observation is in agreement with our initial description of a cluster around C. haemolyticum that
76 exclusively possesses antimalarial activity within the genus. It also explains the lack of purple pigmentation of C. sp. 968, whose colonies are tan as those of C. haemolyticum and C. sp.
Panama, and not purple as C. violaceum and other closely related species due to violacein production. The weight of the evidence, then, points to C. sp .968 not being a C. violaceum strain and belonging to a separate cluster of chromobacteria able to produce romidepsin. The absence of a full genomic sequence for the 968 strain precludes us from making a concrete speciation assignment at this point.
3.4.5. Romidepsin is active against drug resistant isolates and inhibits P. falciparum HDAC
activity
Evaluating all the body of evidence indicating romidepsin as the antimalarial secreted by
Chromobacterium spp. and its relevant activity against different stages of the Plasmodium life cycle, it became important to investigate its mode of action. For this purpose, we decided to study the inhibitory activity of romidepsin against P. falciparum strains known for being resistant to different commonly used antimalarial drugs to understand if romidepsin is able to bypass their resistance mechanism.
In this context, we used P. falciparum strains CamWT_C580Y, a K13-propeller mutant associated with increased resistance to artemisinin; Dd2, resistant to chloroquine, pyrimethamine and mefloquine; GB4, resistant to chloroquine; and SB1-A6, resistant to inhibitors of cytochrome bc1 electron transport such as atovaquone [307]. Compared to P. falciparum NF54, which exhibited an IC50 of 150.7 nM (95% CI: 114.1-198.9 nM, Figure 3.3A), strains
CamWT_C580Y (IC50=190.1 nM, 95% CI: 114.9-249.6 nM), Dd2 (IC50=191.2 nM, 95% CI:
156.5-233.7 nM) and GB4 (IC50=156.9 nM, 95% CI: 120.1-205.0 nM) all showed overlapping intervals for IC50 values with 95% confidence (Figure 3.5A-C), indicating no significant
77 difference in activity of romidepsin against these drug resistant variants. In other words, phenotypic resistance to artemisinin, chloroquine, pyrimethanime and mefloquine does not appear to alter the inhibitory activity of romidepsin towards P. falciparum, pointing to romidepsin having a mechanism of action that differs from that of these drugs. For P. falciparum
SB1-A6, the calculated IC50 was 228.8 nM (95% CI: 214.5-244.1, Figure 3.5D), representing a
1.5 fold-change increase when compared to that of the NF54 strain. While this was a significant change, the low resistance factor of 1.5 – or 1.08 when taking the closest values in the boundaries of each of the confidence intervals – seems rather negligible and does not provide any strong evidence that the anti-Plasmodium activity is hindered when the parasite lacks requirement for electron transport through the cytochrome bc1 complex.
At the same time, given the known role of romidepsin as an HDAC inhibitor, we investigated the
P. falciparum HDAC activity in the presence and absence of the drug. Nuclear protein extracts of asexual P. falciparum NF54 were obtained as described in the methods section, and HDAC inhibition of romidepsin was evaluated by incubating the nuclear protein extract with an acetylated histone substrate in different concentrations of the drug and measuring by ELISA the ratio of remaining acetylated histone substrate in treatment vs. no-drug control after 1 h incubation. As seen in Figure 3.5E, romidepsin was able to suppress the HDAC activity of the P. falciparum nuclear protein extract from 21% at 1 nM to 80-84% at 1-100 µM. Romidepsin has been characterized as a potent inhibitor of human class I HDAC enzymes and a weak inhibitor of class II HDACs, while there is no known activity against class III enzymes [297]. P. falciparum, in turn, expresses at least 5 HDAC enzymes: one homologue of class I (PfHDAC1), two of class
II (PfHDAC2 and PfHDAC3) and two of class III (PfSir2A and PfSir2B) [308]. A maximum inhibition of ~80% of P. falciparum HDAC activity by romidepsin even at high concentrations
78 can be explained, then, by its lack of effect against the class III enzymes together with a potential partial inhibition of the class II HDACs, explaining why the deacetylating activity is never completely ablated by this drug.
In summary, our evidence points to romidepsin exerting its anti-Plasmodium effect by inhibiting
Plasmodium HDAC enzymes and, thus, interfering with the parasite’s gene expression. This is shown by the significant ability of romidepsin to interfere with the HDAC activity of a P. falciparum nuclear protein extract in our in vitro assay. There are no relevant differences in activity of this drug against P. falciparum strains resistant to common antimalarial compounds, indicating that romidepsin is not likely to share a mechanism of action with currently approved antimalarial regimens, none of which is believed to be an HDAC inhibitor.
79 3.5. Discussion
We had previously established that C. sp. Panama is able to limit the development of the malaria parasite when it colonizes the midgut of An. gambiae, and that its bacteria-free culture supernatant inhibits Plasmodium growth in vitro [142]. In this current body of work, we further probe into these observations, showing by bioassay-guided fractionation and multigenomic comparative analysis that romidepsin is the most likely Chromobacterium-produced metabolite that explains its anti-Plasmodium activity. Using romidepsin obtained from a commercial source, we show its potent inhibitory effect against asexual and mosquito stages of the parasite’s life cycle. Furthermore, we demonstrate that a romidepsin-null Chromobacterium mutant loses its anti-Plasmodium effect both against blood and mosquito stages. As comparable amounts of drug are needed to inhibit resistant variants of P. falciparum to commonly deployed antimalarial drugs, romidepsin seems to exert is activity by a distinct mechanism; in vitro measurements show its ability to limit HDAC activity of P. falciparum nuclear protein extracts, pointing to its putative mode of action as an HDAC inhibitor also against the parasite homologues of these enzymes crucial for regulation of gene expression.
As stated, romidepsin was first characterized as an inhibitor of human class I and class II HDAC enzymes and is currently FDA-approved for treatment of T cell lymphoma [304]. Our data indicates that this drug is also potentially effective as an antimalarial with an IC50 of ~150 nM against asexual P. falciparum NF54. This is in agreement with data generated against also drug- sensitive P. falciparum 3D7 of asexual IC50 in the 90-140 nM range for this drug, reported in the context of screening multiple clinically-approved HDAC inhibitors for anti-parasitic activity
[309], [310]. Previous observations showing that treatments with romidepsin lead to hyperacetylation of both histone and non-histone P. falciparum proteins, and relative inhibition
80 of recombinant PfHDAC1 [309], further reinforce the conclusion that it is the HDAC inhibitory activity of romidepsin that is likely to be behind its antimalarial effects.
As drug repurposing goes, however, romidepsin does not appear to be an ideal candidate as indicated by other groups declining from performing further tests with this drug when looking at its selectivity index [310]: the ratio between effective inhibitory dosages against mammalian and
Plasmodium targets indicates that there would be an unbearable amount of side-effects if this drug would be used in its current form [309], [310]. This fact is further reinforced by the majority of patients experiencing nausea, vomiting and anorexia, and some progressive fatigue and occasional fever, when undergoing romidepsin regimens [304]. There is, nonetheless, a great promise in considering romidepsin as a lead compound and use its chemical scaffold to develop a compound that overcomes the selectivity problems outlined by making it significantly more specific to inhibit Plasmodium HDAC enzymes when compared to their human counterparts.
While the direct use of romidepsin as a therapeutic drug against malaria cannot be advocated for at this time, the same is far from true when it comes to its transmission-blocking capabilities.
Faithful to our original approach of using Chromobacterium spp. to suppress parasite infection of the mosquito vector, uncovering romidepsin as the causal agent of the anti-Plasmodium effect further reinforces the value of this strategy. As discussed, chromobacteria are able to exert entomopathogenic activity in the malaria vector An. gambiae when colonizing their midgut through a mechanism that appears to be independent of their ability to secrete romidepsin (Figure
3.S4); control of mosquito populations remains the most widespread and perhaps valuable strategy for malaria control. For those mosquitoes that survive this colonization, however, the mechanisms for a second-line control level are now substantiated. Chromobacterium spp. will secrete romidepsin and control the parasite levels in the mosquito, leading to a potential
81 hindrance of transmission. This compound is shown for the first time to have a significant limiting effect on mosquito stages of the Plasmodium life cycle, and it appears to be doing so by a mechanism distinct from that of currently deployed antimalarials. As such, it is not expected that any pressure applied on the parasite by this HDAC inhibitor as it cycles through the mosquito will result in resistance to any antimalarial drug, placing the use of a chromobacteria- based strategy against anopheline mosquitoes as a relevant additional tool for an integrated approach to malaria control. Further studies currently underway in the semi-field with
Chromobacterium-spiked attractive sugar baits will be able to definitely conclude on the viability of this tactic.
82 Authors’ contributions
To this work also contributed Callie R. Huitt-Roehl, Abhai Tripathi, Yi-Qiang Cheng, Jürgen
Bosch, Craig A. Townsend and George Dimopoulos. The candidate conceived and performed or oversaw the conduction of all experiments. In addition, the candidate was responsible for analyzing the entire set of data presented and was the primary contributor towards the writing of the manuscript.
This work has been submitted for publishing as:
R. G. Saraiva, C. R. Huitt-Roehl, A. Tripathi, Y.-Q. Cheng, J. Bosch, C. A. Townsend and G.
Dimopoulos, “Chromobacterium spp. mediate their antimalarial activity through secretion of the
HDAC inhibitor romidepsin”.
83
CHAPTER 4. Aminopeptidase secreted by Chromobacterium sp.
Panama inhibits dengue virus infection by degrading its E protein
84 4.1. Summary
Our previous work has established that Chromobacterium sp. Panama can inhibit DENV infection of the mosquito midgut and cell lines [142]. While the mechanisms underlying this inhibition are generally unknown, the in vitro inhibition suggests it is mosquito-independent
[142], and indicate a possible production of C. sp. Panama-derived anti-DENV factors with transmission-blocking and therapeutic potential.
In this study, we aim to identify the anti-DENV factors being produced by C. sp. Panama and describe their mechanism of action. We employ a bioassay-guided fractionation approach to discover an aminopeptidase secreted by C. sp. Panama as mediating its anti-DENV activity, as it is able to degrade the viral E protein and thus prevent virus attachment to the host cell and subsequent infection.
85 4.2. Methodology
4.2.1. Cell culture maintenance, virus propagation and bacterial growth
Cell lines were maintained and viral stocks prepared as previously described [126]. BHK-21(15) hamster kidney cells were grown at 37 °C, 5% CO2, in DMEM containing 10% fetal bovine serum (FBS), 1% penicillin- streptomycin and 5 µg/ml Plasmocin. C6/36 Aedes albopictus cells were grown at 32 °C, 5% CO2, in MEM, with the same additives plus 1% non-essential amino acids. C6/36 cells were used to propagate DENV2 New Guinea C strain; after 6 dpi, the cells were lysed and supernatant clarified by centrifugation at 1,500-2,000 g for 10 minutes. The viral stock was aliquoted and stored at −80 °C for later use. Unless otherwise indicated,
Chromobacterium sp. Panama was grown in LB medium at 30 ºC for 72 hours in a shaking incubator. Culture supernatants were obtained by filtering this preparation through a 0.22 µm syringe or disk filter.
4.2.2. Dengue virus titration
Levels of DENV2 infection were assayed as before: by plaque assay in BHK-21 cells [126] or focus forming assay in C6/36 cells [311]. Both cell lines were seeded at ~8.4×105 cells/well and then incubated at 37 ºC or 32 °C overnight, respectively, before reaching 80-90% confluence.
Cells were then incubated for 1 h in the DENV2 viral stock or in a 1:1 mixture of the stock with the different treatments and covered with 0.8% methylcellulose 2% FBS DMEM medium. After
5-6 dpi BHK-21 cells were fixed and stained with 1% crystal violet in methanol/acetone (1:1).
For the focus forming assay, C6/36 cells were fixed with methanol/acetone (1:1) and blocked with 5% skim milk in PBS. Fixed C6-36 cells were incubated with mouse anti-DENV2 4G2 antibody (ATCC; 1:1,000-1:2,000) followed by goat anti-mouse IgG-HRP (1:1,500), and
86 developed with True Blue Peroxidase Substrate (KPL). In each case, the number of plaque or focus forming units (PFU, FFU) per ml was determined.
4.2.3. Solvent extractions of bacterial supernatants
For the water-immiscible solvents, C. sp. Panama supernatant was mixed with butanol, chloroform, ethyl acetate and hexanes, at 1:1 and the non-aqueous phase was recovered. For acetone, acetonitrile, ethanol, methanol and water, C. sp. Panama supernatants were desiccated in an Eppendorf Vacufuge and the resulting residue resuspended in the same volume of the different solvents for 4 hours under continuous shaking (1400 rpm). Particles that did not resuspend were then pelleted by 10 minute centrifugation at 8,000 g in a table top centrifuge and the supernatants were recovered. All extracts were then desiccated in an Eppendorf Vacufuge and the resulting residue resuspended in the same volume of water for 4 hours under continuous shaking (1400 rpm). After being filtered through a 0.22 µm syringe filter, each extract was added
1:1 to the DENV2 stock and incubated at room temperature for 45 minutes, and infectivity was assayed by plaque assay in BHK-21 cells. Desiccated LB medium resuspended in water was used as negative control.
4.2.4. Protein extract preparation
Ammonium sulfate, (NH4)2SO4, was slowly added to the C. sp. Panama supernatant until reaching 70% saturation. After rapid stirring at 4 °C for 1 hour, proteins were pelleted down by centrifugation at 10,000 g for 20 minutes. The protein pellet was then gently resuspended in 0.1
M Tris-HCl pH 7.2 buffer and concentrated using an ultra-centrifugal filter unit (nominal molecular weight limit, 30 kDa; Amicon). For comparison, the ammonium sulfate-rich supernatant was desalted and concentrated to the same extent.
87 4.2.5. Pre- and post-virus attachment neutralization assays
The following protocols were adapted from the literature [312]. Pre-attachment: BHK-21 cells and reagents were cooled to 4 °C. Virus and C. sp. Panama extract were incubated at 4 °C for 1 hour, then transferred to chilled BHK-21 cells and incubated for another hour at 4 °C. After incubation, cell monolayers were washed 3× with cold PBS, overlay medium was added and cells were incubated at 37 °C for 5 days before fixing and staining for plaque detection.
Post-attachment: BHK-21 cells and reagents were cooled to 4 °C. Virus was added to cells and allowed to attach for 1 hour at 4 °C. Unbound virus was washed off by washing twice with cold
PBS. C. sp. Panama extract was then added to virus bound cells and incubated for another hour at 4 °C. Following incubation, cell monolayers were washed once with cold PBS. Overlay medium was added and cells were incubated at 37 °C for 5 days before fixing and staining for plaque forming unit.
4.2.6. Transmission electron microscopy
DENV2 viral stocks were purified as described previously [313]. Pure DENV2 viral particles suspended in 1:1 v/v 0.1 M Tris-HCl (pH 7.2) or in the C. sp. Panama proteinous extract were incubated at room temperature for 1 h and virus within the samples was sequentially inactivated in 4% paraformaldehyde in PBS for 30 min at 25 ºC followed by 30 min at 37 ºC. The samples in
0.08 M phosphate were then applied to a carbon grid, washed 3 times with 50 mM TBS and negatively stained with 1% w/v uranyl acetate 0.04% w/v tyloindicines. The grids were allowed to air-dry and images were acquired in a Hitachi 7600 TEM microscope at 80.0 kV.
4.2.7. Protein gels and Western blotting
88 Viral stocks were incubated 1:1 v/v with the differentially treated C. sp. Panama culture supernatants, their protein extracts or control buffers for 1 h at room temperature. For heat inactivation, the protein extract was incubated at 90 ºC for 1 hour prior to being added to the virus suspension. Samples were then incubated at 85 ºC for 2 min in SDS-containing loading buffer and ran on stacked 4-20% tris-glycine gels under denaturing conditions. Protein bands were dry-transferred to a nitrocellulose membrane, followed by overnight blocking at 4°C in 5% skim milk 0.1% Tween 20 PBS. Membranes were probed with anti-DENV2-E rabbit polyclonal
(1:2,000) followed by anti-rabbit HRP-linked donkey (1:20,000, Amerhsam) antibodies for 1 h each, with abundant wash with 0.1% Tween 20 PBS after each probing. Chemiluminescence signals were detected using Amersham ECL Prime Western Blotting detection reagents.
4.2.8. Immunofluorescence microscopy
Immunostaining and fluorescence microscopy was conducted as described before [314], with adaptations. After 72 h growth at 37 °C, BHK-21 cells were infected with DENV2 (MOI = 10) in the presence and absence of C. sp. Panama proteins for 1 h at 4 °C. Cells were washed with PBS and fixed with 4% paraformaldehyde for 1 h at 4 °C. Fixed cells were blocked with 0.5% BSA in
PBS for 1 h at room temperature, washed with PBS and incubated with mouse 4G2 anti-DENV2 antibody (ATCC) for 2 h at room temperature. Alexa Fluor 568 goat anti-mouse IgG (Invitrogen) was used as secondary antibody. Beta-actin was stained with Alexa Fluor 488 Phalloidin
(Invitrogen) and nuclei were stained with DAPI. Samples were mounted with ProLong Gold antifade reagent and visualized on a Leica DM 2500 fluorescence microscope.
4.2.9. Hydrophobic interaction chromatography
89 Fast protein liquid chromatography was performed using an ÄKTA Explorer system.
Hydrophobic interaction FPLC was conducted on protein extracts of C. sp. Panama resuspended using HiTrap Butyl FF or Butyl HP columns (GE Healthcare Life Sciences) under gradient elution from 1.5 to 0 M ammonium sulfate in 50 mM sodium phosphate (pH 7.0) at constant 1 mL/min flow rate. Collected fractions were dialyzed using ultra-centrifugal filter units (nominal molecular weight limit, 30 kDa; Amicon) for solvent replacement as needed.
4.2.10. Proteomics and protein sequence analysis
Fraction H2p in 0.1M Tris-HCl was submitted to the Johns Hopkins Medicine Mass
Spectrometry Core for identification using iTRAQ as before [315]. Peptides obtained were searched against the theoretical proteomes of Chromobacterium spp. following trypsin digestion using Mascot (Matrix Science); Scaffold 4.8 (Proteome Softwares, Inc.) was used to validate protein identification at 99.0% protein probability threshold, 4 as the minimum number of peptides and 95% as the peptide probability threshold. Amino acid sequences of identified peptides and proteins were further searched against the C. sp. Panama theoretical proteome based on its genomic information for validation of the findings. Sequences of proteins of interest were subjected to basic local alignment tools (BLAST) against the UniProt database and further aligned to the closest related annotated entry using the Geneious v5.4 algorithm (Biomatters
Limited).
4.2.11. Protease inhibiton
Bestatin and phosphoramidon (Sigma) were diluted in water and supplemented to LB at the indicated concentrations prior to inoculation of C. sp. Panama. Following 72 h incubation at 30
90 ºC, bacterial culture supernatants were collected and either processed as before for Western blotting or evaluation of effect on DENV2 titer by plaque assay on BHK-21 cells.
91 4.3. Results
4.3.1. C. sp. Panama culture supernatant shows potent anti-DENV activity
In our previous work, Ramirez & Short et al. [142] explored the in vitro anti-DENV activity of
C. sp. Panama by incubating unfiltered bacterial cultures with DENV2 particles, removing the bacterial cells only prior to assessing infection in hamster or mosquito cells. To understand if live bacteria are required to impair DENV infection, C. sp. Panama cultures were first filtered through a 0.22 µm membrane and only then mixed with the DENV2 viral stock. DENV replication in both BHK-21 cells (mammalian) and C6/36 cells (mosquito) was reduced by 4-5 logs (BHK21: p < 0.0001, C6/36: p = 0.0009) upon treatment with this sterile supernatant when compared to the control (Figure 4.1A). This indicates that C. sp. Panama does not exert any direct effect on the viral particles, but does so through secreted or otherwise released mediators.
Similar level of C. sp. Panama-induced anti-DENV2 activity in both cell lines indicates that the target of such bacteria-derived factors are likely to be viral proteins or RNA, or host factors conserved across species [316].
4.3.2. Anti-DENV activity is associated with C. sp. Panama-produced proteinous factor(s)
To understand the nature of the C. sp. Panama produced factors that influence DENV2 replication, we first partitioned the C. sp. Panama supernatant with commonly used solvents, both miscible and immiscible in water. C. sp. Panama molecules soluble in acetone, acetonitrile, butanol, chloroform, ethanol, ethyl acetate and hexanes showed no anti-DENV activity (Figure
4.1B). Even though methanol partially extracted molecules with anti-DENV properties (< 1-log reduction in DENV2 titer compared to control, p = 0.002), water was the only solvent that was able to retain a significant majority of the activity of the initial extract (4-log reduction, p =
92 0.0002), providing early evidence that the relevant C. sp. Panama factors are likely charged or otherwise polar biomolecules, such as hydrophilic proteins or sugars, and not lipidic in nature.
Next, we assessed the thermostability of such factors, concluding that 1 hour incubation at 50 ºC partially inactivated (2-log reduction in DENV2 titer compared to control, p = 0.0005) the level of anti-DENV activity seen when samples were handled at room temperature (as before) or 37
ºC, for which no viral replication was detected (Figure 4.1C). Incubation at 70 ºC or 99 ºC for 1 hour resulted in total loss of activity (Figure 4.1C). Taken together, these data strongly make the case for a hydrophilic protein being the mediator of the C. sp. Panama-induced anti-DENV effects, given the very strong preference for water as solvent, the only that would ensure proper protein folding [287], [288], and the dramatic loss of activity at 50 ºC and above, a pattern not to be as readily expected from sugars or other small biomolecules.
To further test our theory, we used ammonium sulfate precipitation [317] to create whole-protein extracts of the C. sp. Panama supernatant. The resulting protein pellet carried all the anti-DENV activity (4-log reduction in DENV2 titer compared to control, p < 0.0001, Figure 4.1D), whereas the supernatant, after appropriate desalting, was ineffective in preventing DENV2 infection and replication. This result clearly indicates that proteinous factors, such as peptides, proteins or protein complexes, are the likely mediators of the anti-DENV activity of C. sp. Panama.
4.3.3. C. sp. Panama supernatant proteins inhibit DENV attachment to host cells
Attachment of viral particles to host cells is the first step that initiates virus entry, and thus precedes all the following steps necessary to viral replication. To understand if our C. sp.
Panama proteinous extract exerts its anti-DENV effect by interfering with viral attachment, we performed neutralization assays pre- and post-attachment using low temperature to control
93 receptor-mediated endocytosis. For our pre-attachment assay, we first incubated DENV with C. sp. Panama proteins at room temperature and then exposed this mixture to pre-chilled cells at 4
ºC to allow for viral attachment without endocytosis. Unbound virions were washed off, and attached virions were allowed to complete infection. In our post-attachment assay, DENV virions were first exposed to pre-chilled cells at 4 °C and, only after extensive wash, C. sp. Panama proteins were added to treat attached virions. Our results showed a significant 3-log decrease in
DENV2 titers compared to control (p < 0.0001) when DENV was exposed to the C. sp. Panama proteins before attachment (Figure 4.2A). On the contrary, C. sp. Panama proteins failed to inhibit DENV replication when attachment had already occurred (Figure 4.2A), indicating that our active factor(s) is likely to either interfere with the viral particle itself and/or prevent its attachment to host cells, and not exert effect later in the viral replication cycle.
4.3.4. C. sp. Panama interferes with DENV attachment by mediating degradation of the E
protein
To investigate if C. sp. Panama proteins affect viral particles, we incubated the protein extract with purified DENV2 virions and processed the samples for transmission electron microscopy
(Figure 4.2B). The dengue virus envelope has a diameter of 50-60 nm [318], [319] and, fully consistent with that, we obtained a measurement of 56.7±1.1 nm for the size of the viral particles in our control group (Figure 4.2C). Upon treatment with C. sp. Panama proteins, however, the viral particles exhibited a significantly different (p < 0.0001) average size of 34.0±3.2 nm
(Figure 4.2C). The E protein of dengue virus occupies from radius 180-280 Å (i.e. 18-28 nm) of the particle, as shown by analysis of a class of virus with loose or absent E protein with diameter of 36 nm [319]. As such, we hypothesized that treatment with our bacterial protein extract compromised the integrity of the E protein coating layer of the dengue virus particle. This is in
94 agreement with our previous finding that viral attachment is compromised, as the E protein is a crucial mediator of viral attachment and fusion [320], [321].
