Exploring the Untapped Biosynthetic Potential of Apicomplexan

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Exploring the untapped biosynthetic potential of apicomplexan parasites Jack Ganley, Maria Toro-Moreno, and Emily R. Derbyshire Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.7b00877 • Publication Date (Web): 04 Oct 2017 Downloaded from http://pubs.acs.org on October 5, 2017

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Exploring the untapped biosynthetic potential of apicomplexan parasites Jack G. Ganley‡*, Maria Toro-Moreno‡*, and Emily R. Derbyshire‡†** ‡

Department of Chemistry, Duke University, 124 Science Drive, Durham, North Carolina 27708

(USA) †

Department of Molecular Genetics and Microbiology, Duke University Medical Center, 213

Research Drive, Durham, North Carolina 27710 (USA)

ABSTRACT Apicomplexan parasites encompass a diverse group of eukaryotic intracellular pathogens that infect various animal hosts to cause disease. Intriguingly, apicomplexans possess a unique organelle of algal origin, the apicoplast, which phylogenetically links these parasites to dinoflagellates and photosynthetic, coral-associated organisms. While production of secondary metabolites in closely-related organisms has been thoroughly examined, it remains widely unexplored in apicomplexans. In this perspective, we discuss previous work toward understanding secondary metabolite building block biosynthesis in apicomplexans and highlight the unexplored enzymology and biosynthetic potential of these parasites in the context of evolution.

ACS Paragon Plus Environment

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Introduction As the vast majority of eukaryotic diversity, protists fulfill important and diverse ecological roles in both terrestrial and aquatic environments.1 For example, some protists are photosynthetic and responsible for large portions of primary production, while others are responsible for harmful algal blooms.2 However, a completely separate class of protists, the apicomplexans, are best known for being notorious animal parasites. Apicomplexan parasites cause diseases such as malaria, toxoplasmosis and cryptosporidiosis, generating tremendous medical and socio-economic burden in endemic areas.3,4 Apicomplexan parasites employ diverse lifestyle strategies for survival,5 collectively infecting almost all animals from mollusks to mammals, and they exhibit a variety of complex life cycles. Most species undergo both asexual and sexual reproduction, with these reproductive cycles often occurring in specific hosts (Figure 1A). For instance, the human malaria parasite Plasmodium falciparum requires a human host for asexual reproduction in both its liver and blood stages, and also requires a mosquito to undergo recombination and transmission.6 For Toxoplasma gondii, the sexual cycle only occurs within felids, while asexual reproduction can occur in virtually any warm-blooded animal.7 On the other hand, Cryptosporidium parvum can complete its life cycle—both sexual and asexual stages—within a single animal host.8 Due to the complexity of their life cycles, parasites are difficult to study in a laboratory setting. Among the apicomplexan laboratory models that exist, most have been developed relatively recently and are often limited to a single life stage. These challenges are exacerbated by the complications involved with genetic manipulation, as many parasite genomes are intractable or require exhaustive techniques.9 Apicomplexans have highly disparate genetic


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architecture and they do not appear to cluster biosynthetic genes like other organisms, thus bioinformatic analyses are extraordinarily problematic.10 Therefore, much of our current understanding of parasite biology is limited to a disease pathogenesis context, leaving a gap in our understanding of parasite ecology. For example, the ability of parasites to adapt, develop, and interact in dramatically different environments during their life cycle is still poorly understood. In infectious agents like pathogenic bacteria and fungi, elucidation of ecological cues is often more accessible due to simplified culture conditions, well-established genetic tools, and predictable bioinformatics that are facilitated by genomic organization of functionally linked genes. As a result, it has been established that many of these organisms utilize their metabolism as an instrument to adjust to a changing environment, specifically by the production of bioactive secondary metabolites.11–13 In contrast to primary metabolism, which is highly conserved among all living organisms, secondary metabolism is distinctive to specific organisms or groups of organisms. These secondary metabolites are structurally complex and highly diverse, which often underlies their characteristic bioactivities. Strikingly, such complexity and diversity is derived from a relatively simple set of building blocks: isoprene, shikimate and acetate.14 While partial biosynthetic pathways for these building blocks are present within apicomplexans, there has yet to be an isolated and completely characterized secondary metabolite from any apicomplexan, likely due to the aforementioned complications. In this perspective, we discuss previous work toward understanding secondary metabolite building block biosynthesis in apicomplexans and highlight the unexplored enzymology and biosynthetic potential in the context of parasite evolution. Finally, we examine secondary metabolism in the context of pathogenesis, transmission, and ecology.