To test this hypothesis, we exposed DENV2 to the C. sp. Panama protein extract as before and probed the resulting proteins for the E protein with a polyclonal antibody by Western blotting.
As seen in Figure 4.2D, a significantly reduced signal for the E protein is detected at the expected size of 53 kDa [321], the same not being true for when the virus is incubated with a heat-inactivated C. sp. Panama protein sample. Our immunofluorescence microscopy results were somewhat consistent with this finding by failing to detect attached virions to BHK-21 cells with another anti-E protein antibody (4G2, ATCC; Figure 4.2E). However, as the cells were washed prior to staining, it is possible to interpret that any remaining bound virus (as hinted in
Figure 4.2A) was simply below the detection limit for this technique. Nonetheless, there is clear evidence that there is severe E protein degradation upon treatment with C. sp. Panama, pointing to a potential protease secreted by this bacterium as the most likely factor responsible for this effect.
4.3.5. Secreted proteases are present in active anti-DENV protein fraction
Independently from these efforts, we also sought to further partition the C. sp. Panama culture supernatant protein extract in search of the causative agent for the anti-DENV activity. For this purpose, we employed hydrophobic interaction chromatography (HIC) to separate proteins based on their content in hydrophobic amino acids. Following an initial fractionation in butyl
Sepharose under a salt gradient, we were able to obtain a discrete fraction (H2, Figure 4.3A) that preserved its anti-DENV properties. This fraction was further purified by a second round of fractionation where conductivity along the gradient was monitored to match ~60 mS/cm, resulting in collection H2p (Figure 4.3B). SDS-PAGE gels of these fractions revealed that H2p is
95 considerably less diverse than H2, with the exception of a band at approximately 30 kDa and other small proteins or peptides less than 16 kDa in size (Figure 4.3C). Most importantly, this fraction retained proteolytic activity, as seen in Figure 4.3C by its ability to degrade DENV proteins, including the E protein whose signal is presumed to be corresponding to the large band at ~53 kDa.
Satisfied with the low complexity of the H2p fraction, we next chose to perform a proteomic analysis by mass spectrometry to understand which C. sp. Panama proteins were contained within this sample and the results are summarized on Table 4.1. Of the 8 detected proteins, 2 were of particular relevance given their assignment as proteolytic, CSPP0261 and CSPP0262. As hinted by the numbering, the neutral protease and the aminopeptidase are encoded in tandem in the C. sp. Panama genome and both are assigned to be secreted according to the most closely related UniProt reviewed entries, P14756 from Pseudomonas aeruginosa for CSPP0261 and
Q01693 from Vibrio proteolyticus for CSPP0262, making them ideal candidates for subsequent studies.
4.3.6. Aminopeptidase secreted by C. sp. Panama mediates its anti-DENV activity
Having putatively narrowed down the anti-DENV activity of the C. sp. Panama culture supernatant to two proteases, we aimed at a better understanding of these two enzymes by comparing their sequence to similar proteins already discussed in the literature. For the neutral protease CSPP0261, BLAST analysis returned significant similarity to pseudolysin (59.3% pairwise identity) from P. aeruginosa and vibriolysin (49.7% pairwise identity) from V. proteolyticus, two secreted zinc metallopeptidases of the M4 family that form a divergent branch within the clan [322]. They have been described as having a nutritional role by scavenging amino acids for the bacteria, as well as a virulence role in pathogenic species either by directly
96 mediating the destruction of tissue or indirectly by interfering with host defense mechanisms or the normal function of host proteases [323]. Sequence alignment showed that CSPP0261 conserves the active and metal binding sites of these enzymes (Figure 4.4A), and that it lacks the secondary C-terminal catalytic domain of vibriolysin [324].
For the aminopeptidase CSPP_0262, the only matching fully reviewed entry in UniProt corresponded to aminopeptidase Ap1 (39.6% pairwise identity) from V. proteolyticus, also a secreted zinc metallopeptidase but belonging to the M28 family as it binds two zinc ions that co- catalyze the proteolytic reaction [325]. As the name hints, this enzyme is able to remove free N- terminal amino acids of peptides or proteins that are hydrophobic and large (preferably), basic or proline, but never aspartyl, glutamyl or cysteic acid residues; this capability has been harvested in biotechnological settings namely in the production of antiviral and anticancer agent interferon alfa-2b [325], [326]. Also for CSPP0262, sequence alignment showed perfect agreement in all the annotated active metal binding sites for the V. proteolyticus version (Figure 4.4B);
CSPP0262 lacks the C-terminal propeptide present in the Vibrio aminopeptidase for which, however, no function has been assigned [327], [328].
Of note, both proteases contain a conserved signal peptide sequence potentially also governing their secretion by the Chromobacterium species. Also conserved is an N-terminal propeptide that has been shown for both ortholog proteins to be crucial for correct folding of the mature protein, having the ability to inhibit its activity until secretion to prevent intracellular damage [323],
[327], [328]. This N-terminal propeptide is autocatalytically cleaved in pseudolysin/vibriolysin
[323], [324] and also in the Vibrio aminopeptidase [329], even though for the latter a role has been proposed for vibriolysin as the biologically relevant activating peptidase [330]. Vibriolysin and the Vibrio aminopeptidase are co-expressed [330], as also appear to be CSPP0261 and
97 CSPP0262, which further validates this observation. P. aeruginosa also secretes an aminopeptidase belonging to the M28 family that is processed to its mature form by pseudolysin
[331], but BLAST analysis revealed weak identity between this aminopeptidase and CSPP0262
(not shown).
After processing, both enzymes would assume an active form of ~32 kDa, consistent with our observation that our proteolytically active H2p fraction is enriched with a protein of this size
(Figure 4.3C). Different between the two peptidase families, however, is their susceptibility to inhibitors: phosphoramidon is known to inhibit the activity of the M4 proteases [332] whereas bestatin has been described as an inhibitor of the M28 peptidase [325]. In an attempt to determine which of these enzymes is responsible for the anti-DENV activity of C. sp. Panama, we grew the bacteria in media supplemented with each of these inhibitors and evaluated the culture supernatant for its ability to degrade the DENV2 E protein as described above.
As shown in Figure 4.4C, treatment with 10 µM bestatin, and not 100 µM phosphoramidon, was able to revert E protein degradation by the C. sp. Panama culture supernatant. This indicates that the activity of the CSPP0262 aminopeptidase is crucial for digestion of the E protein, with minimal contribution of CSPP0261. These results are replicated when looking at the effect of these inhibitors in DENV titers by plaque assay with total ablation of anti-DENV activity in the presence of bestatin (Figure 4.4D). However, a more prominent role can be postulated for
CSPP0261: even though supplementation of bacterial growth media with phosphoramidon is not able to fully counteract the inhibitory activity of the C. sp. Panama culture supernatant, it is apparent that this inhibitor indeed aids in rescuing the phenotype. These findings are compatible with the observation made in V. proteolyticus that the aminopeptidase is able to be activated by autocatalysis, even though that route lacks in efficiency when compared to the activation by the
98 respective neutral protease [329]. The bulk of the evidence, then, points to the CSPP0262 aminopeptidase as the causative agent of the anti-DENV activity by C. sp. Panama, and supports a facultative role of the CSPP0261 neutral protease in the activation of this peptidase.
99 4.4. Discussion
The anti-DENV activity of C. sp. Panama was further investigated in the present work, building on the initial observation that this bacteria can inhibit viral replication in vivo and in vitro [142].
First, we validated that the activity is mediated by a secreted factor and does not require live bacteria; retention of activity by protein extracts of the culture supernatant further indicated that a protein was responsible for the effect. Then, we demonstrate that the C. sp. Panama protein extract interfered with viral attachment by promoting the degradation of its E protein. Successive fractionation of this extract allowed for isolation and identification of two tandemly expressed proteases, CSPP0261 and CSPP0262, that appear to function together towards the anti-DENV activity of C. sp. Panama, as indicated by experiments conducted with specific inhibitors to each of the peptidases.
This is the first study characterizing the mechanisms of the anti-DENV activity of a bacterium that was originally isolated from the Aedes mosquito midgut. Knowing that the midgut environment of the mosquito is highly proteolytic, particularly when blood digestion is occurring
(cf. Chapter 1), it is interesting to conclude that a bacterially produced protease, and not any other, is potentially able to significantly impact DENV infection in a timely fashion. Of course, this fact still requires final validation currently underway: we are actively investigating if all our observations translate to our in vivo Aedes-DENV model, as well as the specificity of this activity against the DENV E protein to understand putative therapeutic applications of our findings. So far, we have preliminarily observed that C. sp. Panama is indeed also able to limit
ZIKV infection in vitro and that E protein degradation is also involved and selectively rescued by bestatin. Given the similarities between the two flaviviruses this fact does not provide any evidence that the activity is nonspecific and thus prone to secondary effects; in fact, it is rather
100 encouraging and broadens the putative applicability of the discovery of this aminopeptidase in the therapeutic and transmission-blocking arenas.
101 4.5. Authors’ contributions
To this work also contributed Jingru Fang, Seokyoung Kang, Yesseinia I. Angleró-Rodríguez,
Yuemei Dong and George Dimopoulos. The candidate conceived and performed experiments towards the identification and characterization of the anti-DENV factor of this bacterium. In addition, the candidate was responsible for analyzing the entire set of data presented and was the primary contributor towards the writing of the manuscript.
This work has been submitted for publishing as:
R. G. Saraiva*, J. Fang*, S. Kang, Y. I. Angleró-Rodríguez, Y. Dong and G. Dimopoulos,
“Aminopeptidase secreted by Chromobacterium sp. Panama inhibits dengue virus infection by degrading its E protein”.
* Equally contributing authors
102
CHAPTER 5. Concluding Remarks
103 In this work, we set out to characterize molecular mechanisms through which individual natural mosquito midgut-associated bacteria can limit pathogens within the mosquito vector. Through the study of K. cowanii and C. sp. Panama and their interactions with Plasmodium – An. gambiae for both bacteria, and DENV – Ae. aegypti for the latter, we were able to unfold how these bacteria influence pathogen load in the insect.
Chapter 2 details how Kco_Z is able to partially and temporarily induce a shutdown of the antioxidant response of the motile Plasmodium ookinete in addition to stimulating the production of ROS. The combination of these factors is likely to result in the arrest of parasite development seen both in vitro and in vivo. This effect comes at no fitness cost to the vector, which shows comparable parameters when colonized with this bacterium.
In Chapter 3, romidepsin is presented as the mediator of the anti-Plasmodium activity of
Chromobacterium spp., as shown by bioassay-guided fractionation of C. sp. Panama culture supernatants and comparative genomic analyses across the type strains of chromobacteria. Tests ran on a romidepsin-null mutant further confirm this hypothesis, as activity is lost when the bacterium is unable to produce and secrete this compound. Validation with the romidepsin compound alone shows that it is able to inhibit P. falciparum NF54 in vitro and in vivo within
Ae. gambiae. Surveying for romidepsin’s effect on drug resistant P. falciparum isolates revealed that this compound is able to bypass resistance mechanisms that arose against commonly deployed antimalarials. Further probing indicates that romidepsin also acts as an HDAC inhibitor against P. falciparum, a mechanism of action that is not proposed to any of the currently approved drugs.
The results of the evaluation of the anti-DENV activity of C. sp. Panama are described in
Chapter 4. It is conclusively demonstrated that the anti-DENV activity, as is true for the anti-
104 Plasmodium effect, is mediated by secreted factors that directly interfere with the virus, inhibiting its attachment to host cells. The envelope protein of DENV appears to be targeted, as seen by transmission electron microscopy images of viral particles and confirmed by western blotting. Bioassay-guided fractionation coupled with mass spectrometry suggests a role for an aminopeptidase secreted by C. sp. Panama mediating degradation of the viral E protein with the help of an also secreted neutral protease. The absence of functional E protein prevents viral infection before the DENV has a chance to replicate, this severely impacting viral titers.
These three bacteria-mediated anti-pathogen mechanisms, despite the obvious differences already discussed in detail, share some commonalities. In all cases, the bacteria studied seem to act on the pathogen directly and not require mosquito factors to mediate their effect. To our knowledge, these are the first descriptions with mechanistic detail of such processes and present compelling evidence that the mosquito gut microbiota, besides their role as immune primers and as competitors for resources, can indeed interfere directly on pathogen load.
A case can be made for a symbiotic relationship between Kco_Z and its anopheline host when we consider that Enterobacteriacae are frequently isolated from the gut of mosquitoes and this isolate does not cause fitness costs for its host. The oxidative environment elicited by this bacterium, thus, can be postulated to be of benefit to the mosquito: it prevents Plasmodium invasion and refrains the insect from having to share resources with the parasite during its two- week long extrinsic incubation period. However, evidence for a mutualistic association is lacking: Kco_Z does not readily colonize the mosquito gut and only after serial passages were we able to obtain an isolate that could persist for longer than one week.
For the chromobacteria, the same argument does not hold. It is clear that these bacteria induce entomopathogenic effects and so it is not expected that either romidepsin or the two-protease
105 system described has evolved to the mosquito’s advantage or specifically against the malaria parasite or the dengue virus. The validity of the findings from a natural product discovery point of view cannot be discarded, and propose exciting opportunities to harvest this new information towards development of novel strategies for control of disease transmission.
One would be to develop the compounds or proteases described for therapeutic or transmission- blocking purposes independent of its secreting bacteria. Antimalarial drugs are suffering at the hands of an increasing incidence of drug resistance and there are no therapeutic regimens approved against DENV. Albeit challenging, one can see the path where the chemical scaffold of romidepsin would be used to design a semi-synthetic drug with increase tropism to the
Plasmodium HDACs and not their human counterparts. For the anti-DENV aminopeptidase from
C. sp. Panama, one potential application could be the genetic engineering of mosquitos to be able to produce and secrete this enzyme into their midgut lumen upon blood meal ingestion, thus putatively inactivating the virus before it has a chance to invade the midgut epithelial cells.
Far from saying these two approaches would not be daring within themselves, they would be mostly using currently available technology that has been in development for quite some time now: the former, lead optimization, with obvious success and the latter, mosquito genetic engineering, with proven results in laboratorial setting but still lacking a tested pathway to applicability. A truly novel strategy, still in need of much conceptual work before attempting deployment, would be to devise ways of using the live bacteria as is and think of ways to promote a shift on the mosquito microbiota of a given mosquito population towards enrichment in these putative disease-protective members.
To that extent, some preparative work has already been done with Kco_Z as shown in Chapter 2.
By serially passaging this bacterium through the mosquito we were able not only to improve its
106 colonization rate and persistence time, but also to identify genes that are putatively involved in the process that can be further explored towards optimizing these two variables. We can now conceive of creating attractive sugar bait stations spiked with this bacterium and evaluate their effectiveness outside of the laboratorial setting and into the semi-field for validation, and those experiments are underway. Through this strategy, we would mimic the use of probiotics in humans that have been proven successful in replacing flora and resolve maladies related to dysbiosis, but apply it to mosquitoes and understand if a similar approach is viable in nature.
For C. sp. Panama, its insecticidal properties force us to probably approach it differently. One can conceive following the same protocol and serially adapt chromobacteria to the mosquito gut, thereby selecting against said entomopathogenic effects while retaining the antipathogenic ones, but that would involve stripping the bacterial agent from an activity that we are very keen on preserving: that of limiting the mosquito’s lifespan and, thus, also limit the chances of pathogen transmission. On an ideal scenario, chromobacteria would act as a two-tiered control agent – it would promote death of the vector while also preventing it from acquiring and transmitting pathogens. This would simulate a combination approach, where different processes are targeted at the same time maximizing the chances of success.
For that purpose, the fact that romidepsin limits Plasmodium development through a mechanism putatively different from that which mediates Chromobacterium spp.’s insecticidal activity is of particular interest. Also of interest is the fact that romidepsin acts on a target that remains unexplored by approved antimalarials, meaning that this approach would not apply pressure on the parasite to acquire traits that would contribute to its resistance against drugs in use to control disease in humans. While it is not unconceivable DENV could acquire mutations in its E protein that renders the activity of the aminopeptidase from C. sp. Panama ineffective, we can
107 hypothesize that that would come at a high fitness cost for the virus given the high specialization of this envelope protein and its sequential role in both attachment and entry.
Many more considerations are probably needed before any of the information portrayed here can be translated into applications. Until then, we have uncovered previously unexplored processes by which individual members of the mosquito microbiota can inhibit two major pathogens of relevance for human disease, malaria and dengue, and are willing and eager to see this new knowledge propel the field further towards the end goal of elimination of these tropical diseases.
108
TABLES
109 Table 1.1. Examples of compounds with antimalarial activity isolated from bacteria. SI, selectivity index; calculated as the ratio between activity against mammalian cell line and against
P. falciparum.
IC50 P. PubChem Name Source falciparum Cytotoxicity SI Notes References CID / µM
2-undecen-1’- Pseudomonas 17 µM, IC CAS: 678176- 11 50 2 [333] yl-4-quinolone sp. 1531 E7 KB cells 79-9
>33 µM, 2-undecyl-4- Pseudomonas 10063343 3 IC KB >11 [333] quinolone sp. 1531 E7 50 cells 853.56 µM, Streptomyces [334], acrylamidine 3481477 167.49 IC MRC-5 5 eurythermits 50 [335] cells >100 µM, actinoramide Streptomyces 53483403 0.19±0.02 IC HepG2 >476 [336] A bangulaensis 50 cells other Microcystis aerucyclamide 120 µM, aerucyclamides 24970831 aeruginosa 0.7 171 [337] B IC L6 cells in the same PCC 7806 50 paper 0.027 µM, Streptomyces [164], alborixin 73359 0.0007 EC U937 39 albus 50 [338] cells
Fischerella 68 µM, MIC ambigol A 475341 1.74 39 [339] ambigua L6 cells
206 µM, Fischerella ambigol C 5276614 3.17 MIC L6 65 [339] ambigua cells Streptomyces 1.61 µM, [335], amidinomycin 160703 flavochromog 0.857 IC MRC-5 2 50 [340] enes cells 64.3% Bacillus sp. growth of bacilosarcin A 101439757 2.2 ND [341] GL0033 MCF-7 cells at 20 µM 0.32±0.06, bafilomycin Streptomyces 0.066±0.01 6436223 IC BC-1 3-8 [342] A1 spectabilis 6 50 cells Microcystis balgacyclamid >150 µM, 86579110 aeruguinosa 9.0 >17 [343] e A IC L6 cells EAWAG 251 50
110 Microcystis balgacyclamid >150 µM, 86579111 aeruguinosa 8.2 >18 [343] e B IC L6 cells EAWAG 251 50
Okeania 2.1 µM, IC bastimolide A 122232645 0.140 50 15 [344] hirsuta Vero cells
40±9 nM, 0.058±0.00 calothrixin A 9926867 Calothrix sp. IC HeLa <1 [345] 8 50 cells 350±82 nM, 0.180±0.04 calothrixin B 9817721 Calothrix sp. IC HeLa 1-3 [345] 4 50 cells
Lyngbya 9.8 µM, IC carmabin A 16757536 4.3 50 2 [346] majuscula Vero cells
Streptomyces 0.00019 µM, concanamycin [347]– 6438151 diastatochrom 0.0002 IC HeLa 1 A 50 [349] ogenes S-45 cells Streptomyces 10.93 µM, cyclamidomyc [335], 5368895 sp. MA 130- 8.39 IC MRC-5 1 in 50 [350] A1 cells 3244±11 dibutyl Actinomycete µM, IC 147- 3026 17.5±4.5 50 [351] phthalate sp. H11809 Chang liver 250 cells 182.3 µM, Lyngbya dragomabin 16737469 6.0 IC Vero 30 [346] majuscula 50 cells 67.8 µM, dragonamide Lyngbya 11671902 7.7 IC Vero 9 [346] A majuscula 50 cells
Streptomyces 0.00370±0. 4.4±1.0 µM, 916- [157], echinomycin 3197 echinatus 00001 IC50 HMEC 1463 [352]
10.4 µM, Schizothrix gallinamide A 25209862 8.2 TC Vero 1 [353] sp. 50 cells 0.015 µM, Bacillus [164], gramicidin D 16130140 0.000019 EC U937 789 brevis 50 [354] cells 0.035 µM, Streptomyces [164], grisorixin 12780142 0.0012 EC U937 29 griseus 50 [355] cells isopropylstilbe Photorhabdus 6439522 7.67 ND [356] ne sp.
111 Lyngbya 6.4 nM, IC lagunamide A 50901239 0.19 50 <1 [357] majuscula P388 cells
20.5 nM, Lyngbya lagunamide B 50901240 0.91 IC P388 <1 [357] majuscula 50 cells 24.4 nM, Lyngbya lagunamide C 0.29 IC P388 <1 [358] majuscula 50 cells 30 mg/kg Streptomyces [165], lasalocid A 5360807 0.047 LD in mice ND lasaliensis 50 [359] (i.p.) Streptomyces 0.283 µM, [164], lonomycin A 10101992 ribosidifcus 0.0017 EC U937 166 50 [360] TM-481 cells 0.111 µM, Streptomyces [164], lysocellin 3036964 0.012 EC U937 9 cacaoi 50 [361] cells 55 µM, 10% malyngolide Lyngbya 46211780 19 survival H- ND [362] dimer majuscula 460 cells
Micromonosp 2.2 µM, IC [363], manzamine A 6509753 0.015 50 147 ora sp. M42 Vero cells [364]
(other marinoquinoli Ohtaekwangi 30.6 µM, marinoquinoline 54583213 1.8 17 [365] ne B a kribbensis IC50 L6 cells s in the same paper) 0.63±0.02 metacycloprod Streptomyces 0.012±0.00 44521126 µM, IC 44-65 [342] igiosin spectabilis 2 50 BC-1 cells >150 µM, mevalagmape Photorhabdus 73332828 38.4 IC Rat L6 >4 [366] ptide A luminescens 50 cells Micrococcus varians, >30 µM, Bacillus [160]– micrococcin 6446886 0.035 IC HepG2 >857 pumilus, 50 [162] cells Staphylococc us equorum Streptomyces 0.095 µM, [164], monensin A 441145 cinnamonensi 0.0022 EC U937 43 50 [367] s cells 0.399 µM, Streptomyces [164], narasin A 65452 0.0013 EC U937 307 aureofacies 50 [368] cells
112 Streoptomyce 1.49 µM, [335], neothramycin 122745 s sp. MC916- 4.16 IC MRC-5 <1 50 [369] C4 cells Streptomyces 0.266 µM, [164], nigericin 34230 hygroscopicus 0.0014 EC U937 190 50 [370] var. Geldanus cells Streptomyces 3.53 µM, [164], 0.433– nonactin 72519 griseus ETH CC Huh7 6-8 [371], 0.597 50 A7796 cells [372]
Nostoc sp. 78- 120.9 µM, nostocarboline 5326150 0.194 623 [166] 12A IC50 L6 cells
0.095 µM, Streptomyces [335], pactamycin 5289124 0.014 IC MRC-5 7 pactum 50 [373] cells >50 µM, Salinospora >100 salinipostin A 122222702 0.05 EC HFF [154] sp. 50 0 cells 157.5±45.0 Gametocyte Streptomyces 0.0219±0.0 nM, IC salinomycin 3085092 50 3-21 activity reported [374] spp. 123 HMEC-1 in the nM range cells 0.082 µM, salinosporami Salinispora 0.0114±0.0 [375], 11347535 IC RPMI 6-9 de A tropica 019 50 [376] 8226 cells symplocamide 40 nM, IC 44448139 Symploca sp. 0.95 50 <1 [377] A H460 cells
Xenorhabdus 68.5 µM, ChemSpider ID: szentiamide szentirmaii 1.19 58 [378] IC L6 cells 27444379 DSM 16338T 50
Xenorhabdus 14 µM, IC taxlllaid A 122205185 7.19 50 2 [379] indica L6 cells
27.8±10.9 Streptomycete [380], thiostrepton 16129666 1.8 µM, IC 9-22 s sp. 50 [381] HeLa cells
Streptomyces 0.0017±0.0 1.8±0.1 nM, trioxacarcin A 3067465 1 [382] sp. B8652 001 IC50 K1 cells
Streptomyces 0.091±0.00 114±5 nM, trioxacarcin B 3086133 1 [382] sp. B8652 7 IC50 K1 cells
Streptomyces 0.0020±0.0 2.8±0.2 nM, trioxacarcin D 49799759 1-2 [382] sp. B8652 001 IC50 K1 cells
113 Blennothrix tumonoic acid 25058102 cantharidosm 2 ND [383] I um tychonamide Tychonema 2.79 µM, 101846893 0.998 3 [384] A sp. IC50 L6 cells tychonamide Tychonema 3.34 µM, 101846894 1.25 3 [384] B sp. IC50 L6 cells
0.0022 µM, Streptomyces [164], valinomycin 5649 0.0048 EC U937 <1 fulvissimus 50 [385] cells
Oscillatoria 86 µM, IC venturamide A 11225555 8.2 50 10 [386] sp. Vero cells
Oscillatoria 56 µM, IC venturamide B 16115401 5.2 50 11 [386] sp. Vero cells
Oscillatoria viridamide A 25111596 nigro-Wiridis 5.8±0.6 ND [387] OSC3L Xenorhabdus 52.7 µM, xenoamicin A mauleonii 1.81 29 [388] IC L6 cells DSM17908 50
Xenorhabdus No toxicity xenobactin 16.45 ND [389] sp. PB30.3 Rat L6 cells
0.197 µM, Xenorhabdus xenorhabdin-2 185189 4.36 IC Rat L6 <1 [366] doucetiae 50 cells 0.107 µM, Xenorhabdus xenorhabdin-5 185192 1.84 IC Rat L6 <1 [366] doucetiae 50 cells
114 Table 2.1. Colonization-related candidate genes that are differentially regulated between the parental (P1) and passaged (P3) Kco_Z strains.
Fold Change Gene ID Description Process GO Category (log2) Parental Upregulated yadA-like C-terminal region KCO_Z_1310 N/A 0.999 family protein KCO_Z_1067 SOS cell division inhibitor SOS response 0.897 Regulation of transcription, KCO_Z_3679 uxuR transcriptional repressor 0.834 DNA-dependent Adapted upregulated KCO_Z_2042 type III secretion apparatus protein Transport 1.065 glutathione S-transferase, C- KCO_Z_4047 Metabolic process 0.878 terminal domain protein oxidoreductase FAD-binding KCO_Z_0990 Oxidation-reduction process 0.836 domain protein KCO_Z_4206 sugar abc transporter permease Transport 0.827
115 Table 2.2. ROS-producing candidate genes that are differentially regulated between Kco_Z grown in ookinete medium conducive to ROS production, and LB medium.
Fold Change Gene ID Description Process GO Category (log2) KCO_Z_0666 2,3-DHB dehydrogenase Metabolic process 1.466 Riboflavin biosynthetic KCO_Z_1948 GTP cyclohydrolase II 1.384 process KCO_Z_1388 NADPH nitroreductase Oxidation-reduction process 1.359 molybdenum-pterin binding domain KCO_Z_4666 Transmembrane transport 1.169 protein Respiratory electron transport KCO_Z_1849 cytochrome b561 family protein 1.105 chain
116 Table 3.1. Bacterial strains in this study.
Species Strain Source Abbreviation Reference C. aquaticum CC-SEYA-1 DSMZ (DSM 19852) CAQU [262] C. haemolyticum MDA0585 DSMZ (DSM 19808) CHAE [263] C. piscinae LMG 3947 DSMZ (DSM 23278) CPIS [264] C. pseudoviolaceum LMG 3953 DSMZ (DSM 23279) CPSE [264] C. subtsugae PRAA4-1 DSMZ (DSM 17043) CSUB [261] C. vaccinii MWU205 DSMZ (DSM 25150) CVAC [265] C. violaceum ATCC 12472 ATCC CVIO Chromobacterium sp. 968 Yi-Qiang Cheng C968W [289] Chromobacterium sp. 968 ΔdepA Yi-Qiang Cheng C968A [280] Chromobacterium sp. Panama CSPP [141] Chromobacterium sp. W10 NRRL (B-11053) CW10 [390]
117 Table 3.2. Plasmodium falciparum strains in this study.
Strain Source Resistance traits CamWT_C580Y BEI Resources (MRA-1251) artemisinin Dd2 BEI Resources (MRA-150) chloroquine, pyrimethamine, mefloquine GB4 BEI Resources (MRA-925) chloroquine NF54 drug sensitive cytochrome bc1 inhibitors (e.g. atovaquone) SB1-A6 BEI Resources (MRA-1002) [307]
118 Table 3.3. Sources of genome sequences of the Chromobacterium species/strains analyzed in this study.
Species Strain NCBI RefSeq Reference Chromobacterium sp. Panama This study Chromobacterium sp. W10 This study C. aquaticum CC-SEYA-1 NZ_MQZY00000000.1 [26] C. haemolyticum MDA0585 NZ_JONK00000000.1 [27] C. piscinae ND17 NZ_JTGE00000000.1 [28] C. pseudoviolaceum LMG 3953 NZ_MQZX00000000.1 [29] C. subtsugae MWU2920 NZ_LCWP00000000.1 [30] C. vaccinii IIBBL 21-1 NZ_CP017707.1 [8] C. violaceum ATCC 12472 NC_005085.1 [31]
119 Table 3.4. Nucleotide alignment results (megaBLAST) querying putative biosynthetic gene clusters detected in the C. sp. Panama genome by antiSMASH against the genomes of the remaining Chromobacterium spp. used in this study. Similarity index compounded by multiplying query cover with % identity. A similarity index of 90% or above (green) indicates that said C. sp. Panama gene cluster is present in the genome of that particular species; one of
30% or below (red) indicates with confidence that the cluster is not replicated in the other genome.
megaBLAST similarity index Cluster Type Source Size (kb) CAQU CHAE CPIS CPSE CSUB CVAC CVIO CW10 Cluster 01 Homoserine Lactone antiSMASH 20.7 20% 89% 9% 23% 21% 25% 23% 90% Cluster 02 Saccharide ClusterFinder 25.0 65% 86% 66% 66% 65% 60% 70% 86% Cluster 03 NRPS antiSMASH 73.0 28% 86% 25% 28% 28% 28% 28% 86% Cluster 04 Fatty Acid ClusterFinder 33.7 68% 91% 60% 65% 69% 66% 65% 91% Cluster 05 NRPS antiSMASH 45.1 27% 91% 24% 28% 27% 28% 28% 90% Cluster 06 Terpene antiSMASH 21.7 31% 93% 31% 27% 31% 24% 27% 84% Cluster 07 NRPS antiSMASH 43.2 23% 47% 23% 26% 23% 24% 25% 52% Cluster 08 Putative ClusterFinder 15.3 49% 83% 37% 50% 50% 54% 57% 87% Cluster 09 Putative ClusterFinder 4.4 52% 88% 54% 39% 51% 54% 39% 80% Cluster 10 Putative ClusterFinder 9.5 71% 94% 71% 72% 72% 72% 72% 94% Cluster 11 Polyunsaturated Fatty Acid / Other Ketosynthase antiSMASH 51.3 16% 89% 16% 13% 16% 14% 12% 64% Cluster 12 Putative ClusterFinder 8.8 39% 93% 43% 40% 39% 43% 43% 92% Cluster 13 NRPS antiSMASH 51.5 21% 47% 19% 27% 23% 23% 26% 83% Cluster 14 Putative ClusterFinder 7.2 53% 84% 52% 53% 53% 55% 54% 84% Cluster 15 Bacteriocin antiSMASH 10.8 28% 42% 28% 28% 28% 28% 23% 72% Cluster 16 Putative ClusterFinder 8.5 6% 85% 0% 3% 3% 5% 3% 81% Cluster 17 Bacteriocin antiSMASH 10.8 49% 91% 45% 61% 49% 60% 61% 93% Cluster 18 NRPS antiSMASH 43.9 34% 86% 33% 27% 34% 29% 30% 83% Cluster 19 Fatty Acid ClusterFinder 21.1 74% 92% 75% 76% 74% 75% 76% 92% Cluster 20 Putative ClusterFinder 8.9 9% 95% 9% 65% 9% 65% 65% 95% Cluster 21 Saccharide/NRPS ClusterFinder 81.2 40% 60% 27% 36% 38% 31% 34% 70% Cluster 22 Saccharide ClusterFinder 30.1 30% 33% 28% 29% 30% 31% 28% 33% Cluster 23 Putative ClusterFinder 6.5 0% 0% 0% 0% 0% 0% 0% 87% Cluster 24 Putative ClusterFinder 4.6 84% 91% 83% 80% 84% 83% 81% 91% Cluster 25 Putative ClusterFinder 4.8 74% 89% 66% 78% 74% 74% 79% 91% Cluster 26 NRPS antiSMASH 49.6 16% 63% 18% 17% 16% 13% 13% 60% Cluster 27 Putative ClusterFinder 8.5 20% 89% 20% 20% 20% 22% 20% 89% Cluster 28 Putative ClusterFinder 11.7 13% 22% 0% 6% 6% 13% 6% 21% Cluster 29 Saccharide ClusterFinder 31.2 30% 90% 26% 32% 30% 26% 30% 91% Cluster 30 NRPS antiSMASH 6.1 0% 4% 0% 2% 0% 2% 0% 92% Cluster 31 NRPS antiSMASH 3.1 5% 90% 0% 0% 5% 0% 0% 89%
120 Table 4.1. Proteins recovered from active anti-DENV fraction of C. sp. Panama.
ID Name EC number GO - biological process MW 0261 Neutral protease 3.4.24.25 proteolysis 53.4 0262 Aminopeptidase 3.4.11.10 proteolysis 44.7 1115 Ferredoxin--NADP(+) reductase 1.18.1.2 oxidation-reduction process 29.5 1272 Arginine deiminase 3.5.3.6 protein citrullination 46.1 carbohydrate metabolic 1432 Beta-N-acetylhexosaminidase 3.2.1.52 73.1 process L-serine & pyridoxine 3050 Phosphoserine transaminase 2.6.1.52 40.3 biosynthetic process 4002 Bacterioferritin 1.16.3.1 cellular iron ion homeostasis 18.5 4290 Dihydrolipoyl dehydrogenase 1.8.1.4 oxidation-reduction process 50
121
FIGURES
122
Figure 1.1. Plasmodium interactions with the mosquito gut. The Plasmodium ookinete matures in the midgut lumen (L) within the blood bolus and starts its invasion process of the mosquito gut. First, it must cross the peritrophic matrix (PM) and the midgut epithelium (E). The invaded midgut cells quickly initiate apoptosis, forcing the parasite to cross multiple cells before successfully reaching the basement membrane (BM), where it starts maturing into an oocyst.
While traversing the epithelium, the parasite can suffer nitration, facilitating recognition by the mosquito's complement-like system once it exits on the basal side. TEP1, chaperoned by
LRIM1/APL1, binds the parasite and tags it for lysis or melanization. Other mediators, including
LRRD7, TEP3, TEP4, TEP9, and SPCLIP1, have been postulated to be part of this complement- like system. In contrast, CTL4, CTLMA2, SRPN2, and SRPN6 have been identified as Plasmodium agonists, since they act by preventing melanization. Recognition of the midgut microbiota by the midgut epithelium primes the mosquito's immune response and influences the anti-Plasmodium response in this tissue.
123
Figure 1.2. Antiviral immunity in the mosquito gut. Arboviruses can enter the epithelial cells of the midgut epithelium (E) by endocytosis. Mediators of the RNAi pathway detect viral double-stranded RNA (dsRNA) and degrade complementary viral RNA. Other mechanisms of defense result from the recognition of virus-derived ligands by pattern recognition receptors
(PRR), activating the main immune pathways in the mosquito: the Toll, immune deficiency
(IMD), and Janus kinase/signal transducers and activators of transcription (JAK-STAT) pathways. Induction of these pathways triggers signaling cascades that result in the nuclear translocation of the transcription factors Rel1, Rel2, and STAT, respectively. The pathway- activated response leads to the transcription of effector genes that include antimicrobial peptides
(AMPs). AMPs can act intracellularly or extracellularly to mediate virus killing. Apoptotic cells often coincide with viral infection, and it has been suggested that apoptosis can either play an
124 antiviral or a proviral role. L – midgut lumen, PM – peritrophic matrix, BM – basement membrane.
125
Figure 2.1. Colonization of the Anopheles gambiae midgut by Kco_Z and impact on
Plasmodium falciparum sporogonic development. (A) Kco_Z was introduced into a mosquito cohort already containing a cocktail of naturally occurring bacteria through either a blood or sugar meal, and DNA extracted from 10 midguts was sampled daily. The Kco_Z to 16S rRNA ratio was determined using Kco_Z-specific and universal 16S rRNA standard curves. (B) Female
An. gambiae containing their endogenous microflora were provided with PBS or with Kco_Z at the indicated concentration (10x CFU/mL) suspended within 3% sucrose. After 3 to 4 days of being allowed to feed on these suspensions, they were given a P. falciparum-infected blood meal, and oocyst numbers were determined 7 days later. Oocyst counts for three independent replicates are shown (Rep 1-3). Horizontal bars represent the median number of oocysts per treatment; inhibition (%) was estimated based on the comparison of these values to that of the
PBS control. Prevalence represents the proportion of infected mosquitoes per group. Significance
126 was determined using the Mann-Whitney test by comparing treatment groups to their respective control cohorts (*, p<0.05; **, p<0.01; ***, p<0.001).
127
Figure 2.2. Sugar-meal introduction of Kco_Z has no impact on Anopheles gambiae fitness.
(A) Longevity studies were performed following continuous sugar-meal introduction of either
Kco_Z, a bacterial cocktail, or PBS into aseptic mosquito cohorts. A sterile blood meal was provided on day 4 and unfed mosquitoes were censored from the analysis. Survival was monitored daily and continued until 100% mortality was reached. The curves represent the average percent mortality across three replicates; the error bars indicate standard error.
Significance was determined using the log-rank test (Mantel-Cox) with Bonferroni correction using a Kaplan-Meier survival analysis. (B) Comparative fecundity analysis of the Kco_Z-, bacterial cocktail- and PBS-fed, sugar-fed mosquito cohorts. Separate cohorts were provided with a blood meal 72 h after introduction, and circles represent the number of eggs laid per female. Horizontal bars represent the median number of eggs, and error bars indicate the standard error; three pooled biological replicates are shown. p = 0.0596 (Kruskal-Wallis test).
(C) Fertility analysis in Kco_Z-, bacterial cocktail-, and PBS-fed mosquitoes. At 72 h after introduction, mosquitoes were offered a blood meal, and those not engorged were removed. Eggs were collected at 48 h post-blood meal and allowed to hatch in rearing trays. The hatch rate indicates the percentage of eggs giving rise to 1st instar larvae; the error bars indicate the standard error of the mean. p = 0.0741 (Kruskal-Wallis test).
128
Figure 2.3. Kco_Z genome. (A) Circular representation of the Kco_Z genome. The coding regions of both the forward and reverse strands are shown in green; blue represents a negative deviation and red a positive deviation of the average G-C content for this genome of 55.8%. (B)
Distribution of the functional role categories of the protein-encoding genes of Kco_Z using the
SEED database; 47% of those genes were not assigned a functional role and are omitted from this representation.
129
Figure 2.4. Differential gene expression in Kco_Z on different culture media and upon serial passage in the Anopheles gambiae midgut. (A) A parental strain of Kco_Z (P1) was provided to An. gambiae mosquitoes through a blood meal and sampled daily. Colonies persisting for the longest time were reintroduced into a second cohort (P2) and the process repeated a third time to isolate the passaged (P3) strain. The ability of a selected strain of Kco_Z to persist within the An. gambiae midgut was determined through qRT-PCR copy number analysis. (B) Distribution of the functional role category of upregulated (+) and downregulated (-
) transcripts of parental P1 when compared to passaged P3. Functional role categories were assigned to 59/98 upregulated and 29/61 downregulated transcripts using the SEED database.
(C) Distribution of the functional role category of upregulated (+) and downregulated (-)
130 transcripts of Kco_Z grown in ookinete medium when compared to LB medium. Functional role categories were assigned to 15/21 upregulated and 14/20 downregulated transcripts using the
SEED database.
131
Figure 2.5. Exposure to Kco_Z limits Plasmodium’s antioxidant response. Relative P. berghei pre-ookinete expression of thioredoxin reductase (trxr), thioredoxin (trx), thioredoxin 1
(trx1), peroxiredoxin-1 (tpx-1), and 1-Cys peroxiredoxin (1-cys-prx) upon in vitro co-culture
132 with Kco_Z (blue) or Pseudomonas putida (Ppu, black). RT-qPCR datasets were normalized to the level of expression of 18SrRNA, a PBS control, and the expression at 1 h post-exposure for each experimental group. Bars represent the mean ± the standard error of the mean of six biological replicates over two independent experiments (3+3). Significance was determined using a two-tailed one sample t-test to determine whether each transcript was significantly up- or down-regulated (*, p<0.05; **, p<0.01; ***, p<0.001).
133
Figure 2.S1. Impact of blood feeding Kco_Z on mosquito fitness. (A) Longevity studies were performed following blood-meal introduction of either Kco_Z, bacterial cocktail, or PBS into aseptic mosquito cohorts. A sterile blood meal was provided on day 4 (black arrow), and unfed mosquitoes were censored from the analysis. Survival was monitored daily and continued until
100% mortality was reached. The curves represent the average percent mortality across three replicates, and the error bars indicate the standard error. Significance was determined using the log-rank test (Mantel-Cox) using a Kaplan-Meier survival analysis. (B) Fecundity analysis between Kco_Z-, bacterial cocktail- and PBS-fed blood-fed mosquito cohorts. Separate cohorts were provided a second blood meal 72 h after blood-meal introduction of Kco_Z, and circles represent the number of eggs laid per female. Horizontal bars represent the median number of eggs, error bars indicate the standard error, and three pooled biological replicates are shown.
Significance was determined using the Mann-Whitney test. (C) Fertility analysis in Kco_Z-, bacterial cocktail-, and PBS-fed blood-fed mosquitoes. At 72 h after introduction, mosquitoes were offered a second, naive blood meal, and those not engorged were removed. Eggs were collected 48 h post-blood meal and allowed to hatch in rearing trays. The hatch rate indicates the percentage of eggs giving rise to 1st instar larvae; the error bars indicate the standard error of the mean, and significance was determined using an unpaired t-test.
134
Figure 3.1. Bioassay-guided fractionation of Chromobacterium sp. Panama culture supernatant points to romidepsin as the main antimalarial compound. (A) Variation in growth of asexual stage Plasmodium falciparum NF54 upon incubation with culture supernatant (LB, 72 h, 30 ºC) of C. sp. Panama, Sup., when compared to that upon incubation with the respective n-butanol extract,
135 BuOH and successive 1:2 dilutions. The equivalent of 5 mL of culture was used in each case;
Sup. was prepared by filtering the bacterial supernatant (pore size: 0.22 µm), dissecating it, and resuspending in 20% DMSO. 0% inhibition is adjusted for parasite growth in vehicle control
(1% DMSO) and 100% inhibition is matched to that of 250 nM chloroquine. Results are shown as mean ± standard deviation; significance was determined using a one-tailed one sample t-test to determine whether each treatment significantly lowered parasite growth compared to control (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001). CQ, chloroquine. (B) Reverse-phase FPLC chromatogram (absorbance at 210 nm) of n-butanol extract of 250 mL of culture supernantant of
C. sp. Panama (LB, 72h, 30ºC). Fraction boundaries (A-F) indicated in red; fraction F highlighted. Column: RESOURCE RPC 3 mL. Flow rate: 2 mL/min. Gradient elution with 2%-
85% methanol in water 0.1% TFA (dashed line). (C) Variation in growth of asexual stage P. falciparum NF54 upon incubation with fractions recovered from reverse-phase FPLC (cf. 1B).
Fractions were dried and resuspended in proportional amounts of 20% DMSO according to their initial volume. 0% inhibition is adjusted for parasite growth in vehicle control (1% DMSO) and
100% inhibition is matched to that of 250 nM chloroquine. Results are shown as mean ± standard deviation; significance was determined using a one-tailed one sample t-test to determine whether each treatment significantly lowered parasite growth compared to control (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001). CQ, chloroquine. (D) Total ion chromatogram from UPLC-ESI-MS analysis of fraction F6. Structures of romidepsin and previously characterized fragmentation products are shown. (E) Size exclusion FPLC chromatogram
(absorbance at 210 nm) of fraction F (cf. 1B). Fraction boundaries (F1-F10) indicated in red; fractions F6 and F7 highlighted. Column: Superdex Peptide 10/300 GL. Flow rate: 1 mL/min.
Isocratic elution with 20% DMSO in water. (F) Variation in growth of asexual stage P.
136 falciparum NF54 upon incubation with fractions recovered from size-exclusion FPLC (cf. 1E).
Fractions were dried and resuspended in proportional amounts of 20% DMSO according to their initial volume. 0% inhibition is adjusted for parasite growth in vehicle control (1% DMSO) and
100% inhibition is matched to that of 250 nM chloroquine. Results are shown as mean ± standard deviation; significance was determined using a one-tailed one sample t-test to determine whether each treatment significantly lowered parasite growth compared to control (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001). CQ, chloroquine.
137
Figure 3.2. Antimalarial activity of chromobacteria is restricted to the romidepsin- producing subcluster comprised of C. sp. Panama and C. haemolyticum. (A) Variation in growth of asexual stage P. falciparum NF54 upon incubation with n-butanol extracts of approximately 5mL of culture supernatants of different Chromobacterium species (LB, 72h,
30ºC). 0% inhibition is adjusted for parasite growth in vehicle control (1% DMSO) and 100% inhibition is matched to that of 250 nM chloroquine. Results are shown as mean ± standard deviation; significance was determined using a one-tailed one sample t-test to determine whether each treatment significantly lowered parasite growth compared to control (ns, not significant;
*p < 0.05; **p < 0.01; ***p < 0.001). CQ, chloroquine. CAQU, C. aquaticum; CHAE, C. haemolyticum; CPIS, C. piscinae; CPSE, C. pseudoviolaceum; CSPP, C. sp. Panama; CSUB, C. subtsugae; CVAC, C. vaccinii; CVIO, C. violaceum; CW10, C. haemolyticum W10. (B)
UPGMA tree generated from a distance matrix inferred from pairwise genome average nucleotide identity (gANI). A distance of 0.02 indicates that the gANI of two species differs by 2
138 percent points. Cophenetic Correlation Coefficient (CP) = 0.997961877789881. (C) Alignment between the reference romidepsin biosynthetic gene cluster in C. sp. 968 (C968; GenBank:
EF210776.1) to the ones present in the genomes of C. sp. Panama (CSPP) and C. haemolyticum, both MDA0585 (CHAE) and W10 (CW10) strains. Geneious alignment in Geneious v5.4 (global alignment with free end gaps; cost matrix: 65% similarity).
139
Figure 3.3. Romidepsin strongly inhibits Plasmodium. (A) Dose-response curve of romidepsin against P. falciparum NF54. 0% inhibition is adjusted for parasite growth in vehicle control (1%
DMSO) and 100% inhibition is matched to that of 250 nM chloroquine. IC50 is indicated together with the 95% confidence interval returned after fitting a curve to the normalized data using the least squares method with variable slope. (B) Female An. gambiae were allowed to feed on romidepsin at the indicated concentrations in 0.5% DMSO 3% sucrose for 1 day prior to being given a P. falciparum NF54-infected blood meal; control group was fed on vehicle alone. Oocyst numbers determined 7 days later for three independent replicates are shown. Horizontal bars represent the median number of oocysts per treatment. Significance was determined using the
Kruskal-Wallis test by comparing treatment groups to the control and correcting for multiple comparisons by the Dunn’s test (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001).
140
Figure 3.4. Romidepsin production is necessary and sufficient for the antimalarial activity of Chromobacterium spp.. (A) Variation in growth of asexual stage P. falciparum NF54 upon incubation with n-butanol extracts of approximately 5mL of culture supernatants (LB, 72h, 30ºC) of C. sp. 968 wildtype (wt) and depA-null mutant (∆depA). 0% inhibition is adjusted for parasite growth in vehicle control (1% DMSO) and 100% inhibition is matched to that of 250 nM chloroquine. Results are shown as mean ± standard deviation; significance was determined using a one-tailed one sample t-test to determine whether each treatment significantly lowered parasite growth compared to control (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001). CQ, chloroquine. (B) Female An. gambiae were provided either PBS, C. sp. 968 wildtype (wt) or its
∆depA mutant suspended within 3 % sucrose. After 1 day of being allowed to feed on these suspensions, they were given a P. falciparum NF54-infected blood meal, and oocyst numbers were determined 7 days later; oocyst counts for three independent replicates are shown.
Horizontal bars represent the median number of oocysts per treatment; inhibition (%) was estimated based on the comparison of these values to that of the PBS control. Prevalence represents the proportion of infected mosquitoes per group. Significance was determined using the Kruskal-Wallis test by comparing treatment groups to the control and correcting for multiple comparisons by the Dunn’s test (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001). (C)
141 Evolutionary relationships between C. sp. 968, other chromobacteria and other related species
(Neisseria gonorrhoeae NCTC13800; Neisseria meningitidis LNP21362; Pseudomonas aeruginosa DSM 50071) based on 16S rRNA genomic sequences as inferred by the UPGMA method. The optimal tree with the sum of branch lengths equal to 0.33018119 is shown. The tree is drawn to scale, with branch lengths in the same units as those of the evolutionary distances used to infer the phylogenetic tree. The analysis was conducted in MEGA6 and involved 13 nucleotide sequences for a total of 925 positions in the final dataset.
142
Figure 3.5. Romidepsin surpasses common drug resistance mechanisms and interferes with
P. falciparum HDAC activity. Dose-response curve of romidepsin against P. falciparum
CamWT_C580Y (A, resistant to artemisinin), Dd2 (B, resistant to chloroquine, pyrimethamine
143 and mefloquine), GB4 (C, resistant to chloroquine) and SB1-A6 (D, resistant to cytochrome bc1 inhibitors such as atovaquone). 0% inhibition is adjusted for parasite growth in vehicle control
(1% DMSO) and 100% inhibition is matched to that of 250 nM chloroquine. IC50 is indicated together with the 95% confidence interval returned after fitting a curve to the normalized data using the least squares method with variable slope. (E) Inhibition of HDAC activity of P. falciparum NF54 nuclear protein extracts in the presence of romidepsin, as measured by endpoint immunodetection of remaining acetylated histone substrate following incubation.
Results are normalized to vehicle control of 1.3% DMSO (0% inhibition) and absence of P. falciparum nuclear protein extract (100% inhibition).
144
Figure 3.S1. UPLC chromatograms of Fraction F. (top) base peak ion chromatogram (BPI),
(bottom) absorbance at 210 nm.
145
Figure 3.S2. Predicted NRPS organization and function of the Sch 20561/2 biosynthetic cluster. Listed in the table are Csp_P genes and predicted functions of encoded proteins in the putative Sch 20561/2 biosynthetic cluster. In bold are the NRPS domains that allowed prediction of a hexa(hepta)peptide core. Synthesis is hypothesized to be initiated by a (R)-β- hydroxylmyristoylation of the starting Dhb unit and continuing through presently indeterminent steps to the lipidated tripeptide in the first module of CspP_3681. Subsequent peptide elongation steps are assigned consistent with A domain substrate preference shown in bold above each.
Final macrolactonization is well-precedented by the action of the C-terminal thioesterase (TE) domain to release the aglycone, Sch 20561, which is regiospecifically glycosylated to Sch 20562. adenylation domain (A), peptidyl-carrier protein (PCP), condensation domain (C), epimerization domain (E), N-methylation domain (nMT).
146
Figure 3.S3. Tentative structural assignments of C. sp Panama-isolated Sch 20561/2.
147 B o th F e m a le s M a le s
1 0 0 d e p A 1 0 0 d e p A 1 0 0 d e p A
l l
wt wt l wt
a
a
a
v
v
v
i i
C o n tro l C o n tro l i C o n tro l
v
v
v
r
r
r
u
u
u
s
s
s
t t
5 0 5 0 t 5 0
n
n
n
e
e
e
c
c
c
r
r
r
e
e
e
P
P P
0 0 0 0 2 4 6 8 0 2 4 6 8 0 2 4 6 8 D a y s D a y s D a y s
Figure 3.S4. Survival curves of Anopheles gambiae (females only, males only and both) upon ingestion of Chromobacterium sp. 968 wildtype or ∆depA at approximately 106
CFU/mL.
148
Figure 4.1. Anti-DENV activity of Chromobacterium sp. Panama is mediated by secreted proteinous factors. DENV virus titers (A) in the presence or absence of culture supernatant of
C. cp. Panama following infection in BHK-21 and C6/36 cells, as determined by plaque or focus forming assays, respectively; (B) in BHK-21 cells upon exposure to different solvent extracts of culture supernatants of C. sp. Panama; (C) in BHK-21 cells in the presence or absence of culture supernatant of C. cp. Panama incubated for 1 h at the indicated temperatures; and (D) upon exposure to protein-rich ammonium sulfate pellet following extraction at 70% of the culture supernatant of C. sp. Panama (Pellet), the desalted supernatant of that extraction (Sup.) or
149 control buffer (0.1M Tris-Hcl). Significance determined using unpaired t tests; (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001).
150
Figure 4.2. Protein extracts of culture supernatants of C. sp. Panama inhibit DENV entry by mediating degradation of the E protein. (A) DENV titers following exposure to a protein extract of the C. sp. Panama culture supernatant before or after viral attachment to BHK-21 cells; significance determined using unpaired t tests. (B) Representative negative stained
151 transmission EM images of control DENV particles (arrow head) and those treated with C. sp.
Panama culture supernatant protein extract (empty arrow head); low magnification: 93,000X, high magnification: 245,000X. (C) Size distribution of the viral particles as measured from the
TEM images collected in B; mean and standard deviation indicated; significance determined using unpaired t test. (D) Western blot (anti-E-DENV) of DENV2 proteins when exposed to the
C. sp. Panama culture supernatant protein extract. h.i., heat-inactivated. (E) Fluorescence microscopy of DENV infection dynamics in the presence of the protein extract of the C. sp.
Panama. Cells were fixed with 4% paraformaldehyde, stained for DNA (DAPI-blue), and F- actin (Phalloidin-green). Membrane-bound DENV (arrow head) were immunostained with anti-E antibody (Red). (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001)
152
Figure 4.3. Purification of the anti-DENV proteins of C. sp. Panama. (A) Hydrophobic interaction FPLC chromatogram (absorbance at 214 nm) of the protein extract of C. sp. Panama.
Fraction H2 highlighted. Column: HiTrap FF 1 mL. Flow rate: 1 mL/min. Gradient elution from
1.5 to 0 M ammonium sulfate in 50 mM sodium phosphate (pH 7.0). (B) Hydrophobic interaction FPLC chromatogram (absorbance at 214 nm) of H2 from A. Fraction H2p highlighted. Column: HiTrap HP 1 mL. Flow rate: 1 mL/min. Gradient elution from 1.5 to 0 M ammonium sulfate in 50 mM sodium phosphate (pH 7.0); conductivity (mS/cm) showed by dashed line. (C) SDS-PAGE protein gel of H2, H2p, DENV2 protein extract and co-incubation of this extract with H2p.
153
Figure 4.4. Aminopeptidase secreted by C. sp. Panama induces DENV E protein degradation, thereby inhibiting viral entry. (A) Protein sequence alignment of CSPP0261 and orthologues in Pseudomonas aeruginosa and Vibrio proteolyticus. (B) Protein sequence alignment of CSPP0262 and ortholog in Vibrio proteolyticus. (C) Western blot (anti-E-DENV) of DENV2 proteins when exposed to the culture supernatant of C. sp. Panama with or without supplementation with 10 µM bestatin or 100 µM phosphoramidon (P-ramidon). (D) DENV titers in BHK-21 cells following incubation of the virus with the C. sp. Panama culture supernatant with or without supplementation with 10 µM bestatin or 100 µM phosphoramidon (P-ramidon); significance determined using unpaired t tests. (ns, not significant; *p < 0.05; **p < 0.01;
***p < 0.001)
154 ABBREVIATIONS
1-cys-prx 1-cys-peroxiredoxin Ag Anopheles gambiae Ago argonaute AMP antimicrobial peptide APL Anopheles Plasmodium-responsive leucine-rich repeat protein APOD apolipoprotein D ATCC American Type Culture Collection ATP adenosine triphosphate cDNA complementary DNA CFU colony-forming unit CID compound identifier CLIP/SPCLIP Clip-domain serine protease CTL C-type lectin CTLMA mannose-binding C-type lectin DENV Dengue virus DMEM Dulbecco's modified Eagle's medium DMSO dimethyl sulfoxide DNA deoxyribonucleic acid DOPA dihydroxyphenylalanine Dscam Down syndrome cell adhesion molecule DTT dithiothreitol EBP enolase-binding protein EDTA ethylenediaminetetraacetic acid ESI/MS electrospray ionization mass spectrometry FAD flavin adenine dinucleotide FBN fibrinogen-related protein FBS fetal bovine serum FDA US Food and Drug Administration FMN flavin mononucleotide FPLC fast performance liquid chromatography FREP fibrinogen-related protein gANI genome average nucleotide identity GFP green fluorescence protein GO gene ontology GST glutathione S-transferase GTP guanosine triphosphate HDAC histone deacetylase HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
155 HIC hydrophobic interaction chromatography HMBPP (E)-4-hydroxy-3-methyl-but-2-enyl pyrophosphate HPLC high performance liquid chromatography IC50 half maximal inhibitory concentration IMD immune deficiency JAK-STAT Janus kinase/signal transducers and activators of transcription JNK Jun-N-terminal kinase Kco_Z Kosakonia cowanii Zambiae LB lysogeny broth Log2FC log base 2 fold change Lp Lipophorin LRIM leucine-rich repeat immune protein LRR leucine-rich repeat LRRD leucine-rich repeat domain containing protein MDL myeloid DAP12- associating lectin MS mass spectrometry MyD88 myeloid differentiation primary response 88 NADPH nicotinamide adenine dinucleotide phosphate NCBI National Center for Biotechnology Information NOS nitric oxide synthase NRPS non-ribosomal peptide OD600 optical density at wavelength 600 nm PAGE polyacrylamide gel electrophoresis PAMP pathogen-associated molecular pattern PBS phosphate-buffered saline PCR polymerase chain reaction PGRP-LC peptidoglycan recognition protein LC PM peritrophic matrix PMSF phenylmethane sulfonyl fluoride PPO prophenoloxidase Ppu Pseudomonas putida PRR pattern recognition receptor qPCR quantitative polymerase chain reaction RBC red blood cell Rel NF-κB relish-like transcription factor RNA ribonucleic acid RNAi RNA interference RNS reactive nitrogen species ROS reactive oxygen species rRNA ribosomal ribonucleic acid
156 SDS sodium dodecyl sulfate SI selective index SRPN serpin STAT signal transducers and activators of transcription TBS Tris-buffered saline TEP thioester protein Tpx-1 peroxiredoxin-1 TPxGl thioredoxin peroxidase Tris tris(hydroxymethyl)aminomethane Trx-1 thioredoxin-1 TrxR thioredoxin reductase UPGMA unweighted pair group method with arithmetic mean UPLC ultra performance liquid chromatography WHO World Health Organization WNV West Nile virus YFV yellow fever virus ZIKV Zika virus
157 REFERENCES
[1] B. Gates, “The Deadliest Animal in the World,” Gates Notes, 2014.
[2] WHO, “World Malaria Report 2016,” 2017.
[3] S. Bhatt et al., “The global distribution and burden of dengue.,” Nature, vol. 496, no.
7446, pp. 504–7, Apr. 2013.
[4] N. J. Dennison, N. Jupatanakul, and G. Dimopoulos, “The mosquito microbiota influences
vector competence for human pathogens,” Curr. Opin. Insect Sci., vol. 3, pp. 6–13, Sep.
2014.
[5] J. A. Vaughan, “Population dynamics of Plasmodium sporogony,” Trends Parasitol., vol.
23, no. 2, pp. 63–70, Feb. 2007.
[6] S. Wang and M. Jacobs-Lorena, “Genetic approaches to interfere with malaria
transmission by vector mosquitoes,” Trends Biotechnol., vol. 31, no. 3, pp. 185–193,
2013.
[7] Y. Alavi et al., “The dynamics of interactions between Plasmodium and the mosquito: a
study of the infectivity of Plasmodium berghei and Plasmodium gallinaceum, and their
transmission by Anopheles stephensi, Anopheles gambiae and Aedes aegypti,” Int J
Parasitol, vol. 33, no. 9, pp. 933–943, 2003.
[8] A. Cohuet et al., “Anopheles and Plasmodium: from laboratory models to natural systems
in the field,” EMBO Rep., vol. 7, no. 12, pp. 1285–1289, 2006.
[9] A. M. Mendes et al., “Conserved mosquito/parasite interactions affect development of
Plasmodium falciparum in Africa,” PLoS Pathog., vol. 4, no. 5, p. e1000069, 2008.
158 [10] G. Jaramillo-Gutierrez, J. Rodrigues, G. Ndikuyeze, M. Povelones, A. Molina-Cruz, and
C. Barillas-Mury, “Mosquito immune responses and compatibility between Plasmodium
parasites and anopheline mosquitoes,” BMC Microbiol., vol. 9, no. 1, p. 154, 2009.
[11] R. C. Smith, J. Vega-Rodr??guez, and M. Jacobs-Lorena, “The Plasmodium bottleneck:
Malaria parasite losses in the mosquito vector,” Mem. Inst. Oswaldo Cruz, vol. 109, no. 5,
pp. 644–661, 2014.
[12] H. M. Müller, J. M. Crampton, a della Torre, R. Sinden, and a Crisanti, “Members of a
trypsin gene family in Anopheles gambiae are induced in the gut by blood meal.,” EMBO
J., vol. 12, no. 7, pp. 2891–2900, 1993.
[13] A. N. Dana et al., “Gene expression patterns associated with blood-feeding in the malaria
mosquito Anopheles gambiae,” BMC Genomics, vol. 6, p. 5, 2005.
[14] E. G. Abraham and M. Jacobs-Lorena, “Mosquito midgut barriers to malaria parasite
development,” Insect Biochem. Mol. Biol., vol. 34, no. 7, pp. 667–671, 2004.
[15] B. T. Beerntsen, A. A. James, and B. M. Christensen, “Genetics of Mosquito Vector
Competence,” Microbiol. Mol. Biol. Rev., vol. 64, no. 1, pp. 115–137, 2000.
[16] R. R. Dinglasan et al., “The Anopheles gambiae adult midgut peritrophic matrix
proteome,” Insect Biochem. Mol. Biol., vol. 39, no. 2, pp. 125–134, 2009.
[17] A. G. Richards and P. A. Richards, “The peritrophic membranes of insects,” Annu Rev
Entomol, vol. 22, pp. 219–240, 1977.
[18] M. Moreno-García, B. Recio-Tótoro, F. Claudio-Piedras, and H. Lanz-Mendoza, “Injury
and immune response: applying the danger theory to mosquitoes.,” Front. Plant Sci., vol.
159 5, no. September, p. 451, 2014.
[19] L. Shao, M. Devenport, and M. Jacobs-Lorena, “The peritrophic matrix of hematophagous
insects.,” Arch. Insect Biochem. Physiol., vol. 47, no. 2, pp. 119–125, 2001.
[20] M. Devenport, H. Fujioka, and M. Jacobs-Lorena, “Storage and secretion of the
peritrophic matrix protein Ag-Aper1 and trypsin in the midgut of Anopheles gambiae,”
Insect Mol Biol, vol. 13, no. 4, pp. 349–358, 2004.
[21] M. Shahabuddin and D. C. Kaslow, “Plasmodium: parasite chitinase and its role in
malaria transmission.,” Exp. Parasitol., vol. 79, no. 1, pp. 85–88, 1994.
[22] R. K. Bhatnagar, N. Arora, S. Sachidanand, M. Shahabuddin, D. Keister, and V. S.
Chauhan, “Synthetic propeptide inhibits mosquito midgut chitinase and blocks sporogonic
development of malaria parasite,” Biochem. Biophys. Res. Commun., vol. 304, no. 4, pp.
783–787, 2003.
[23] Y. L. Tsai, R. E. Hayward, R. C. Langer, D. A. Fidock, and J. M. Vinetz, “Disruption of
Plasmodium falciparum chitinase markedly impairs parasite invasion of mosquito
midgut,” Infect. Immun., vol. 69, no. 6, pp. 4048–4054, 2001.
[24] J. T. Dessens et al., “Knockout of the rodent malaria parasite chitinase PbCHT1 reduces
infectivity to mosquitoes,” Infect. Immun., vol. 69, no. 6, pp. 4041–4047, 2001.
[25] M. Huber, E. Cabib, and L. H. Miller, “Malaria parasite chitinase and penetration of the
mosquito peritrophic membrane.,” Proc. Natl. Acad. Sci. U. S. A., vol. 88, no. 7, pp. 2807–
2810, 1991.
[26] M. Di Luca et al., “Human chitotriosidase helps Plasmodium falciparum in the Anopheles
160 midgut,” J. Vector Borne Dis., vol. 43, no. 3, pp. 144–146, 2006.
[27] A. Giansanti, M. Bocchieri, V. Rosato, and S. Musumeci, “A fine functional homology
between chitinases from host and parasite is relevant for malaria transmissibility,”
Parasitol. Res., vol. 101, no. 3, pp. 639–645, 2007.
[28] Z. Shen and M. Jacobs-Lorena, “Characterization of a novel gut-specific chitinase gene
from the human malaria vector Anopheles gambiae.,” J. Biol. Chem., vol. 272, no. 46, pp.
28895–28900, 1997.
[29] J. Zhang et al., “Comparative Genomic Analysis of Chitinase and Chitinase-Like Genes in
the African Malaria Mosquito (Anopheles gambiae),” PLoS One, vol. 6, no. 5, p. e19899,
2011.
[30] G. Zhang et al., “Anopheles midgut FREP1 mediates plasmodium invasion,” J. Biol.
Chem., vol. 290, no. 27, pp. 16490–16501, 2015.
[31] G. Niu, B. Wang, G. Zhang, J. B. King, R. H. Cichewicz, and J. Li, “Targeting mosquito
FREP1 with a fungal metabolite blocks malaria transmission,” Sci Rep, vol. 5, p. 14694,
2015.
[32] H. Zieler and J. a Dvorak, “Invasion in vitro of mosquito midgut cells by the malaria
parasite proceeds by a conserved mechanism and results in death of the invaded midgut
cells.,” Proc. Natl. Acad. Sci. U. S. A., vol. 97, no. 21, pp. 11516–11521, 2000.
[33] R. W. Moon et al., “A cyclic GMP signalling module that regulates gliding motility in a
malaria parasite,” PLoS Pathog, vol. 5, no. 9, p. e1000599, 2009.
[34] J. Vega-Rodríguez et al., “Multiple pathways for Plasmodium ookinete invasion of the
161 mosquito midgut.,” Proc. Natl. Acad. Sci. U. S. A., vol. 111, no. 4, pp. E492-500, 2014.
[35] L. A. Baton and L. C. Ranford-Cartwright, “How do malaria ookinetes cross the mosquito
midgut wall?,” Trends Parasitol., vol. 21, no. 1, pp. 22–28, 2005.
[36] Y. S. Han, J. Thompson, F. C. Kafatos, and C. Barillas-Mury, “Molecular interactions
between Anopheles stephensi midgut cells and Plasmodium berghei: the time bomb theory
of ookinete invasion of mosquitoes,” EMBO J., vol. 19, no. 22, pp. 6030–6040, 2000.
[37] G. K. Christophides, D. Vlachou, and F. C. Kafatos, “Comparative and functional
genomics of the innate immune system in the malaria vector Anopheles gambiae,”
Immunol Rev, vol. 198, pp. 127–148, 2004.
[38] L. S. Garver, G. de Almeida Oliveira, and C. Barillas-Mury, “The JNK Pathway Is a Key
Mediator of Anopheles gambiae Antiplasmodial Immunity,” PLoS Pathog., vol. 9, no. 9,
p. e1003622, 2013.
[39] S. Meister et al., “Anopheles gambiae PGRPLC-mediated defense against bacteria
modulates infections with malaria parasites,” PLoS Pathog., vol. 5, no. 8, p. e1000542,
2009.
[40] L. S. Garver, Y. Dong, and G. Dimopoulos, “Caspar controls resistance to Plasmodium
falciparum in diverse anopheline species,” PLoS Pathog., vol. 5, no. 3, p. e1000335, 2009.
[41] A. M. Clayton, C. M. Cirimotich, Y. Dong, and G. Dimopoulos, “Caudal is a negative
regulator of the Anopheles IMD Pathway that controls resistance to Plasmodium
falciparum infection,” Dev. Comp. Immunol., vol. 39, no. 4, pp. 323–332, 2013.
[42] C. M. Cirimotich, Y. Dong, L. S. Garver, S. Sim, and G. Dimopoulos, “Mosquito immune
162 defenses against Plasmodium infection,” Dev. Comp. Immunol., vol. 34, no. 4, pp. 387–
395, 2010.
[43] A. M. Clayton, Y. Dong, and G. Dimopoulos, “The anopheles innate immune system in
the defense against malaria infection,” J. Innate Immun., vol. 6, no. 2, pp. 169–181, 2014.
[44] N. J. Dennison, O. J. BenMarzouk-Hidalgo, and G. Dimopoulos, “MicroRNA-regulation
of Anopheles gambiae immunity to Plasmodium falciparum infection and midgut
microbiota,” Dev. Comp. Immunol., vol. 49, no. 1, pp. 170–178, 2015.
[45] Y. Dong, R. Aguilar, Z. Xi, E. Warr, E. Mongin, and G. Dimopoulos, “Anopheles
gambiae immune responses to human and rodent Plasmodium parasite species,” PLoS
Pathog., vol. 2, no. 6, pp. 0513–0525, 2006.
[46] Y. Dong and G. Dimopoulos, “Anopheles fibrinogen-related proteins provide expanded
pattern recognition capacity against bacteria and malaria parasites,” J. Biol. Chem., vol.
284, no. 15, pp. 9835–9844, 2009.
[47] Y. Dong, C. M. Cirimotich, A. Pike, R. Chandra, and G. Dimopoulos, “Anopheles NF-
kappaB-regulated splicing factors direct pathogen-specific repertoires of the hypervariable
pattern recognition receptor AgDscam,” Cell Host Microbe, vol. 12, no. 4, pp. 521–530,
2012.
[48] Y. Dong, H. E. Taylor, and G. Dimopoulos, “AgDscam, a hypervariable immunoglobulin
domain-containing receptor of the Anopheles gambiae innate immune system,” PLoS
Biol., vol. 4, no. 7, p. e229, 2006.
[49] P. H. Smith et al., “Alternative splicing of the Anopheles gambiae Dscam gene in diverse
163 Plasmodium falciparum infections,” Malar. J., vol. 10, p. 156, 2011.
[50] S. Blandin et al., “Complement-like protein TEP1 is a determinant of vectorial capacity in
the malaria vector Anopheles gambiae,” Cell, vol. 116, no. 5, pp. 661–670, 2004.
[51] E. A. Levashina, L. F. Moita, S. Blandin, G. Vriend, M. Lagueux, and F. C. Kafatos,
“Conserved Role of a Complement-like Protein in Phagocytosis Revealed by dsRNA
Knockout in Cultured Cells of the Mosquito, Anopheles gambiae,” Cell, vol. 104, no. 5,
pp. 709–718, 2001.
[52] S. A. Blandin et al., “Dissecting the genetic basis of resistance to malaria parasites in
Anopheles gambiae.,” Science, vol. 326, no. 5949, pp. 147–50, 2009.
[53] M. Fraiture et al., “Two Mosquito LRR Proteins Function as Complement Control Factors
in the TEP1-Mediated Killing of Plasmodium,” Cell Host Microbe, vol. 5, no. 3, pp. 273–
284, 2009.
[54] M. Povelones, R. M. Waterhouse, F. C. Kafatos, and G. K. Christophides, “Leucine-Rich
Repeat Protein Complex Activates Mosquito Complement in Defense Against
Plasmodium Parasites,” Science (80-. )., vol. 324, no. 5924, pp. 258–261, 2009.
[55] M. M. Riehle et al., “Anopheles gambiae APL1 is a family of variable LRR proteins
required for Rel1-mediated protection from themalaria parasite, Plasmodium berghei,”
PLoS One, vol. 3, no. 11, p. e3672, 2008.
[56] I. Holm et al., “Diverged Alleles of the Anopheles gambiae Leucine-Rich Repeat Gene
APL1A Display Distinct Protective Profiles against Plasmodium falciparum,” PLoS One,
vol. 7, no. 12, p. e52684, 2012.
164 [57] S. M. Rottschaefer et al., “Exceptional diversity, maintenance of polymorphism, and
recent directional selection on the APL1 malaria resistance genes of Anopheles gambiae,”
PLoS Biol., vol. 9, no. 3, p. e1000600, 2011.
[58] C. Mitri et al., “Fine pathogen discrimination within the APL1 gene family protects
Anopheles gambiae against human and rodent malaria species,” PLoS Pathog., vol. 5, no.
9, p. e1000576, 2009.
[59] L. S. Garver et al., “Anopheles Imd pathway factors and effectors in infection intensity-
dependent anti-Plasmodium action,” PLoS Pathog., vol. 8, no. 6, p. e1002737, 2012.
[60] M. Williams, B. J. Summers, and R. H. G. Baxter, “Biophysical Analysis of Anopheles
gambiae Leucine-Rich Repeat Proteins APL1A1, APLB and APL1C and Their Interaction
with LRIM1,” PLoS One, vol. 10, no. 3, p. e0118911, 2015.
[61] M. Povelones, L. M. Upton, K. A. Sala, and G. K. Christophides, “Structure-function
analysis of the anopheles gambiae LRIM1/APL1C complex and its interaction with
complement C3-like protein TEP1,” PLoS Pathog., vol. 7, no. 4, p. e1002023, 2011.
[62] M. Povelones et al., “The CLIP-Domain Serine Protease Homolog SPCLIP1 Regulates
Complement Recruitment to Microbial Surfaces in the Malaria Mosquito Anopheles
gambiae,” PLoS Pathog., vol. 9, no. 9, p. e1003623, 2013.
[63] S. A. Blandin, E. Marois, and E. A. Levashina, “Antimalarial Responses in Anopheles
gambiae: From a Complement-like Protein to a Complement-like Pathway,” Cell Host
Microbe, vol. 3, no. 6, pp. 364–374, 2008.
[64] C. Barillas-Mury, “CLIP proteases and Plasmodium melanization in Anopheles gambiae,”
165 Trends Parasitol., vol. 23, no. 7, pp. 297–299, 2007.
[65] M. Sugumaran, “Comparative Biochemistry of Eumelanogenesis and the Protective Roles
of Phenoloxidase and Melanin in Insects,” Pigment Cell Res., vol. 15, no. 1, pp. 2–9,
2002.
[66] S. Blandin and E. A. Levashina, “Mosquito immune responses against malaria parasites,”
Curr. Opin. Immunol., vol. 16, no. 1, pp. 16–20, 2004.
[67] S. H. Shiao, M. M. A. Whitten, D. Zachary, J. A. Hoffmann, and E. A. Levashina, “Fz2
and Cdc42 mediate melanization and actin polymerization but are dispensable for
Plasmodium killing in the mosquito midgut,” PLoS Pathog., vol. 2, no. 12, pp. 1152–
1164, 2006.
[68] D. Vlachou, T. Schlegelmilch, G. K. Christophides, and F. C. Kafatos, “Functional
genomic analysis of midgut epithelial responses in Anopheles during Plasmodium
invasion,” Curr. Biol., vol. 15, no. 13, pp. 1185–1195, 2005.
[69] T. Schlegelmilch and D. Vlachou, “Cell biological analysis of mosquito midgut invasion:
the defensive role of the actin-based ookinete hood.,” Pathog. Glob. Health, vol. 107, no.
8, pp. 480–92, 2013.
[70] M. A. Osta, G. K. Christophides, and F. C. Kafatos, “Effects of mosquito genes on
Plasmodium development.,” Science, vol. 303, no. 5666, pp. 2030–2, 2004.
[71] J. Volz, H. M. Muller, A. Zdanowicz, F. C. Kafatos, and M. A. Osta, “A genetic module
regulates the melanization response of Anopheles to Plasmodium,” Cell Microbiol, vol. 8,
no. 9, pp. 1392–1405, 2006.
166 [72] K. Michel, A. Budd, S. Pinto, T. J. Gibson, and F. C. Kafatos, “Anopheles gambiae
SRPN2 facilitates midgut invasion by the malaria parasite Plasmodium berghei,” EMBO
Rep., vol. 6, no. 9, pp. 891–897, 2005.
[73] E. G. Abraham et al., “An immune-responsive serpin, SRPN6, mediates mosquito defense
against malaria parasites,” Proc. Natl. Acad. Sci. U.S.A., vol. 102, no. 45, pp. 16327–
16332, 2005.
[74] S. Kumar et al., “The role of reactive oxygen species on Plasmodium melanotic
encapsulation in Anopheles gambiae.,” Proc. Natl. Acad. Sci. U. S. A., vol. 100, no. 24,
pp. 14139–44, 2003.
[75] H. Lanz-Mendoza, S. Hernandez-Martinez, M. Ku-Lopez, C. Rodriguez Mdel, A.
Herrera-Ortiz, and M. H. Rodriguez, “Superoxide anion in Anopheles albimanus
hemolymph and midgut is toxic to Plasmodium berghei ookinetes,” J. Parasitol., vol. 88,
no. 4, pp. 702–706, 2002.
[76] J. H. M. Oliveira, R. L. S. Gon??alves, G. A. Oliveira, P. L. Oliveira, M. F. Oliveira, and
C. Barillas-Mury, “Energy metabolism affects susceptibility of Anopheles gambiae
mosquitoes to Plasmodium infection,” Insect Biochem. Mol. Biol., vol. 41, no. 6, pp. 349–
355, 2011.
[77] R. L. S. Gonçalves et al., “Mitochondrial reactive oxygen species modulate mosquito
susceptibility to Plasmodium infection,” PLoS One, vol. 7, no. 7, p. e41083, 2012.
[78] A. Molina-Cruz et al., “Reactive oxygen species modulate Anopheles gambiae immunity
against bacteria and Plasmodium,” J. Biol. Chem., vol. 283, no. 6, pp. 3217–3223, 2008.
167 [79] L. Gupta et al., “The STAT pathway mediates late-phase immunity against Plasmodium in
the mosquito Anopheles gambiae,” Cell Host Microbe, vol. 5, no. 5, pp. 498–507, 2009.
[80] S. Luckhart, Y. Vodovotz, L. Cui, and R. Rosenberg, “The mosquito Anopheles stephensi
limits malaria parasite development with inducible synthesis of nitric oxide.,” Proc. Natl.
Acad. Sci. U. S. A., vol. 95, no. 10, pp. 5700–5705, 1998.
[81] S. Vijay et al., “Parasite killing in malaria non-vector mosquito Anopheles culicifacies
species B: implication of nitric oxide synthase upregulation,” PLoS One, vol. 6, no. 4, p.
e18400, 2011.
[82] R. J. DeJong, L. M. Miller, A. Molina-Cruz, L. Gupta, S. Kumar, and C. Barillas-Mury,
“Reactive oxygen species detoxification by catalase is a major determinant of fecundity in
the mosquito Anopheles gambiae,” Proc. Natl. Acad. Sci. U.S.A., vol. 104, no. 7, pp.
2121–2126, 2007.
[83] M. A. Kang, T. M. Mott, E. C. Tapley, E. E. Lewis, and S. Luckhart, “Insulin regulates
aging and oxidative stress in Anopheles stephensi,” J Exp Biol, vol. 211, no. Pt 5, pp.
741–748, 2008.
[84] R. C. Felix, P. Muller, V. Ribeiro, H. Ranson, and H. Silveira, “Plasmodium infection
alters Anopheles gambiae detoxification gene expression,” BMC Genomics, vol. 11, p.
312, 2010.
[85] G. Jaramillo-Gutierrez, A. Molina-Cruz, S. Kumar, and C. Barillas-Mury, “The Anopheles
gambiae oxidation resistance 1 (OXR1) gene regulates expression of enzymes that
detoxify reactive oxygen species,” PLoS One, vol. 5, no. 6, p. e11168, 2010.
168 [86] W. Surachetpong, N. Pakpour, K. W. Cheung, and S. Luckhart, “Reactive oxygen species-
dependent cell signaling regulates the mosquito immune response to Plasmodium
falciparum,” Antioxid. Redox Signal., vol. 14, no. 6, pp. 943–955, 2011.
[87] A. Herrera-Ortiz, J. Martinez-Barnetche, N. Smit, M. H. Rodriguez, and H. Lanz-
Mendoza, “The effect of nitric oxide and hydrogen peroxide in the activation of the
systemic immune response of Anopheles albimanus infected with Plasmodium berghei,”
Dev. Comp. Immunol., vol. 35, no. 1, pp. 44–50, 2011.
[88] S. Luckhart et al., “Sustained activation of Akt elicits mitochondrial dysfunction to block
Plasmodium falciparum infection in the mosquito host,” PLoS Pathog., vol. 9, no. 2, p.
e1003180, 2013.
[89] A. L. Drexler et al., “Human IGF1 regulates midgut oxidative stress and epithelial
homeostasis to balance lifespan and Plasmodium falciparum resistance in Anopheles
stephensi,” PLoS Pathog., vol. 10, no. 6, p. e1004231, 2014.
[90] B. G. Lindberg et al., “Immunogenic and antioxidant effects of a pathogen-associated
prenyl pyrophosphate in Anopheles gambiae,” PLoS One, vol. 8, no. 8, p. e73868, 2013.
[91] A. C. Bahia et al., “The role of reactive oxygen species in Anopheles aquasalis response
to Plasmodium vivax infection,” PLoS One, vol. 8, no. 2, p. e57014, 2013.
[92] G. Dimopoulos, D. Seeley, A. Wolf, and F. C. Kafatos, “Malaria infection of the mosquito
Anopheles gambiae activates immune-responsive genes during critical transition stages of
the parasite life cycle,” EMBO J., vol. 17, no. 21, pp. 6115–6123, 1998.
[93] A. C. Bahia et al., “The JAK-STAT pathway controls Plasmodium vivax load in early
169 stages of Anopheles aquasalis infection,” PLoS Negl. Trop. Dis., vol. 5, no. 11, p. e1317,
2011.
[94] G. de A. Oliveira, J. Lieberman, and C. Barillas-Mury, “Epithelial nitration by a
peroxidase/NOX5 system mediates mosquito antiplasmodial immunity.,” Science, vol.
335, no. 6070, pp. 856–9, 2012.
[95] R. Tahar, C. Boudin, I. Thiery, and C. Bourgouin, “Immune response of Anopheles
gambiae to the early sporogonic stages of the human malaria parasite Plasmodium
falciparum,” EMBO J., vol. 21, no. 24, pp. 6673–6680, 2002.
[96] A. Molina-Cruz et al., “The human malaria parasite Pfs47 gene mediates evasion of the
mosquito immune system.,” Science, vol. 340, no. May, pp. 984–7, 2013.
[97] U. N. Ramphul, L. S. Garver, A. Molina-Cruz, G. E. Canepa, and C. Barillas-Mury,
“Plasmodium falciparum evades mosquito immunity by disrupting JNK-mediated
apoptosis of invaded midgut cells,” Proc. Natl. Acad. Sci., vol. 112, no. 5, pp. 1273–1280,
2015.
[98] J. Vizioli, P. Bulet, J. A. Hoffmann, F. C. Kafatos, H. M. Muller, and G. Dimopoulos,
“Gambicin: a novel immune responsive antimicrobial peptide from the malaria vector
Anopheles gambiae,” Proc. Natl. Acad. Sci. U.S.A., vol. 98, no. 22, pp. 12630–12635,
2001.
[99] A. N. Clements, The Biology of Mosquitoes, Volume 3 - Transmission of Viruses and
Interactions with Bacteria. Chapman & Hall, 2012.
[100] J. Vizioli et al., “Cloning and analysis of a cecropin gene from the malaria vector
170 mosquito, Anopheles gambiae,” Insect Mol. Biol., vol. 9, no. 1, pp. 75–84, 2000.
[101] G. Dimopoulos, A. Richman, H. M. Muller, and F. C. Kafatos, “Molecular immune
responses of the mosquito Anopheles gambiae to bacteria and malaria parasites,” Proc.
Natl. Acad. Sci. U.S.A., vol. 94, no. 21, pp. 11508–11513, 1997.
[102] S. Blandin, L. F. Moita, T. Kocher, M. Wilm, F. C. Kafatos, and E. A. Levashina,
“Reverse genetics in the mosquito Anopheles gambiae: targeted disruption of the Defensin
gene,” EMBO Rep., vol. 3, no. 9, pp. 852–856, 2002.
[103] W. Kim et al., “Ectopic expression of a cecropin transgene in the human malaria vector
mosquito Anopheles gambiae (Diptera: Culicidae): effects on susceptibility to
Plasmodium,” J. Med. Entomol., vol. 41, no. 3, pp. 447–455, 2004.
[104] L. Gupta et al., “Apolipophorin-III mediates antiplasmodial epithelial responses in
Anopheles gambiae (G3) mosquitoes,” PLoS One, vol. 5, no. 11, p. e15410, 2010.
[105] M. K. Rono, M. M. Whitten, M. Oulad-Abdelghani, E. A. Levashina, and E. Marois, “The
major yolk protein vitellogenin interferes with the anti-plasmodium response in the
malaria mosquito Anopheles gambiae,” PLoS Biol., vol. 8, no. 7, p. e1000434, 2010.
[106] S. L. Sandiford, Y. Dong, A. Pike, B. J. Blumberg, A. C. Bahia, and G. Dimopoulos,
“Cytoplasmic actin is an extracellular insect immune factor which is secreted upon
immune challenge and mediates phagocytosis and direct killing of bacteria, and Is a
Plasmodium Antagonist,” PLoS Pathog., vol. 11, no. 2, p. e1004631, 2015.
[107] Y. Dong, F. Manfredini, and G. Dimopoulos, “Implication of the mosquito midgut
microbiota in the defense against malaria parasites,” PLoS Pathog., vol. 5, no. 5, p.
171 e1000423, 2009.
[108] C. M. Cirimotich et al., “Natural Microbe-Mediated Refractoriness to Plasmodium
Infection in Anopheles gambiae,” Science (80-. )., vol. 332, no. May, pp. 855–858, 2011.
[109] A. Capone et al., “Interactions between Asaia, Plasmodium and Anopheles: new insights
into mosquito symbiosis and implications in malaria symbiotic control.,” Parasit. Vectors,
vol. 6, no. 1, p. 182, Jun. 2013.
[110] G. Minard, P. Mavingui, and C. V. Moro, “Diversity and function of bacterial microbiota
in the mosquito holobiont.,” Parasit. Vectors, vol. 6, no. 1, p. 146, May 2013.
[111] A. C. Bahia et al., “Exploring Anopheles gut bacteria for Plasmodium blocking activity,”
Environ. Microbiol., vol. 16, no. 9, pp. 2980–2994, Sep. 2014.
[112] S. Kumar, M.-C. Alvaro, L. Gupta, J. Rodrigues, and C. Barillas-Mury, “A
Peroxidase/Dual Oxidase System Modulates Midgut Epithelial Immunity in Anopheles
gambiae.,” Science (80-. )., vol. 327, no. 5973, pp. 1644–1648, 2010.
[113] B. Lemaitre and J. Hoffmann, “The host defense of Drosophila melanogaster,” Annu. Rev.
Immunol., vol. 25, pp. 697–743, 2007.
[114] Y. Dong, S. Das, C. Cirimotich, J. A. Souza-Neto, K. J. McLean, and G. Dimopoulos,
“Engineered anopheles immunity to Plasmodium infection,” PLoS Pathog., vol. 7, no. 12,
p. e1002458, 2011.
[115] J. L. Ramirez et al., “A mosquito lipoxin/lipocalin complex mediates innate immune
priming in Anopheles gambiae.,” Nat. Commun., vol. 6, no. May 2015, p. 7403, 2015.
[116] M. L. Simões and G. Dimopoulos, “A mosquito mediator of parasite-induced immune
172 priming,” Trends Parasitol., vol. 31, no. 9, pp. 402–404, 2015.
[117] G. Bian et al., “Wolbachia invades Anopheles stephensi populations and induces
refractoriness to Plasmodium infection,” Science (80-. )., vol. 340, no. 6133, pp. 748–751,
2013.
[118] Z. Kambris et al., “Wolbachia stimulates immune gene expression and inhibits
plasmodium development in anopheles gambiae,” PLoS Pathog., vol. 6, no. 10, p.
e1001143, 2010.
[119] G. L. Hughes, R. Koga, P. Xue, T. Fukatsu, and J. L. Rasgon, “Wolbachia infections are
virulent and inhibit the human malaria parasite Plasmodium falciparum in Anopheles
gambiae,” PLoS Pathog., vol. 7, no. 5, p. e1002043, 2011.
[120] D. J. Gubler, “The economic burden of dengue,” Am. J. Trop. Med. Hyg., vol. 86, no. 5,
pp. 743–744, 2012.
[121] T. Michel, J.-M. Reichhart, J. A. Hoffmann, and J. Royet, “Drosophila Toll is activated by
Gram-positive bacteria through a circulating peptidoglycan recognition protein,” Nature,
vol. 414, no. 6865, pp. 756–759, 2001.
[122] V. Bischoff, C. Vignal, I. G. Boneca, T. Michel, J. A. Hoffmann, and J. Royet, “Function
of the Drosophila pattern-recognition receptor PGRP-SD in the detection of Gram-positive
bacteria,” Nat. Immunol., vol. 5, no. 11, pp. 1175–1180, 2004.
[123] Y. DeLotto and R. DeLotto, “Proteolytic processing of the Drosophila Spätzle protein by
Easter generates a dimeric NGF-like molecule with ventralising activity,” Mech. Dev., vol.
72, no. 1–2, pp. 141–148, 1998.
173 [124] E. Nicolas, J. M. Reichhart, J. A. Hoffmann, and B. Lemaitre, “In vivo regulation of the
IκB homologuecactus during the immune response of Drosophila,” J. Biol. Chem., vol.
273, no. 17, pp. 10463–10469, 1998.
[125] Z. Xi, J. L. Ramirez, and G. Dimopoulos, “The Aedes aegypti toll pathway controls
dengue virus infection,” PLoS Pathog., vol. 4, no. 7, p. e1000098, 2008.
[126] S. Sim, J. L. Ramirez, and G. Dimopoulos, “Dengue virus infection of the Aedes aegypti
salivary gland and chemosensory apparatus induces genes that modulate infection and
blood-feeding behavior,” PLoS Pathog., vol. 8, no. 3, p. e1002631, Mar. 2012.
[127] T. M. Colpitts et al., “Alterations in the Aedes aegypti transcriptome during Infection with
West Nile, Dengue and Yellow Fever viruses,” PLoS Pathog., vol. 7, no. 9, p. e1002189,
2011.
[128] H. R. Sanders et al., “Sindbis virus induces transport processes and alters expression of
innate immunity pathway genes in the midgut of the disease vector, Aedes aegypti,” Insect
Biochem. Mol. Biol., vol. 35, no. 11, pp. 1293–1307, 2005.
[129] N. Luplertlop et al., “Induction of a peptide with activity against a broad spectrum of
pathogens in the Aedes aegypti salivary gland, following Infection with Dengue virus,”
PLoS Pathog., vol. 7, no. 1, p. e1001252, 2011.
[130] S. Sim et al., “Transcriptomic Profiling of Diverse Aedes aegypti Strains Reveals
Increased Basal-level Immune Activation in Dengue Virus-refractory Populations and
Identifies Novel Virus-vector Molecular Interactions,” PLoS Negl. Trop. Dis., vol. 7, no.
7, p. e2295, Jul. 2013.
174 [131] S. Dupuis et al., “Impaired response to interferon-α/β and lethal viral disease in human
STAT1 deficiency,” Nat. Genet., vol. 33, no. 3, pp. 388–391, 2003.
[132] S. M. Karst, C. E. Wobus, M. Lay, J. Davidson, and H. W. Virgin, “STAT1-dependent
innate immunity to a Norwalk-like virus,” Science (80-. )., vol. 299, no. 5612, pp. 1575–
1578, 2003.
[133] N. I. Arbouzova and M. P. Zeidler, “JAK/STAT signalling in Drosophila: insights into
conserved regulatory and cellular functions,” Development, vol. 133, no. 14, pp. 2605–
2616, 2006.
[134] J. A. Souza-Neto, S. Sim, and G. Dimopoulos, “An evolutionary conserved function of the
JAK-STAT pathway in anti-dengue defense.,” Proc. Natl. Acad. Sci. U. S. A., vol. 106,
no. 42, pp. 17841–6, 2009.
[135] P. N. Paradkar, J.-B. Duchemin, R. Voysey, and P. J. Walker, “Dicer-2-dependent
activation of Culex Vago occurs via the TRAF-Rel2 signaling pathway,” PLoS Neglected
Trop. Dis., vol. 8, no. 4, p. e2823, 2014.
[136] P. N. Paradkar, L. Trinidad, R. Voysey, J.-B. Duchemin, and P. J. Walker, “Secreted Vago
restricts West Nile virus infection in Culex mosquito cells by activating the JAK-STAT
pathway,” Proc. Natl. Acad. Sci. U.S.A., vol. 109, no. 46, pp. 18915–18920, 2012.
[137] I. Sánchez-Vargas et al., “Dengue virus type 2 infections of Aedes aegypti are modulated
by the mosquito’s RNA interference pathway,” PLoS Pathog., vol. 5, no. 2, p. e1000299,
2009.
[138] J. M. Hardwick, “Viral interference with apoptosis,” Semin. Cell Dev. Biol., vol. 9, no. 3,
175 pp. 339–349, 1998.
[139] R. Vaidyanathan and T. Scott, “Apoptosis in mosquito midgut epithelia associated with
West Nile virus infection,” Apoptosis, vol. 11, no. 9, pp. 1643–1651, 2006.
[140] B. Liu et al., “P53-mediated rapid induction of apoptosis conveys resistance to viral
infection in Drosophila melanogaster,” PLoS Pathog., vol. 9, no. 2, p. e1003137, 2013.
[141] J. L. Ramirez et al., “Reciprocal tripartite interactions between the Aedes aegypti midgut
microbiota, innate immune system and dengue virus influences vector competence,” PLoS
Negl. Trop. Dis., vol. 6, no. 3, p. e1561, 2012.
[142] J. L. Ramirez et al., “Chromobacterium Csp_P reduces malaria and dengue infection in
vector mosquitoes and has entomopathogenic and in vitro anti-pathogen activities,” PLoS
Pathog., vol. 10, no. 10, p. e1004398, Oct. 2014.
[143] A. de O. Gaio, D. S. Gusmão, A. V Santos, M. A. Berbert-Molina, P. F. P. Pimenta, and F.
J. A. Lemos, “Contribution of midgut bacteria to blood digestion and egg production in
aedes aegypti (diptera: culicidae) (L.).,” Parasit. Vectors, vol. 4, no. 1, p. 105, 2011.
[144] D. Laurent and F. Pietra, “Antiplasmodial marine natural products in the perspective of
current chemotherapy and prevention of malaria. A review,” Marine Biotechnology, vol.
8, no. 5. pp. 433–447, 2006.
[145] World Health Organization, Guidelines for the treatment of malaria, Third edit. 2015.
[146] E. L. Dahl, J. L. Shock, B. R. Shenai, J. Gut, J. L. DeRisi, and P. J. Rosenthal,
“Tetracyclines specifically target the apicoplast of the malaria parasite Plasmodium
falciparum.,” Antimicrob. Agents Chemother., vol. 50, no. 9, pp. 3124–31, Sep. 2006.
176 [147] D. A. Leigh, “Antibacterial activity and pharmacokinetics of clindamycin,” J. Antimicrob.
Chemother., vol. 7, no. suppl A, pp. 3–9, Jan. 1981.
[148] H. J. Smith and M. M. Meremikwu, “Iron-chelating agents for treating malaria,” in
Cochrane Database of Systematic Reviews, H. J. Smith, Ed. Chichester, UK: John Wiley
& Sons, Ltd, 2003.
[149] J. F. Fernandes et al., “Fosmidomycin as an antimalarial drug: a meta-analysis of clinical
trials,” Future Microbiol., vol. 10, no. 8, pp. 1375–1390, Aug. 2015.
[150] J. Wiesner, D. Henschker, D. B. Hutchinson, E. Beck, and H. Jomaa, “In vitro and in vivo
synergy of fosmidomycin, a novel antimalarial drug, with clindamycin.,” Antimicrob.
Agents Chemother., vol. 46, no. 9, pp. 2889–94, Sep. 2002.
[151] P. Cos, A. J. Vlietinck, D. Vanden Berghe, and L. Maes, “Anti-infective potential of
natural products: How to develop a stronger in vitro ‘proof-of-concept,’” J.
Ethnopharmacol., vol. 106, no. 3, pp. 290–302, Jul. 2006.
[152] S. O. Sarr et al., “Icacina senegalensis (Icacinaceae), traditionally used for the treatment of
malaria, inhibits in vitro Plasmodium falciparum growth without host cell toxicity.,”
Malar. J., vol. 10, p. 85, Apr. 2011.
[153] W. C. Van Voorhis et al., “Open Source Drug Discovery with the Malaria Box Compound
Collection for Neglected Diseases and Beyond,” PLOS Pathog., vol. 12, no. 7, p.
e1005763, Jul. 2016.
[154] C. J. Schulze, G. Navarro, D. Ebert, J. DeRisi, and R. G. Linington, “Salinipostins A–K,
Long-Chain Bicyclic Phosphotriesters as a Potent and Selective Antimalarial Chemotype,”
177 J. Org. Chem., vol. 80, no. 3, pp. 1312–1320, Feb. 2015.
[155] V. Point et al., “Synthesis and Kinetic Evaluation of Cyclophostin and Cyclipostins
Phosphonate Analogs As Selective and Potent Inhibitors of Microbial Lipases,” J. Med.
Chem., vol. 55, no. 22, pp. 10204–10219, Nov. 2012.
[156] P. C. Nguyen et al., “Cyclipostins and Cyclophostin analogs as promising compounds in
the fight against tuberculosis,” Sci. Rep., vol. 7, no. 1, p. 11751, Dec. 2017.
[157] J. V Formica and M. J. Waring, “Effect of phosphate and amino acids on echinomycin
biosynthesis by Streptomyces echinatus.,” Antimicrob. Agents Chemother., vol. 24, no. 5,
pp. 735–41, Nov. 1983.
[158] H. Hayase et al., “Inhibition of malaria parasite growth by quinomycin A and its
derivatives through DNA-intercalating activity,” Biosci. Biotechnol. Biochem., vol. 79, no.
4, pp. 633–635, Apr. 2015.
[159] “Phase I trial of echinomycin (NSC 526417), a bifunctional intercalating agent,
administered by 24-hour continuous infusion,” Eur. J. Cancer Clin. Oncol., vol. 25, no. 5,
pp. 797–803, May 1989.
[160] M. C. Carnio, T. Stachelhaus, K. P. Francis, and S. Scherer, “Pyridinyl polythiazole class
peptide antibiotic micrococcin P1, secreted by foodborne Staphylococcus equorum
WS2733, is biosynthesized nonribosomally,” Eur. J. Biochem., vol. 268, no. 24, pp. 6390–
6400, Dec. 2001.
[161] M. J. Rogers, E. Cundliffe, and T. F. McCutchan, “The antibiotic micrococcin is a potent
inhibitor of growth and protein synthesis in the malaria parasite,” Antimicrob. Agents
178 Chemother., vol. 42, no. 3, pp. 715–716, Mar. 1998.
[162] G. Degiacomi et al., “Micrococcin P1 – A bactericidal thiopeptide active against
Mycobacterium tuberculosis,” Tuberculosis, vol. 100, pp. 95–101, Sep. 2016.
[163] G. B. MAHAJAN et al., “Use of a Thiopeptide Compound in the Treatment of
Clostridium Difficile Associated Infections,” 08-May-2015.
[164] C. Gumila, M. L. Ancelin, G. Jeminet, A. M. Delort, G. Miquel, and H. J. Vial,
“Differential in vitro activities of ionophore compounds against Plasmodium falciparum
and mammalian cells,” Antimicrob. Agents Chemother., vol. 40, no. 3, pp. 602–608, Mar.
1996.
[165] C. Gumila, M. L. Ancelin, A. M. Delort, G. Jeminet, and H. J. Vial, “Characterization of
the potent in vitro and in vivo antimalarial activities of ionophore compounds,”
Antimicrob. Agents Chemother., vol. 41, no. 3, pp. 523–529, Mar. 1997.
[166] D. Barbaras, M. Kaiser, R. Brun, and K. Gademann, “Potent and selective antiplasmodial
activity of the cyanobacterial alkaloid nostocarboline and its dimers,” 2008.
[167] P. G. Becher, H. I. Baumann, K. Gademann, and F. Jüttner, “The cyanobacterial alkaloid
nostocarboline: an inhibitor of acetylcholinesterase and trypsin,” J. Appl. Phycol., vol. 21,
no. 1, pp. 103–110, Feb. 2009.
[168] S. Bonazzi et al., “Antimalarial and antitubercular nostocarboline and eudistomin
derivatives: Synthesis, in vitro and in vivo biological evaluation,” Bioorg. Med. Chem.,
vol. 18, no. 4, pp. 1464–1476, 2010.
[169] “Mutasynthesis – uniting chemistry and genetics for drug discovery,” Trends Biotechnol.,
179 vol. 25, no. 4, pp. 139–142, Apr. 2007.
[170] W. Lu, N. Roongsawang, and T. Mahmud, “Biosynthetic studies and genetic engineering
of pactamycin analogs with improved selectivity toward malarial parasites,” Cell Press,
Apr. 2011.
[171] K. H. Almabruk, W. Lu, Y. Li, M. Abugreen, J. X. Kelly, and T. Mahmud,
“Mutasynthesis of fluorinated pactamycin analogues and their antimalarial activity,” Org.
Lett., vol. 15, no. 7, pp. 1678–1681, Apr. 2013.
[172] R. R. Teixeira et al., “Natural products as source of potential dengue antivirals,”
Molecules, vol. 19, no. 6. Multidisciplinary Digital Publishing Institute, pp. 8151–8176,
17-Jun-2014.
[173] T. Yoshimori, A. Yamamoto, Y. Moriyama, M. Futai, and Y. Tashiro, “Bafilomycin A1, a
specific inhibitor of vacuolar-type H(+)-ATPase, inhibits acidification and protein
degradation in lysosomes of cultured cells.,” J. Biol. Chem., vol. 266, no. 26, pp. 17707–
12, Sep. 1991.
[174] “Endocytic pathway followed by dengue virus to infect the mosquito cell line C6/36 HT,”
Virology, vol. 378, no. 1, pp. 193–199, Aug. 2008.
[175] S. Kang, A. R. Shields, N. Jupatanakul, and G. Dimopoulos, “Suppressing Dengue-2
infection by chemical inhibition of Aedes aegypti host factors,” PLoS Neglected Trop.
Dis., vol. 8, no. 8, p. e3084, Aug. 2014.
[176] D. Shum et al., “High-Content Assay to Identify Inhibitors of Dengue Virus Infection,”
Assay Drug Dev. Technol., vol. 8, no. 5, pp. 553–570, Oct. 2010.
180 [177] A. Raveh et al., “Discovery of Potent Broad Spectrum Antivirals Derived from Marine
Actinobacteria,” PLoS One, vol. 8, no. 12, p. e82318, Dec. 2013.
[178] F. Karray, E. Darbon, H. C. Nguyen, J. Gagnat, and J.-L. Pernodet, “Regulation of the
biosynthesis of the macrolide antibiotic spiramycin in Streptomyces ambofaciens.,” J.
Bacteriol., vol. 192, no. 21, pp. 5813–21, Nov. 2010.
[179] K. Naghmouchi, R. Hammami, I. Fliss, R. Teather, J. Baah, and D. Drider, “Colistin A
and colistin B among inhibitory substances of Paenibacillus polymyxa JB05-01-1,” Arch.
Microbiol., vol. 194, no. 5, pp. 363–370, May 2012.
[180] C. K. Lin et al., “Butyrolactones and Diketopiperazines from Marine Microbes: Inhibition
Effects on Dengue Virus Type 2 Replication,” Planta Med., vol. 83, no. 1–2, pp. 158–163,
Aug. 2017.
[181] M. Adamczeski, E. Quinoa, and P. Crews, “Novel sponge-derived amino acids. 5.
Structures, stereochemistry, and synthesis of several new heterocycles,” J. Am. Chem.
Soc., vol. 111, no. 2, pp. 647–654, Jan. 1989.
[182] A. D. Borthwick, “2,5-Diketopiperazines: Synthesis, Reactions, Medicinal Chemistry, and
Bioactive Natural Products,” Chem. Rev., vol. 112, no. 7, pp. 3641–3716, Jul. 2012.
[183] Z. Jiang, K. G. Boyd, A. Mearns-spragg, D. R. Adams, P. C. Wright, and J. G. Burgess,
“Two Diketopiperazines and One Halogenated Phenol from Cultures of the Marine
Bacterium, Pseudoalteromonas luteoviolacea,” Nat. Prod. Lett., vol. 14, no. 6, pp. 435–
440, Dec. 2000.
[184] A. Sajeli Begum et al., “Isolation and Characterization of Antimicrobial Cyclic Dipeptides
181 from Pseudomonas fluorescens and Their Efficacy on Sorghum Grain Mold Fungi,”
Chem. Biodivers., vol. 11, no. 1, pp. 92–100, Jan. 2014.
[185] C. B. Pumpuni, J. Demaio, M. Kent, J. R. Davis, and J. C. Beier, “Bacterial population
dynamics in three anopheline species: The impact on Plasmodium sporogonic
development,” Am. J. Trop. Med. Hyg., vol. 54, no. 2, pp. 214–218, Feb. 1996.
[186] S. Álvarez-Pérez, C. M. Herrera, and C. Vega, “Zooming-in on floral nectar: a first
exploration of nectar-associated bacteria in wild plant communities,” FEMS Microbiol.
Ecol., vol. 80, no. 3, pp. 591–602, Jun. 2012.
[187] Y. Wang, T. M. Gilbreath, P. Kukutla, G. Yan, and J. Xu, “Dynamic Gut Microbiome
across Life History of the Malaria Mosquito Anopheles gambiae in Kenya,” PLoS One,
vol. 6, no. 9, p. e24767, Sep. 2011.
[188] A. Boissière et al., “Midgut microbiota of the malaria mosquito vector Anopheles
gambiae and interactions with Plasmodium falciparum infection,” PLoS Pathog., vol. 8,
no. 5, p. 12, May 2012.
[189] C. J. Ngwa et al., “16S rRNA Gene-Based Identification of <I>Elizabethkingia
meningoseptica</I> (Flavobacteriales: Flavobacteriaceae) as a Dominant Midgut
Bacterium of the Asian Malaria Vector <I>Anopheles stephensi</I> (Dipteria:
Culicidae) With Antimicrobial Activities,” J. Med. Entomol., vol. 50, no. 2, pp. 404–414,
Mar. 2013.
[190] J. Osei-Poku, C. M. Mbogo, W. J. Palmer, and F. M. Jiggins, “Deep sequencing reveals
extensive variation in the gut microbiota of wild mosquitoes from Kenya,” Mol. Ecol., vol.
21, no. 20, pp. 5138–5150, Oct. 2012.
182 [191] K. L. Coon, K. J. Vogel, M. R. Brown, and M. R. Strand, “Mosquitoes rely on their gut
microbiota for development.,” Mol. Ecol., vol. 23, no. 11, pp. 2727–39, Jun. 2014.
[192] D. Duguma et al., “Developmental succession of the microbiome of Culex mosquitoes.,”
BMC Microbiol., vol. 15, p. 140, Jul. 2015.
[193] M. Buck, L. K. J. Nilsson, C. Brunius, R. K. Dabiré, R. Hopkins, and O. Terenius,
“Bacterial associations reveal spatial population dynamics in Anopheles gambiae
mosquitoes,” Sci. Rep., vol. 6, no. 1, p. 22806, Sep. 2016.
[194] N. Segata et al., “The reproductive tracts of two malaria vectors are populated by a core
microbiome and by gender- and swarm-enriched microbial biomarkers.,” Sci. Rep., vol. 6,
p. 24207, Apr. 2016.
[195] S. M. Short, E. F. Mongodin, H. J. MacLeod, O. A. C. Talyuli, and G. Dimopoulos,
“Amino acid metabolic signaling influences Aedes aegypti midgut microbiome
variability,” PLoS Negl. Trop. Dis., vol. 11, no. 7, p. e0005677, Jul. 2017.
[196] P. Azambuja, E. S. Garcia, and N. A. Ratcliffe, “Gut microbiota and parasite transmission
by insect vectors,” Trends Parasitol., vol. 21, no. 12, pp. 568–572, Dec. 2005.
[197] P. Azambuja, D. Feder, and E. S. Garcia, “Isolation of Serratia marcescens in the midgut
of Rhodnius prolixus: Impact on the establishment of the parasite Trypanosoma cruzi in
the vector,” Exp. Parasitol., vol. 107, no. 1–2, pp. 89–96, May 2004.
[198] C. Genes et al., “Mitochondrial dysfunction in Trypanosoma cruzi: the role of Serratia
marcescens prodigiosin in the alternative treatment of Chagas disease,” Parasit. Vectors,
vol. 4, no. 1, p. 66, 2011.
183 [199] C. M. Cirimotich et al., “Natural microbe-mediated rfractoriness to Plasmodium infection
in Anopheles gambiae,” Science (80-. )., vol. 332, no. 6031, pp. 855–858, 2011.
[200] C. Brady, I. Cleenwerck, S. Venter, T. Coutinho, and P. De Vos, “Taxonomic evaluation
of the genus Enterobacter based on multilocus sequence analysis (MLSA): Proposal to
reclassify E. nimipressuralis and E. amnigenus into Lelliottia gen. nov. as Lelliottia
nimipressuralis comb. nov. and Lelliottia amnigena comb. nov., ,” Syst. Appl. Microbiol.,
vol. 36, no. 5, pp. 309–319, Jul. 2013.
[201] B. A. Turturice et al., “Expression of Cytosolic Peroxiredoxins in Plasmodium berghei
Ookinetes Is Regulated by Environmental Factors in the Mosquito Bloodmeal,” Plos
Pathog., vol. 9, no. 1, p. 12, 2013.
[202] E. Jortzik and K. Becker, “Thioredoxin and glutathione systems in Plasmodium
falciparum,” Int. J. Med. Microbiol., vol. 302, no. 4–5, pp. 187–194, 2012.
[203] S. Muller, “Redox and antioxidant systems of the malaria parasite Plasmodium
falciparum,” Mol. Microbiol., vol. 53, no. 5, pp. 1291–1305, 2004.
[204] H. Sztajer et al., “The putative glutathione peroxidase gene of Plasmodium falciparum
codes for a thioredoxin peroxidase,” J. Biol. Chem., vol. 276, no. 10, pp. 7397–7403,
2001.
[205] N. Sturm et al., “Identification of Proteins Targeted by the Thioredoxin Superfamily in
Plasmodium falciparum,” Plos Pathog., vol. 5, no. 4, p. 12, 2009.
[206] K. Komaki-Yasuda, S. Kawazu, and S. Kano, “Disruption of the Plasmodium falciparum
2-Cys peroxiredoxin gene renders parasites hypersensitive to reactive oxygen and nitrogen
184 species,” Febs Lett., vol. 547, no. 1–3, pp. 140–144, 2003.
[207] K. J. Haselton et al., “Molecular cloning, characterization and expression profile of a
glutathione peroxidase-like thioredoxin peroxidase (TPxGl) of the rodent malaria parasite
Plasmodium berghei,” Parsitology Int., vol. http://dx., 2014.
[208] R. Munigunti, S. Gathiaka, O. Acevedo, R. Sahu, B. Tekwani, and A. I. Calderon,
“Characterization of PfTrxR inhibitors using antimalarial assays and in silico techniques,”
Chem. Cent. J., vol. 7, p. 7, 2013.
[209] R. E. Gary and W. A. Foster, “Effects of available sugar on the reproductive fitness and
vectorial capacity of the malaria vector Anopheles gambiae (Diptera : Culicidae),” J. Med.
Entomol., vol. 38, no. 1, pp. 22–28, 2001.
[210] G. Favia, G. Dimopoulos, A. Dellatorre, Y. T. Toure, M. Coluzzi, and C. Louis,
“Polymorphisms detected by random PCR distinguish between different chromosomal
forms of Anopheles gambiae ,” Proc. Natl. Acad. Sci. U. S. A., vol. 91, no. 22, pp. 10315–
10319, 1994.
[211] M. W. Pfaffl, “A new mathematical model for relative quantification in real-time RT-
PCR,” Nucleic Acids Res., vol. 29, no. 9, p. e45, 2001.
[212] S. V Angiuoli et al., “CloVR: A virtual machine for automated and portable sequence
analysis from the desktop using cloud computing,” BMC Bioinformatics, vol. 12, no. 1, p.
356, 2011.
[213] R. Aguilar et al., “Genome-wide analysis of transcriptomic divergence between laboratory
colony and field Anopheles gambiae mosquitoes of the M and S molecular forms,” Insect
185 Mol. Biol., vol. 19, no. 5, pp. 695–705, 2010.
[214] J. Herrero et al., “GEPAS: A web-based resource for microarray gene expression data
analysis,” Nucleic Acids Res., vol. 31, no. 13, pp. 3461–3467, 2003.
[215] S. Dudoit, R. C. Gentleman, and J. Quackenbush, “Open source software for the analysis
of microarray data,” Biotechniques, vol. 34, no. 3 SUPPL., pp. 45–51, 2003.
[216] I. V Yang et al., “Within the fold: assessing differential expression measures and
reproducibility in microarray assays.,” Genome Biol., vol. 3, no. 11, p. research0062,
2002.
[217] A. Rani, A. Sharma, R. Rajagopal, T. Adak, and R. K. Bhatnagar, “Bacterial diversity
analysis of larvae and adult midgut microflora using culture-dependent and culture-
independent methods in lab-reared and field-collected Anopheles stephensi-an Asian
malarial vector,” Bmc Microbiol., vol. 9, p. 22, 2009.
[218] J. K. Nayar and Vanhande.E, “Fuel for sustained mosquito flight,” J. Insect Physiol., vol.
17, no. 3, p. 471-, 1971.
[219] W. A. Foster, “Mosquito sugar feeding and reporoductive energetics,” Annu. Rev.
Entomol., vol. 40, pp. 443–474, 1995.
[220] M. L. Quinones, J. D. Lines, M. C. Thomson, M. Jawara, J. Morris, and B. M.
Greenwood, “Anopheles gambiae gonotrophic cycle duration, biting and exiting
behaviour unaffected by permethrin-impregnated bednets in The Gambia,” Med Vet
Entomol, vol. 11, no. 1, pp. 71–78, 1997.
[221] R. Morrow and W. Moss, “The Epidemiology and Control of Malaria,” in Infectious
186 Disease Epidemiology, 3rd ed., K. E. Nelson and C. Williams, Eds. Burlington, MA,
USA: Jones & Bartlett Learning, 2014.
[222] G. C. Muller et al., “Successful field trial of attractive toxic sugar bait (ATSB) plant-
spraying methods against malaria vectors in the Anopheles gambiae complex in Mali,
West Africa,” Malar. J., vol. 9, p. 7, 2010.
[223] L. C. Gouagna, B. Mulder, E. Noubissi, T. Tchuinkam, J. P. Verhave, and C. Boudin,
“The early sporogonic cycle of Plasmodium falciparum in laboratory-infected Anopheles
gambiae: an estimation of parasite efficacy,” Trop. Med. Int. Heal., vol. 3, no. 1, pp. 21–
28, 1998.
[224] M. A. Riehle, C. K. Moreira, D. Lampe, C. Lauzon, and M. Jacobs-Lorena, “Using
bacteria to express and display anti-Plasmodium molecules in the mosquito midgut,” Int.
J. Parasitol., vol. 37, no. 6, pp. 595–603, 2007.
[225] C. J. Hueck, “Type III protein secretion systems in bacterial pathogens of animals and
plants,” Microbiol Mol Biol Rev, vol. 62, no. 2, pp. 379–433, 1998.
[226] N. K. Fennelly et al., “Bordetella pertussis expresses a functional type III secretion system
that subverts protective innate and adaptive immune responses,” Infect Immun, vol. 76, no.
3, pp. 1257–1266, 2008.
[227] V. R. Correa et al., “The Bacterium Pantoea stewartii Uses Two Different Type III
Secretion Systems To Colonize Its Plant Host and Insect Vector,” Appl. Environ.
Microbiol., vol. 78, no. 17, pp. 6327–6336, 2012.
[228] A. Polissi et al., “Large-scale identification of virulence genes from Streptococcus
187 pneumoniae,” Infect Immun, vol. 66, no. 12, pp. 5620–5629, 1998.
[229] F. Song et al., “A multicomponent sugar phosphate sensor system specifically induced in
Bacillus cereus during infection of the insect gut,” Faseb j, vol. 26, no. 8, pp. 3336–3350,
2012.
[230] S. W. Ryter and R. M. Tyrrell, “The Heme synthesis and degradation pathways: role in
oxidant sensitivity - Heme oxygenase has both pro- and antioxidant properties,” Free
Radic. Biol. Med., vol. 28, no. 2, pp. 289–309, 2000.
[231] J. H. M. Oliveira et al., “Blood Meal-Derived Heme Decreases ROS Levels in the Midgut
of Aedes aegypti and Allows Proliferation of Intestinal Microbiota,” Plos Pathog., vol. 7,
no. 3, p. 14, 2011.
[232] J. B. Bliska, M. C. Copass, and S. Falkow, “The Yersinia pseudotuberculosis adhesin
YadA mediates intimate bacterial attachment to and entry into HEp-2 cells,” Infect.
Immun., vol. 61, no. 9, pp. 3914–3921, 1993.
[233] J. C. Leo et al., “First analysis of a bacterial collagen-binding protein with collagen
Toolkits: promiscuous binding of YadA to collagens may explain how YadA interferes
with host processes,” Infect Immun, vol. 78, no. 7, pp. 3226–3236, 2010.
[234] C. Blanco, “Transcriptional and translational signals of the uidA gene in Escherichia coli
K12,” Mol Gen Genet, vol. 208, no. 3, pp. 490–498, 1987.
[235] M. Novel and G. Novel, “Regulation of beta-glucuronidase synthesis in Escherichia coli
K-12: pleiotropic constitutive mutations affecting uxu and uidA expression,” J Bacteriol,
vol. 127, no. 1, pp. 418–432, 1976.
188 [236] K. Poole, “Bacterial stress responses as determinants of antimicrobial resistance,” J.
Antimicrob. Chemother., vol. 67, no. 9, pp. 2069–2089, 2012.
[237] G. Schwarz, R. R. Mendel, and M. W. Ribbe, “Molybdenum cofactors, enzymes and
pathways,” Nature, vol. 460, no. 7257, pp. 839–847, 2009.
[238] F. Foor and G. M. Brown, “Purification and properties of guanosine triphosphate
cyclohydrolase II from Eschericia coli,” J. Biol. Chem., vol. 250, no. 9, pp. 3545–3551,
1975.
[239] J. Green and M. S. Paget, “Bacterial redox sensors,” Nat. Rev. Microbiol., vol. 2, no. 12,
pp. 954–966, 2004.
[240] F. J. Peterson, R. P. Mason, J. Hovsepian, and J. L. Holtzman, “Oxygen-sensitive and
oxygen-insensitive nitroreduction by Escherichia coli and rat hepatic microsomes,” J.
Biol. Chem., vol. 254, no. 10, pp. 4009–4014, 1979.
[241] B. S. Hall, C. Bot, and S. R. Wilkinson, “Nifurtimox activation by trypanosomal type I
nitroreductases generates cytotoxic nitrile metabolites,” J Biol Chem, vol. 286, no. 15, pp.
13088–13095, 2011.
[242] M. Watanabe, T. Nishino, K. Takio, T. Sofuni, and T. Nohmi, “Purification and
characterization of wild-type and mutant ‘Classical’ nitroreductases of Salmonella
typhimurium - L33R mutation greatly diminishes binding of FMN to the nitroreductase of
S. typhimurium,” J. Biol. Chem., vol. 273, no. 37, pp. 23922–23928, 1998.
[243] D. Njus, P. M. Kelley, G. J. Harnadek, and Y. V Pacquing, “Mechanisms of ascorbic acid
regeneration emdiate by cytochrome b561 ,” Ann. N. Y. Acad. Sci., vol. 493, pp. 108–119,
189 1987.
[244] I. G. Young and F. Gibson, “Regulation of the enzymes involved in the biosynthesis of
2,3-dihydroxybenzoic acid in Aerobacter aerogenes and Escherichia coli,” Biochim.
Biophys. Acta - Gen. Subj., vol. 177, no. 3, pp. 401–411, May 1969.
[245] M. A. Skinnider et al., “Genomes to natural products PRediction Informatics for
Secondary Metabolomes (PRISM),” Nucleic Acids Res., vol. 43, no. 20, p. gkv1012, Oct.
2015.
[246] R. J. Abergel, A. M. Zawadzka, T. M. Hoette, and K. N. Raymond, “Enzymatic hydrolysis
of trilactone siderophores: where chiral recognition occurs in enterobactin and
bacillibactin iron transport.,” J. Am. Chem. Soc., vol. 131, no. 35, pp. 12682–92, Sep.
2009.
[247] T. Madden, “The BLAST Sequence Analysis Tool,” 2013.
[248] D. Galaris and K. Pantopoulos, “Oxidative stress and iron homeostasis: mechanistic and
health aspects.,” Crit. Rev. Clin. Lab. Sci., vol. 45, no. 1, pp. 1–23, 2008.
[249] N. Bresgen and P. M. Eckl, “Oxidative stress and the homeodynamics of iron
metabolism.,” Biomolecules, vol. 5, no. 2, pp. 808–47, 2015.
[250] A. Rotheneder et al., “Effects of synthetic siderophores on proliferation of Plasmodium
falciparum in infected human erythrocytes.,” Antimicrob. Agents Chemother., vol. 46, no.
6, pp. 2010–3, Jun. 2002.
[251] B. Pradines et al., “Iron chelators as antimalarial agents: in vitro activity of dicatecholate
against Plasmodium falciparum.,” J. Antimicrob. Chemother., vol. 50, no. 2, pp. 177–87,
190 Aug. 2002.
[252] K. Yano, K. Komaki-Yasuda, T. Tsuboi, M. Torii, S. Kana, and S. Kawazu, “2-Cys
peroxiredoxin TPx-1 is involved in gametocyte development in Plasmodium berghei,”
Mol. Biochem. Parasitol., vol. 148, no. 1, pp. 44–51, 2006.
[253] K. Yano et al., “Disruption of the Plasmodium berghei 2-Cys peroxiredoxin TPx-1 gene
hinders the sporozoite development in the vector mosquito,” Mol. Biochem. Parasitol.,
vol. 159, no. 2, pp. 142–145, 2008.
[254] S. Kawazu, N. Ikenoue, H. Takemae, K. Komaki-Yasuda, and S. Kano, “Roles of 1-Cys
peroxiredoxin in haem detoxification in the human malaria parasite Plasmodium
falciparum,” Febs J., vol. 272, no. 7, pp. 1784–1791, 2005.
[255] J. M. Lindh, A. K. Borg-Karlson, and I. Faye, “Transstadial and horizontal transfer of
bacteria within a colony of Anopheles gambiae (Diptera: Culicidae) and oviposition
response to bacteria-containing water,” Acta Trop., vol. 107, no. 3, pp. 242–250, 2008.
[256] R. M. Moll, W. S. Romoser, M. C. Modrzakowski, A. C. Moncayo, and K. Lerdthusnee,
“Meconial peritrophic membranes and the fate of midgut bacteria during mosquito
(Diptera : culicidae) metamorphosis,” J. Med. Entomol., vol. 38, no. 1, pp. 29–32, 2001.
[257] Z. Y. Xi, C. C. H. Khoo, and S. L. Dobson, “Wolbachia establishment and invasion in an
Aedes aegypti laboratory population,” Science (80-. )., vol. 310, no. 5746, pp. 326–328,
2005.
[258] G. Bian et al., “Wolbachia Invades Anopheles stephensi,” Science (80-. )., vol. 340, no.
May, pp. 748–751, 2013.
191 [259] B. Chouaia et al., “Delayed larval development in Anopheles mosquitoes deprived of
Asaia bacterial symbionts,” Bmc Microbiol., vol. 12, p. 8, 2012.
[260] M. Gillis and N. A. Logan, “Chromobacterium,” in Bergey’s Manual of Systematics of
Archaea and Bacteria, Chichester, UK: John Wiley & Sons, Ltd, 2015, pp. 1–9.
[261] P. A. W. Martin, D. Gundersen-Rindal, M. Blackburn, and J. Buyer, “Chromobacterium
subtsugae sp. nov., a betaproteobacterium toxic to Colorado potato beetle and other insect
pests,” Int. J. Syst. Evol. Microbiol., vol. 57, no. 5, pp. 993–999, May 2007.
[262] C.-C. Young et al., “Chromobacterium aquaticum sp. nov., isolated from spring water
samples.,” Int. J. Syst. Evol. Microbiol., vol. 58, no. 2008, pp. 877–880, Apr. 2008.
[263] X. Y. Han, F. S. Han, and J. Segal, “Chromobacterium haemolyticum sp. nov., a strongly
haemolytic species,” Int. J. Syst. Evol. Microbiol., vol. 58, no. 6, pp. 1398–1403, Jun.
2008.
[264] P. Kämpfer, H. J. Busse, and H. C. Scholz, “Chromobacterium piscinae sp. nov. and
Chromobacterium pseudoviolaceum sp. nov., from environmental samples,” Int. J. Syst.
Evol. Microbiol., vol. 59, no. 10, pp. 2486–2490, Oct. 2009.
[265] S. D. Soby, S. R. Gadagkar, C. Contreras, and F. L. Caruso, “Chromobacterium vaccinii
sp. nov., isolated from native and cultivated cranberry (Vaccinium macrocarpon Ait.) bogs
and irrigation ponds,” Int. J. Syst. Evol. Microbiol., vol. 63, no. PART 5, pp. 1840–1846,
May 2013.
[266] M. B. Blackburn, R. R. Farrar, M. E. Sparks, D. Kuhar, A. Mitchell, and D. E. Gundersen-
Rindal, “Chromobacterium sphagni sp. nov., an insecticidal bacterium isolated from
192 Sphagnum bogs,” Int. J. Syst. Evol. Microbiol., vol. 67, no. 9, pp. 3417–3422, 2017.
[267] W. Trager and J. B. Jensen, “Human malaria parasites in continuous culture,” J.
Parasitol., vol. 91, no. 3, pp. 484–486, 2005.
[268] T. N. Bennett et al., “Novel, rapid, and inexpensive cell-based quantification of
antimalarial drug efficacy,” Antimicrob. Agents Chemother., vol. 48, no. 5, pp. 1807–
1810, 2004.
[269] P. Ferrer, A. K. Tripathi, M. A. Clark, C. C. Hand, H. Y. Rienhoff, and D. J. Sullivan,
“Antimalarial iron chelator, FBS0701, shows asexual and gametocyte Plasmodium
falciparum activity and single oral dose cure in a murine malaria model,” PLoS One, vol.
7, no. 5, p. e37171, May 2012.
[270] C. Lambros and J. P. Vanderberg, “Synchronization of Plasmodium falciparum
erythrocytic stages in culture,” J. Parasitol., vol. 65, no. 3, p. 418, Jun. 1979.
[271] M. Adamek, M. Spohn, E. Stegmann, and N. Ziemert, “Mining bacterial genomes for
secondary metabolite gene clusters,” in Methods in Molecular Biology, vol. 1520,
Humana Press, New York, NY, 2017, pp. 23–47.
[272] E. Bosi et al., “MeDuSa: A multi-draft based scaffolder,” Bioinformatics, vol. 31, no. 15,
pp. 2443–2451, Aug. 2015.
[273] S. H. Yoon, S. min Ha, J. Lim, S. Kwon, and J. Chun, “A large-scale evaluation of
algorithms to calculate average nucleotide identity,” Antonie van Leeuwenhoek,
International Journal of General and Molecular Microbiology, pp. 1–6, 15-Feb-2017.
[274] S. Garcia-Vallvé, J. Palau, and A. Romeu, “Horizontal gene transfer in glycosyl
193 hydrolases inferred from codon usage in Escherichia coli and Bacillus subtilis,” Mol. Biol.
Evol., vol. 16, no. 9, pp. 1125–34, Sep. 1999.
[275] M. H. Medema et al., “AntiSMASH: Rapid identification, annotation and analysis of
secondary metabolite biosynthesis gene clusters in bacterial and fungal genome
sequences,” Nucleic Acids Res., vol. 39, no. SUPPL. 2, pp. W339–W346, Jul. 2011.
[276] T. Weber et al., “antiSMASH 3.0-a comprehensive resource for the genome mining of
biosynthetic gene clusters.,” Nucleic Acids Res., vol. 43, no. W1, pp. W237-43, Jul. 2015.
[277] P. Cimermancic et al., “Insights into secondary metabolism from a global analysis of
prokaryotic biosynthetic gene clusters,” Cell, vol. 158, no. 2, pp. 412–421, Jul. 2014.
[278] A. Morgulis, G. Coulouris, Y. Raytselis, T. L. Madden, R. Agarwala, and A. A. Schäffer,
“Database indexing for production MegaBLAST searches,” in Bioinformatics, 2008, vol.
24, no. 16, pp. 1757–1764.
[279] Y. Q. Cheng, M. Yang, and A. M. Matter, “Characterization of a gene cluster responsible
for the biosynthesis of anticancer agent FK228 in Chromobacterium violaceum No. 968,”
Appl. Environ. Microbiol., vol. 73, no. 11, pp. 3460–3469, 2007.
[280] V. Y. Potharla, S. R. Wesener, and Y.-Q. Cheng, “New insights into the genetic
organization of the FK228 biosynthetic gene cluster in Chromobacterium violaceum no.
968.,” Appl. Environ. Microbiol., vol. 77, no. 4, pp. 1508–11, Feb. 2011.
[281] D. Hyatt, G.-L. Chen, P. F. Locascio, M. L. Land, F. W. Larimer, and L. J. Hauser,
“Prodigal: prokaryotic gene recognition and translation initiation site identification.,”
BMC Bioinformatics, vol. 11, p. 119, Mar. 2010.
194 [282] K. Tamura, G. Stecher, D. Peterson, A. Filipski, and S. Kumar, “MEGA6: Molecular
Evolutionary Genetics Analysis version 6.0.,” Mol. Biol. Evol., vol. 30, no. 12, pp. 2725–
9, Dec. 2013.
[283] J. D. Thompson, D. G. Higgins, and T. J. Gibson, “CLUSTAL W: improving the
sensitivity of progressive multiple sequence alignment through sequence weighting,
position-specific gap penalties and weight matrix choice.,” Nucleic Acids Res., vol. 22, no.
22, pp. 4673–80, Nov. 1994.
[284] A. Seath and R. . Sokal, “Numerical Taxonomy. The Principles and Practice of Numerical
Classification,” Systamatic Zoology, vol. 24, no. 2. W.H. Freeman, pp. 263–268, 1973.
[285] K. Tamura, M. Nei, and S. Kumar, “Prospects for inferring very large phylogenies by
using the neighbor-joining method.,” Proc. Natl. Acad. Sci. U. S. A., vol. 101, no. 30, pp.
11030–5, Jul. 2004.
[286] T. S. Voss, T. Mini, P. Jenoe, and H.-P. Beck, “Plasmodium falciparum possesses a cell
cycle-regulated short type replication protein A large subunit encoded by an unusual
transcript.,” J. Biol. Chem., vol. 277, no. 20, pp. 17493–501, May 2002.
[287] G. Houen et al., “The Solubility of Proteins in Organic Solvents.,” Acta Chem. Scand.,
vol. 50, pp. 68–70, 1996.
[288] C. Nick Pace, S. Trevino, E. Prabhakaran, and J. Martin Scholtz, “Protein structure,
stability and solubility in water and other solvents,” Philos. Trans. R. Soc. B Biol. Sci.,
vol. 359, no. 1448, pp. 1225–1235, Aug. 2004.
[289] N. SHIGEMATSU, H. UEDA, S. TAKASE, H. TANAKA, K. YAMAMOTO, and T.
195 TADA, “FR901228, a novel antitumor bicyclic depsipeptide produced by
Chromobacterium violaceum No. 968. II. Structure determination.,” J. Antibiot. (Tokyo).,
vol. 47, no. 3, pp. 311–314, 1994.
[290] J. J. Xiao, J. Byrd, G. Marcucci, M. Grever, and K. K. Chan, “Identification of thiols and
glutathione conjugates of depsipeptide FK228 (FR901228), a novel histone protein
deacetylase inhibitor, in the blood,” Rapid Commun. Mass Spectrom., vol. 17, no. 8, pp.
757–766, Apr. 2003.
[291] G. L. Challis, J. Ravel, and C. A. Townsend, “Predictive, structure-based model of amino
acid recognition by nonribosomal peptide synthetase adenylation domains,” Chem. Biol.,
vol. 7, no. 3, pp. 211–224, 2000.
[292] T. Stachelhaus, H. D. Mootz, and M. A. Marahiel, “The specificity-conferring code of
adenylation domains in nonribosomal peptide synthetases,” Chem. Biol., vol. 6, no. 8, pp.
493–505, 1999.
[293] M. Rottig, M. H. Medema, K. Blin, T. Weber, C. Rausch, and O. Kohlbacher,
“NRPSpredictor2—a web server for predicting NRPS adenylation domain specificity,”
Nucleic Acids Res., vol. 39, no. Web Server, pp. W362–W367, 2011.
[294] C. Rausch, I. Hoof, T. Weber, W. Wohlleben, and D. H. Huson, “Phylogenetic analysis of
condensation domains in NRPS sheds light on their functional evolution.,” BMC Evol.
Biol., vol. 7, p. 78, 2007.
[295] A. Afonso, F. Hon, and R. Brambilla, “Structure elucidation of Sch 20562, a glucosidic
cyclic dehydropeptide lactone--the major component of W-10 antifungal antibiotic.,” J.
Antibiot. (Tokyo)., vol. 52, no. 4, pp. 383–97, Apr. 1999.
196 [296] A. Afonso, F. Hon, and R. Brambilla, “Structure Elucidation of Sch 20561, a Cyclic
Dehydropeptide Lactone,” J. Antibiot., vol. 52, no. 4, pp. 398–406, Apr. 2015.
[297] R. Furumai et al., “FK228 (depsipeptide) as a natural prodrug that inhibits class I histone
deacetylases,” Cancer Res., vol. 62, no. 17, pp. 4916–4921, 2002.
[298] K. T. Konstantinidis and J. M. Tiedje, “Genomic insights that advance the species
definition for prokaryotes,” Proc. Natl. Acad. Sci., vol. 102, no. 7, pp. 2567–2572, Feb.
2005.
[299] J. Goris, K. T. Konstantinidis, J. A. Klappenbach, T. Coenye, P. Vandamme, and J. M.
Tiedje, “DNA-DNA hybridization values and their relationship to whole-genome
sequence similarities,” Int. J. Syst. Evol. Microbiol., vol. 57, no. 1, pp. 81–91, Jan. 2007.
[300] C. Fuqua and E. P. Greenberg, “Signalling: Listening in on bacteria: acyl-homoserine
lactone signalling,” Nat. Rev. Mol. Cell Biol., vol. 3, no. 9, pp. 685–695, Sep. 2002.
[301] M. Schuster, D. Joseph Sexton, S. P. Diggle, and E. Peter Greenberg, “Acyl-Homoserine
Lactone Quorum Sensing: From Evolution to Application,” Annu. Rev. Microbiol., vol.
67, no. 1, pp. 43–63, Sep. 2013.
[302] H. Ueda, H. Nakajima, Y. Hori, T. Goto, and M. Okuhara, “Action of FR901228, a novel
antitumor bicyclic depsipeptide produced by Chromobacterium violaceum no. 968, on Ha-
ras transformed NIH3T3 cells,” Biosci. Biotechnol. Biochem., vol. 58, no. 9, pp. 1579–
1583, Jan. 1994.
[303] H. Nakajima, Y. B. Kim, H. Terano, M. Yoshida, and S. Horinouchi, “FR901228, a potent
antitumor antibiotic, is a novel histone deacetylase inhibitor,” Exp Cell Res, vol. 241, no.
197 1, pp. 126–133, 1998.
[304] K. M. VanderMolen, W. McCulloch, C. J. Pearce, and N. H. Oberlies, “Romidepsin
(Istodax, NSC 630176, FR901228, FK228, depsipeptide): a natural product recently
approved for cutaneous T-cell lymphoma,” J. Antibiot. (Tokyo)., vol. 64, no. 8, pp. 525–
531, Aug. 2011.
[305] E. M. Bertino and G. A. Otterson, “Romidepsin: a novel histone deacetylase inhibitor for
cancer,” Expert Opin. Investig. Drugs, vol. 20, no. 8, pp. 1151–1158, Aug. 2011.
[306] P. Smolewski and T. Robak, “The discovery and development of romidepsin for the
treatment of T-cell lymphoma,” Expert Opin. Drug Discov., pp. 1–15, Jun. 2017.
[307] M. J. Smilkstein et al., “A drug-selected Plasmodium falciparum lacking the need for
conventional electron transport,” Mol. Biochem. Parasitol., vol. 159, no. 1, pp. 64–68,
May 2008.
[308] K. T. Andrews, T. N. Tran, N. C. Wheatley, and D. P. Fairlie, “Targeting histone
deacetylase inhibitors for anti-malarial therapy.,” Curr. Top. Med. Chem., vol. 9, no. 3, pp.
292–308, 2009.
[309] J. A. Engel et al., “Profiling the anti-protozoal activity of anti-cancer HDAC inhibitors
against Plasmodium and Trypanosoma parasites.,” Int. J. Parasitol. Drugs drug Resist.,
vol. 5, no. 3, pp. 117–26, Dec. 2015.
[310] M. J. Chua et al., “Effect of clinically approved HDAC inhibitors on Plasmodium,
Leishmania and Schistosoma parasite growth.,” Int. J. Parasitol. Drugs drug Resist., vol.
7, no. 1, pp. 42–50, Apr. 2017.
198 [311] S. Das, L. Garver, J. R. Ramirez, Z. Xi, and G. Dimopoulos, “Protocol for dengue
infections in mosquitoes (A. aegypti) and infection phenotype determination.,” J. Vis.
Exp., no. 5, p. 220, Jul. 2007.
[312] G. Fibriansah et al., “A potent anti-dengue human antibody preferentially recognizes the
conformation of E protein monomers assembled on the virus surface.,” EMBO Mol. Med.,
vol. 6, no. 3, pp. 358–71, 2014.
[313] J. L. Tan and S. M. Lok, “Dengue Virus Purification and Sample Preparation for Cryo-
Electron Microscopy,” Humana Press, New York, NY, 2014, pp. 41–52.
[314] P. Vervaeke, M. Alen, S. Noppen, D. Schols, P. Oreste, and S. Liekens, “Sulfated
Escherichia coli K5 Polysaccharide Derivatives Inhibit Dengue Virus Infection of Human
Microvascular Endothelial Cells by Interacting with the Viral Envelope Protein E Domain
III,” PLoS One, vol. 8, no. 8, p. e74035, Aug. 2013.
[315] A. Pike, A. Vadlamani, S. L. Sandiford, A. Gacita, and G. Dimopoulos, “Characterization
of the Rel2-regulated transcriptome and proteome of Anopheles stephensi identifies new
anti-Plasmodium factors.,” Insect Biochem. Mol. Biol., vol. 52, pp. 82–93, Sep. 2014.
[316] V. Haridas et al., “Bispidine-Amino Acid Conjugates Act as a Novel Scaffold for the
Design of Antivirals That Block Japanese Encephalitis Virus Replication,” PLoS Negl.
Trop. Dis., vol. 7, no. 1, p. e2005, Jan. 2013.
[317] R. R. Burgess, “Protein Precipitation Techniques,” Methods in Enzymology, vol. 463, no.
C. pp. 331–342, 2009.
[318] R. J. Kuhn et al., “Structure of dengue virus: implications for flavivirus organization,
199 maturation, and fusion.,” Cell, vol. 108, no. 5, pp. 717–25, Mar. 2002.
[319] G. Fibriansah et al., “Structural changes in dengue virus when exposed to a temperature of
37°C.,” J. Virol., vol. 87, no. 13, pp. 7585–92, Jul. 2013.
[320] F. X. Heinz and S. L. Allison, “Structures and mechanisms in flavivirus fusion.,” Adv.
Virus Res., vol. 55, pp. 231–69, 2000.
[321] J. M. Smit, B. Moesker, I. Rodenhuis-Zybert, and J. Wilschut, “Flavivirus cell entry and
membrane fusion.,” Viruses, vol. 3, no. 2, pp. 160–71, Feb. 2011.
[322] N. D. Rawlings and A. J. Barrett, “Introduction: Metallopeptidases and Their Clans,” in
Handbook of Proteolytic Enzymes, Elsevier, 2013, pp. 325–370.
[323] E. Kessler and D. E. Ohman, “Pseudolysin,” in Handbook of Proteolytic Enzymes,
Elsevier, 2013, pp. 582–592.
[324] S. Miyoshi, K. Okamoto, and E. Takahashi, “Vibriolysin,” in Handbook of Proteolytic
Enzymes, Elsevier, 2013, pp. 579–582.
[325] B. Chevrier and H. D’Orchymont, “Vibrio Aminopeptidase,” in Handbook of Proteolytic
Enzymes, Elsevier, 2013, pp. 1627–1630.
[326] G. Pérez-Sánchez, L. I. Leal-Guadarrama, I. Trelles, N. O. Pérez, and E. Medina-Rivero,
“High-level production of a recombinant Vibrio proteolyticus leucine aminopeptidase and
its use for N-terminal methionine excision from interferon alpha-2b,” Process Biochem.,
vol. 46, no. 9, pp. 1825–1830, Sep. 2011.
[327] Z. Z. Zhang, S. Nirasawa, Y. Nakajima, M. Yoshida, and K. Hayashi, “Function of the N-
terminal propeptide of an aminopeptidase from Vibrio proteolyticus.,” Biochem. J., vol.
200 350 Pt 3, no. Pt 3, pp. 671–6, Sep. 2000.
[328] K. P. Bzymek, V. M. D’Souza, G. Chen, H. Campbell, A. Mitchell, and R. C. Holz,
“Function of the signal peptide and N- and C-terminal propeptides in the leucine
aminopeptidase from Aeromonas proteolytica,” Protein Expr. Purif., vol. 37, no. 2, pp.
294–305, Oct. 2004.
[329] M. Hartley, W. Yong, and B. Bennett, “Heterologous expression and purification of
Vibrio proteolyticus (Aeromonas proteolytica) aminopeptidase: a rapid protocol.,” Protein
Expr. Purif., vol. 66, no. 1, pp. 91–101, Jul. 2009.
[330] H. Sonoda, K. Daimon, H. Yamaji, and A. Sugimura, “Efficient production of active
Vibrio proteolyticus aminopeptidase in Escherichia coli by co-expression with engineered
vibriolysin,” Appl. Microbiol. Biotechnol., vol. 84, no. 1, pp. 191–198, Aug. 2009.
[331] R. Cahan, I. Axelrad, M. Safrin, D. E. Ohman, and E. Kessler, “A secreted
aminopeptidase of Pseudomonas aeruginosa. Identification, primary structure, and
relationship to other aminopeptidases.,” J. Biol. Chem., vol. 276, no. 47, pp. 43645–52,
Nov. 2001.
[332] K. Morihara and H. Tsuzuki, “Phosphoramidon as an inhibitor of elastase from
Pseudomonas aeruginosa.,” Jpn. J. Exp. Med., vol. 48, no. 1, pp. 81–4, Feb. 1978.
[333] V. Bultel-Poncé, J.-P. Berge, C. Debitus, J.-L. Nicolas, and M. Guyot, “Metabolites from
the Sponge-Associated Bacterium Pseudomonas Species,” Mar. Biotechnol., vol. 1, no. 4,
pp. 384–390, Jul. 1999.
[334] K. YAGISHITA, R. UTAHARA, K. MAEDA, M. HAMADA, and H. UMEZAWA,
201 “ACRYLAMIDINE, AN ANTI-FUNGAL SUBSTANCE PRODUCED BY A
STREPTOMYCES,” J. Antibiot. (Tokyo)., vol. 21, no. 7, pp. 444–450, Jul. 1968.
[335] K. Otoguro et al., “Promising lead compounds for novel antiprotozoals,” J. Antibiot.
(Tokyo)., vol. 63, no. 7, pp. 381–384, Jul. 2010.
[336] K. C. C. Cheng et al., “Actinoramide A Identified as a Potent Antimalarial from Titration-
Based Screening of Marine Natural Product Extracts,” J. Nat. Prod., vol. 78, no. 10, pp.
2411–2422, Oct. 2015.
[337] C. Portmann, J. F. Blom, M. Kaiser, R. Brun, F. Jüttner, and K. Gademann, “Isolation of
Aerucyclamides C and D and Structure Revision of Microcyclamide 7806A: Heterocyclic
Ribosomal Peptides from Microcystis aeruginosa PCC 7806 and Their Antiparasite
Evaluation,” J. Nat. Prod., vol. 71, no. 11, pp. 1891–1896, Dec. 2008.
[338] P. Gachon, C. Farges, and A. Kergomard, “Alborixin, a new antibiotic ionophore:
isolation, structure, physical and chemical properties.,” J. Antibiot. (Tokyo)., vol. 29, no. 6,
pp. 603–10, Jun. 1976.
[339] *,† Anthony D. Wright, ‡ and Olaf Papendorf, and G. M. König‡, “Ambigol C and 2,4-
Dichlorobenzoic Acid, Natural Products Produced by the Terrestrial Cyanobacterium
Fischerella ambigua,” 2005.
[340] S. Nakamura, K. Karasawa, H. Yonehara, N. Tanaka, and H. Umezawa, “New antibiotic,
amidinomycin [N-(2-amidinoethyl)-3aminocyclopentanecarboxamide) produced by a
Streptomyces,” J. Antibiot., vol. 14, no. Ser. A, pp. 103–106, 1961.
[341] M. Gutierrez, C. Boya, L. Herrera, and H. Guzman, “Antiplasmodial activity of
202 bacilosarcin A isolated from the octocoral-associated bacterium Bacillus sp. collected in
Panama,” J. Pharm. Bioallied Sci., vol. 4, no. 1, p. 66, 2012.
[342] M. Isaka, A. Jaturapat, J. Kramyu, M. Tanticharoen, and Y. Thebtaranonth, “Potent in
vitro antimalarial activity of metacycloprodigiosin isolated from Streptomyces spectabilis
BCC 4785,” Antimicrob. Agents Chemother., vol. 46, no. 4, pp. 1112–1113, Apr. 2002.
[343] C. Portmann et al., “Balgacyclamides, Antiplasmodial Heterocyclic Peptides from
Microcystis aeruguinosa EAWAG 251,” J. Nat. Prod., vol. 77, no. 3, pp. 557–562, Mar.
2014.
[344] C.-L. Shao et al., “Bastimolide A, a Potent Antimalarial Polyhydroxy Macrolide from the
Marine Cyanobacterium Okeania hirsuta,” J. Org. Chem., vol. 80, no. 16, pp. 7849–7855,
Aug. 2015.
[345] R. W. Rickards et al., “Calothrixins A and B, novel pentacyclic metabolites from
Calothrix cyanobacteria with potent activity against malaria parasites and human cancer
cells,” Tetrahedron, vol. 55, no. 47, pp. 13513–13520, 1999.
[346] *,† Kerry L. McPhail et al., “Antimalarial Linear Lipopeptides from a Panamanian Strain
of the Marine Cyanobacterium Lyngbya majuscula,” 2007.
[347] H. KINASHI, K. SOMENO, and K. SAKAGUCHI, “Isolation and characterization of
concanamycins A, B and C.,” J. Antibiot. (Tokyo)., vol. 37, no. 11, pp. 1333–1343, 1984.
[348] S. Auparakkitanon and P. Wilairat, “Antimalarial activity of concanamycin A alone and in
combination with pyronaridine,” Southeast Asian J. Trop. Med. Public Health, vol. 37, no.
4, pp. 619–621, Jul. 2006.
203 [349] K. H. Müller et al., “Inhibition by cellular vacuolar ATPase impairs human
papillomavirus uncoating and infection.,” Antimicrob. Agents Chemother., vol. 58, no. 5,
pp. 2905–11, May 2014.
[350] S. TAKAHASHI et al., “A New Antibiotic, Cyclamidomycin,” J. Antibiot. (Tokyo)., vol.
24, no. 12, pp. 902–903, 1971.
[351] D. E. Dahari, R. M. Salleh, F. Mahmud, L. P. Chin, N. Embi, and H. M. Sidek, “Anti-
malarial Activities of Two Soil Actinomycete Isolates from Sabah via Inhibition of
Glycogen Synthase Kinase 3β.,” Trop. life Sci. Res., vol. 27, no. 2, pp. 53–71, Aug. 2016.
[352] U. Castillo et al., “Kakadumycins, novel antibiotics from Streptomyces sp. NRRL 30566,
an endophyte of Grevillea pteridifolia,” FEMS Microbiol. Lett., vol. 224, no. 2, pp. 183–
190, Jul. 2003.
[353] R. G. Linington et al., “Antimalarial Peptides from Marine Cyanobacteria: Isolation and
Structural Elucidation of Gallinamide A,” J. Nat. Prod., vol. 72, no. 1, pp. 14–17, Jan.
2009.
[354] R. D. Hotchkiss, “Gramicidin, Tyrocidine, and Tyrothricin,” John Wiley & Sons, Inc.,
2006, pp. 153–199.
[355] P. Gachon and A. Kergomard, “Grisorixin, an ionophorous antibiotic of the nigericin
group. II. Chemical and structural study of grisorixin and some derivatives.,” J. Antibiot.
(Tokyo)., vol. 28, no. 5, pp. 351–7, May 1975.
[356] M. Kronenwerth, C. Dauth, M. Kaiser, I. Pemberton, and H. B. Bode, “Facile Synthesis of
Cyclohexanediones and Dialkylresorcinols - Bioactive Natural Products from
204 Entomopathogenic Bacteria,” European J. Org. Chem., vol. 2014, no. 36, pp. 8026–8028,
Dec. 2014.
[357] A. Tripathi, J. Puddick, M. R. Prinsep, M. Rottmann, and L. T. Tan, “Lagunamides A and
B: Cytotoxic and Antimalarial Cyclodepsipeptides from the Marine Cyanobacterium
Lyngbya majuscula,” J. Nat. Prod., vol. 73, no. 11, pp. 1810–1814, Nov. 2010.
[358] A. Tripathi et al., “Lagunamide C, a cytotoxic cyclodepsipeptide from the marine
cyanobacterium Lyngbya majuscula,” Phytochemistry, vol. 72, no. 18, pp. 2369–2375,
2011.
[359] J. W. WESTLEY et al., “BIOSYNTHESIS OF LASALOCID. I,” J. Antibiot. (Tokyo).,
vol. 27, no. 4, pp. 288–297, 1974.
[360] S. Omura, M. Shibata, S. Machida, and J. Sawada, “Isolation of a new polyether
antibiotic, lonomycin.,” J. Antibiot. (Tokyo)., vol. 29, no. 1, pp. 15–20, Jan. 1976.
[361] E. Ebata, H. Kasahara, K. Sekine, and Y. Inoue, “Lysocellin, a new polyether antibiotic. I.
Isolation, purification, physico-chemical and biological properties.,” J. Antibiot. (Tokyo).,
vol. 28, no. 2, pp. 118–21, Feb. 1975.
[362] M. Gutiérrez et al., “Malyngolide Dimer, a Bioactive Symmetric Cyclodepside from the
Panamanian Marine Cyanobacterium Lyngbya majuscula,” J. Nat. Prod., vol. 73, no. 4,
pp. 709–711, Apr. 2010.
[363] A. L. Waters et al., “An analysis of the sponge Acanthostrongylophora igens’ microbiome
yields an actinomycete that produces the natural product manzamine A.,” Front. Mar. Sci.,
vol. 1, p. 54, Oct. 2014.
205 [364] † Karumanchi V. Rao et al., “Manzamine B and E and Ircinal A Related Alkaloids from
an Indonesian Acanthostrongylophora Sponge and Their Activity against Infectious,
Tropical Parasitic, and Alzheimer’s Diseases,” 2006.
[365] P. W. Okanya, K. I. Mohr, K. Gerth, R. Jansen, and R. Müller, “Marinoquinolines A−F,
Pyrroloquinolines from Ohtaekwangia kribbensis (Bacteroidetes),” J. Nat. Prod., vol. 74,
no. 4, pp. 603–608, Apr. 2011.
[366] E. Bode et al., “Simple ‘On-Demand’ Production of Bioactive Natural Products,”
ChemBioChem, vol. 16, no. 7, pp. 1115–1119, May 2015.
[367] A. Agtarap, J. W. Chamberlin, M. Pinkerton, and L. K. Steinrauf, “Structure of monensic
acid, a new biologically active compound,” J. Am. Chem. Soc., vol. 89, no. 22, pp. 5737–
5739, Oct. 1967.
[368] D. E. Dorman, J. W. Paschal, W. M. Nakatsukasa, L. L. Huckstep, and N. Neuss, “The
Use of13C-NMR. Spectroscopy in Biosynthetic Studies. II[1]. Biosynthesis of narasin, a
new polyether ionophore from fermentation ofStreptomyces aureofaciens [2],” Helv.
Chim. Acta, vol. 59, no. 8, pp. 2625–2634, Dec. 1976.
[369] M. MIYAMOTO, S. KONDO, H. NAGANAWA, K. MAEDA, M. OHNO, and H.
UMEZAWA, “Structure and synthesis of neothramycin.,” J. Antibiot. (Tokyo)., vol. 30,
no. 4, pp. 340–343, 1977.
[370] C. BeBoer and A. Dietz, “The description and antibiotic production of Streptomyces
hygroscopicus var. Geldanus.,” J. Antibiot. (Tokyo)., vol. 29, no. 11, pp. 1182–8, Nov.
1976.
206 [371] A. J. Woo, W. R. Strohl, and N. D. Priestley, “Nonactin biosynthesis: the product of nonS
catalyzes the formation of the furan ring of nonactic acid.,” Antimicrob. Agents
Chemother., vol. 43, no. 7, pp. 1662–8, Jul. 1999.
[372] D. Plouffe et al., “In silico activity profiling reveals the mechanism of action of
antimalarials discovered in a high-throughput screen.,” Proc. Natl. Acad. Sci. U. S. A., vol.
105, no. 26, pp. 9059–64, Jul. 2008.
[373] B. K. BHUYAN, “Pactamycin production by Streptomyces pactum.,” Appl. Microbiol.,
vol. 10, pp. 302–4, Jul. 1962.
[374] S. D’Alessandro et al., “Salinomycin and other ionophores as a new class of antimalarial
drugs with transmission-blocking activity.,” Antimicrob. Agents Chemother., vol. 59, no.
9, pp. 5135–44, Sep. 2015.
[375] J. Prudhomme et al., “Marine actinomycetes: A new source of compounds against the
human malaria parasite,” PLoS One, vol. 3, no. 6, p. e2335, 2008.
[376] V. R. Macherla et al., “Structure-activity relationship studies of salinosporamide A (NPI-
0052), a novel marine derived proteasome inhibitor.,” J. Med. Chem., vol. 48, no. 11, pp.
3684–7, Jun. 2005.
[377] R. G. Linington, D. J. Edwards, C. F. Shuman, K. L. McPhail, T. Matainaho, and W. H.
Gerwick, “Symplocamide A, a Potent Cytotoxin and Chymotrypsin Inhibitor from the
Marine Cyanobacterium Symploca sp.,” J. Nat. Prod., vol. 71, no. 1, pp. 22–27, Jan. 2008.
[378] F. I. Nollmann et al., “Synthesis of szentiamide, a depsipeptide from entomopathogenic
Xenorhabdus szentirmaii with activity against Plasmodium falciparum,” Beilstein J. Org.
207 Chem., vol. 8, no. 1, pp. 528–533, Apr. 2012.
[379] M. Kronenwerth et al., “Characterisation of Taxlllaids A-G; Natural Products from
Xenorhabdus indica,” Chem. - A Eur. J., vol. 20, no. 52, pp. 17478–17487, Dec. 2014.
[380] G. A. McConkey, M. J. Rogers, and T. F. McCutchan, “Inhibition of Plasmodium
falciparum protein synthesis. Targeting the plastid-like organelle with thiostrepton.,” J.
Biol. Chem., vol. 272, no. 4, pp. 2046–9, Jan. 1997.
[381] M. N. Aminake et al., “Thiostrepton and derivatives exhibit antimalarial and
gametocytocidal activity by dually targeting parasite proteasome and apicoplast.,”
Antimicrob. Agents Chemother., vol. 55, no. 4, pp. 1338–48, Apr. 2011.
[382] R. P. MASKEY et al., “Anti-cancer and Antibacterial Trioxacarcins with High Anti-
malaria Activity from a Marine Streptomycete and their Absolute Stereochemistry,” J.
Antibiot. (Tokyo)., vol. 57, no. 12, pp. 771–779, 2004.
[383] B. R. Clark et al., “Natural Products Chemistry and Taxonomy of the Marine
Cyanobacterium Blennothrix cantharidosmum,” J. Nat. Prod., vol. 71, no. 9, pp. 1530–
1537, Sep. 2008.
[384] C. Mehner et al., “A Novel β-Amino Acid in Cytotoxic Peptides from the
CyanobacteriumTychonema sp.,” European J. Org. Chem., vol. 2008, no. 10, pp. 1732–
1739, Apr. 2008.
[385] H. Brockmann and G. Schmidt-Kastner, “Valinomycin I, XXVII. Mitteil. über Antibiotica
aus Actinomyceten,” Chem. Ber., vol. 88, no. 1, pp. 57–61, Jan. 1955.
[386] †,‡ Roger G. Linington, ‡,§ José González, ‡ Luis-David Ureña, ‡ Luz I. Romero, ‡ and
208 Eduardo Ortega-Barría, and † William H. Gerwick*, “Venturamides A and B:
Antimalarial Constituents of the Panamanian Marine Cyanobacterium Oscillatoria sp.⊥,”
2007.
[387] T. Luke Simmons et al., “Viridamides A and B, Lipodepsipeptides with Antiprotozoal
Activity from the Marine Cyanobacterium Oscillatoria nigro-viridis,” J. Nat. Prod., vol.
71, no. 9, pp. 1544–1550, Sep. 2008.
[388] Q. Zhou et al., “Structure and Biosynthesis of Xenoamicins from Entomopathogenic
Xenorhabdus,” Chem. - A Eur. J., vol. 19, no. 49, pp. 16772–16779, Dec. 2013.
[389] F. Grundmann et al., “Structure determination of the bioactive depsipeptide xenobactin
from Xenorhabdus sp. PB30.3,” RSC Adv., vol. 3, no. 44, p. 22072, 2013.
[390] D. Taplin, M. J. Weinstein, R. T. Testa, J. A. Marquez, and M. G. Patel, “Process for the
preparation of antibiotic W-10 complex and for the isolation of Antibiotic 20561 and
Antibiotic 20562 therefrom,” 1978.
209 RAÚL G. SARAIVA Porto, Portugal | 14.04.1989
RESEARCH EXPERIENCE OCT 2014 – MAR 2018 PhD Candidate Dimopoulos Group - Johns Hopkins Malaria Research Institute, Johns Hopkins Bloomberg School of Public Health, Baltimore, MD, USA Isolation of fermentation products of natural-occurring bacteria as antiparasitic and antiviral therapeutic and transmission-blocking drugs. OCT 2008 - JUL 2012 Junior Researcher REQUIMTE - Faculdade de Ciências, Universidade do Porto, Porto, Portugal Effect of antibiotics (fluoroquinolones, namely enrofloxacin and copper-complexed derivatives) on E. coli strains and their mechanism of translocation through the bacterial membrane. Guest at University College Dublin (UCD), Republic of Ireland (2011), and Jacobs University Bremen, Germany (2012), for weeks-long projects. EDUCATION 2018 PhD in Molecular Microbiology and Immunology Johns Hopkins University, Baltimore, MD, USA 2011 Mestrado (MSc) in Biochemistry Universidade do Porto, Porto, Portugal 2009 Licenciatura (BSc) in Biochemistry Universidade do Porto, Porto, Portugal SELECT AWARDS 2017 ASTMH 2017 Young Investigator Award – Honorable Mention, ASTMH
2014 - 2017 PhD Fellowship, Boehringer Ingelheim Fonds
2014 - 2017 PhD Emergent BioSolutions Award, Emergent BioSolutions 2014 ASTMH 2014 Annual Meeting Travel Award, ASTMH & Bill and Melinda Gates Foundation 2013 ASM ICAAC 2013 Student Travel Grant, American Society of Microbiology
2012 - 2017 Fulbright Foreign Studies Program Grant, J. William Fulbright Foreign Scholarship Board & US- Portugal Binational Fulbright Commission
PUBLICATIONS Raúl G. Saraiva and George Dimopoulos. Bacterial natural products in the fight against mosquito- borne diseases. (in preparation) Raúl G. Saraiva, Callie R. Huitt-Roehl, Abhai Tripathi, Yi-Qiang Cheng, Jürgen Bosch, Craig A. Townsend and George Dimopoulos. Chromobacterium spp. mediate their anti-Plasmodium activity through secretion of the HDAC inhibitor romidepsin. (in review) Raúl G. Saraiva*, Jingru Fang*, Seokyoung Kang, Yesseinia I. Angleró-Rodríguez, Yuemei Dong and George Dimopoulos. Aminopeptidase secreted by Chromobacterium sp. Panama inhibits dengue virus infection by degrading its E protein. (in review) Nathan J. Dennison*, Raúl G. Saraiva*, et al. (2016). Functional genomic analyses of Enterobacter, Anopheles and Plasmodium reciprocal interactions that impact vector competence. Malaria Journal 15:425 Raúl G. Saraiva, et al. (2016). Mosquito gut antiparasitic and antiviral immunity. Developmental &
210 Comparative Immunology 2016 Nov;64:53-64. Epub 2016 Jan 29. Jose Luis Ramirez*, Sarah M. Short*, Ana C. Bahia, Raúl G. Saraiva, et al. (2014). Chromobacterium Csp_P reduces malaria and dengue infection in vector mosquitoes and has entomopathogenic and in vitro anti-pathogen activities. PLOS Pathogens 10(10): e1004398. Maria J. Feio, Isabel Sousa, Mariana Ferreira, Luís Cunha-Silva, Raúl G. Saraiva, et al. (2014). Fluoroquinolone-metal complexes: a route to counteract bacterial resistance? Journal of Inorganic Biochemistry, 138 (2014) 129-143. Raúl G. Saraiva, et al. (2012). Discrimination of single-porin Escherichia (E.) coli mutants by ATR and transmission mode FTIR spectroscopy. Journal of Biophotonics 7(6):392-400. Epub 2012 Nov 26. Raúl Saraiva, et al. (2010). Solution and biological behaviour of enrofloxacin metalloantibiotics: A route to counteract bacterial resistance? Journal of Inorganic Biochemistry, 104, 8, 843-850.
ORAL COMMUNICATIONS Raúl G. Saraiva, Nathan Dennison and George Dimopoulos (2015). A natural Enterobacter mosquito midgut commensal’s role in arresting malaria parasite development. ASTMH 2015, Philadelphia, Pennsylvania, USA.
Raúl G. Saraiva, Abhai Tripathi and George Dimopoulos (2015). Anti-Plasmodium mechanisms of a
natural Chromobacterium mosquito midgut commensal. Malaria Gordon Research Seminar, Girona,
Spain.
Raúl G. Saraiva, Nathan Dennison, Abhai Tripathi and George Dimopoulos (2015). Fighting bugs with bugs: anti-Plasmodium mechanisms of mosquito midgut bacteria. Tropical Infectious Diseases Gordon Research Conference, Galveston, Texas, USA. Raúl G. Saraiva, Yuemei Dong, Abhai Tripathi, José L. Ramirez and George Dimopoulos (2014). Antimalarial and anti-dengue properties of a natural Chromobacterium mosquito midgut commensal. ASTMH 2014, New Orleans, Louisiana, USA. Raúl G. Saraiva, Matthew P. McCusker, Séamus Fanning, Paula Gameiro and Maria J. Feio (2013). The Role of OmpF and OmpC Porins on the Translocation of Fluoroquinolones and their Metallocomplexes across the Outer Membrane of E. coli. ICAAC 2013, Denver, Colorado, USA. Raúl G. Saraiva, João Almeida Lopes, Paula Gameiro and Maria J. Feio (2012). Bacterial Discrimination by ATR-FTIR Spectroscopy: Towards a Faster Diagnosis of Infection and a More Targeted Antimicrobial Therapy. IJUP 12 - 5th Meeting of Young Researchers of U.Porto. Raúl G. Saraiva, Maria J. Feio and Paula Gameiro (2010). Quinolones as Metalloantibiotics: Surpassing Resistance? - Characterization of Binary and Ternary Complexes of Cu(II)-Enrofloxacin. IJUP 10 - 3rd Meeting of Young Researchers of U.Porto.
POSTER PRESENTATIONS Raúl G. Saraiva, Callie Huitt-Roehl, Abhai Tripathi, Yi-Qiang Cheng, Jürgen Bosch, Craig A. Townsend and George Dimopoulos (2017). Chromobacterium spp. mediate their anti-Plasmodium activity through secretion of the histone deacetylase inhibitor romidepsin. ASTMH 2017, Baltimore, Maryland, USA. Raúl G. Saraiva, Callie Huitt-Roehl, Abhai Tripathi, Craig Townsend and George Dimopoulos (2016). Characterization of Chromobacterium Csp_P-produced malaria transmission-blocking and therapeutic agents. Keystone Symposium on Drug Discovery for Parasitic Diseases, Tahoe City,
211 California, USA. Jingru Fang, Raúl G. Saraiva, Yesseinia Angleró-Rodríguez and George Dimopoulos (2015). Protein-like factors from a mosquito-derived Chromobacterium reduce dengue virus infection. ASTMH 2015, Philadelphia, Pennsylvania, USA. Raúl G. Saraiva, Nathan Dennison, Abhai Tripathi and George Dimopoulos (2015). Fighting bugs with bugs: anti-Plasmodium mechanisms of mosquito midgut bacteria. Science of Eradication - Malaria, São Paulo, Brazil. Raúl G. Saraiva, Nathan Dennison, Abhai Tripathi and George Dimopoulos (2015). Fighting bugs with bugs: anti-Plasmodium mechanisms of mosquito midgut bacteria. Tropical Infectious Diseases Gordon Research Seminar, Galveston, Texas, USA. Raúl G. Saraiva, José L. Ramirez, Sarah M. Short, Ana C. Bahia, Abhai Tripathi, Yuemei Dong and George Dimopoulos (2014). Antimalarial and entomopathogenic properties of a natural Chromobacterium mosquito midgut commensal. 2014 Baltimore-Washington Area Malaria Symposium, JHMRI. Vasco Claro, Raúl G. Saraiva, Paula Gameiro and Maria J. Feio (2012). Influence of porins OmpF and OmpC in the influx of sparfloxacin in Escherichia coli. IJUP 12 - 5th Meeting of Young Researchers of U.Porto. Raúl G. Saraiva, Matthew McCusker, João Lopes, Séamus Fanning, Paula Gameiro and Maria J. Feio (2011). Discrimination of E. coli BE BL21(DE3) porin mutants by attenuated total reflectance Fourier-transformed infrared (ATR-FTIR) spectroscopy. International WE-Heraeus Workshop at Jacobs University, Bremen, Germany. Raúl G. Saraiva, Sofia Santos Costa, Isabel Couto, Miguel Viveiros, Maria J. Feio and Paula Gameiro (2010). The role of enrofloxacin-based metallocomplexes in the war against bacterial resistance. ICAR 2010 - International Conference on Antimicrobial Research, Valladolid, Spain. Raúl G. Saraiva, Maria J. Feio and Paula Gameiro (2010). Quinolones as Metalloantibiotics: Surpassing Resistance? - Characterization of Binary and Ternary Complexes of Cu(II)-Enrofloxacin. Ciência 2010 - Scientific Research in Portugal.
SELECT ORGANIZATIONAL ACTIVITIES
SEP 2013 - PRESENT JHSPH Student Assembly Select roles: President (May 2016 – present), Treasurer (May 2014 - May 2015) Oversaw team of 50 and managed six-figure budget towards the development of student-led initiatives. Represented the student community at department, school and university levels. JUN 2013 - PRESENT PAPS, Portuguese Graduate Society in North America Select roles: Chairman (Jun 2015 – present), President (Jun 2014 - Jun 2015) Led team of 20 and secured five-figure budget for the promotion of Portuguese graduates in the U.S.A. and Canada. MAR 2013 - JUN 2016 National Capital Area Chapter of the Fulbright Association Select roles: Treasurer (Jan 2014 - Jun 2016), Director (Jan 2014 - Jun 2016) Secured and managed five-figure budget towards activities for current and past Fulbright grantees in the D.C.-Maryland-Virginia area. SEP 2014 - MAY 2016 JHSPH Technology Transfer Committee Role: Student Representative Reviewed faculty-submitted grants on transferable technology and voted on their approval.
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