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Evolutionary History of Apicomplexan Parasites Eukaryotic diversity has been driven by various endosymbiotic events throughout the history of life.15 Most modern eukaryotes are the product of a primary endosymbiosis, in which a eukaryote ancestor engulfed a bacterium, giving rise to the ubiquitous mitochondrion, which results in two genomes. Most apicomplexan parasites, similar to plants and algae, contain an additional endosymbiont-derived organelle, resulting in three genomes.16 Indeed, a hallmark of apicomplexan biology is the presence of this unique non-photosynthetic organelle termed the apicoplast.17 Since its discovery almost 20 years ago, the apicoplast has served as a roadmap for apicomplexan evolution.18 Molecular characterization and sequencing of the apicoplast genome revealed that it was plastid-like, thereby strongly indicating that apicomplexan parasites have an algal ancestor.18–20 Although initial studies established an algal origin for the apicoplast,19 controversy rapidly ensued regarding the nature of this alga. Preliminary analyses based on a single plastid gene supported a green algal origin,20 while shared gene duplication of a nucleus-encoded, plastid-targeted protein pointed to the red algal counterpart.21,22 Dinoflagellates, some of which contain a red algal plastid, were investigated to settle the debate. Although dinoflagellates and apicomplexans are considered sister groups based on nuclear rRNA sequences,23 morphological features,24 and many shared nuclear genomic similarities,25 their plastids were found to be highly divergent both in terms of genomic structure and gene content.19,26 Therefore, the discovery of a protist more closely related to apicomplexans was necessary to clarify parasite evolution. In 2008, the discovery of Chromera velia, a photosynthetic alveolate, provided the missing phylogenetic link between dinoflagellates and apicomplexans.27 Originally isolated in


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the Great Barrier Reef, this protist’s nuclear-encoded housekeeping genes are most similar to apicomplexan species, while the photosynthetic genes within the plastid are most closely related to dinoflagellates.27 This connection provided evidence for the first time that the apicoplast was derived from a red alga in the same endosymbiotic event that gave rise to the dinoflagellate plastid, thus confirming a common ancestor for these two phyla (Figure 1B).27 Recently, wholegenome sequencing of C. velia and another recently discovered, closely-related organism, Vitrella brassicaformis, has been completed.28 This new class of protists, termed chromerids, provides a means to decipher genetic differences between free-living and parasitic organisms, thereby enhancing our understanding of apicomplexans. One of the major differences between the free-living chromerids and parasitic apicomplexans is the function of the plastid.29 Endosymbiosis arises when there is a selective advantage to establishing a tight partnership.30 In many cases, this selective advantage is rather easily apparent. For example, chloroplasts are widespread in nature and serve their host cells by providing energy by photosynthesis. Notably, photosynthetic capabilities had been lost in the apicoplast yet this organelle remains essential.31,32 Studies within various apicomplexans have shown that one essential aspect of the apicoplast is the biosynthesis of metabolites.33–36 Despite bioinformatic woes, genomic data for apicomplexan parasites over the past 15 years has unearthed several active apicoplast biosynthetic processes, including isoprenoid, heme, and fatty acid biosynthesis.37 A recent survey of metabolic candidate genes that mediate plastid dependency predicted that it was established early, before the split of apicomplexans and dinoflagellates, and that the key event was the loss of cytosolic isoprenoid biosynthesis in the common ancestor of apicomplexans and dinoflagellates.38 Supporting this hypothesis, plastid


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isoprenoid biosynthesis has been maintained in all apicomplexans, chromerids, and dinoflagellates known to date,38 except for Cryptosporidium, which lack an apicoplast.39

The Methylerythritol Phosphate Pathway: Terpenes and Carotenoids Despite the loss of some metabolic pathways, apicomplexans have retained numerous essential primary metabolic pathways that canonically produce building blocks for bioactive secondary metabolites (summarized in Figure 2). A well-studied example is the non-mevalonate pathway

[or

2-C-methyl-D-erythritol

4-phosphate/1-deoxy-D-xylulose

5-phosphate

(MEP/DOXP) pathway]. Originally discovered within chloroplasts in plants,40 the MEP pathway is also present in apicomplexans, where the enzymes involved are localized to the apicoplast,41 and in some bacteria.42 Specifically, the enzymes (DXS, IspC–IspH) in this pathway have been identified in the genomes of both Plasmodium43 and Toxoplasma.44 This pathway entails seven enzymatic steps to produce isopentenyl pyrophosphate (IPP, 6) and dimethylallyl pyrophosphate (DMAPP, 7) from the glucose (1) catabolites pyruvate (3) and glyceraldehyde-3-phosphate (4). IPP and DMAPP are activated isoprene starter units that are often incorporated into diverse biomolecules including proteins, cell-wall components, and secondary metabolites. These isoprenyl units can be polymerized to varying degrees and subsequently cyclized via various mechanisms to produce discrete chemical scaffolds called terpenes.45 Although the biosynthesis of IPP and DMAPP building blocks has been thoroughly characterized in both Plasmodium and Toxoplasma, the identity and function of downstream products have received less attention.43,46 Terpenes comprise the largest class of secondary metabolites, with thousands of structurally diverse metabolites characterized to date. It is among these secondary metabolites that many historically important pharmaceuticals have been discovered and isolated. An intriguing example


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of this is a class of diterpene glycosides named pseudopterosins, which have potent antiinflammatory and analgesic activities.47 Interestingly, biosynthetic studies with radiolabeled precursors uncovered that the pseudopterosins are synthesized by an apicomplexan cousin, the Symbiodinium dinoflagellate, revealing the capacity of alveolates to synthesize complex terpenes.48 Indeed, our analysis of the recently sequenced Symbiodinium minutum, C. velia, and V. brassicaformis genomes using the natural product biosynthetic gene cluster (BGC) algorithm antiSMASH49 (using the eukaryotic fungiSMASH version) putatively identified diverse BGCs including many terpene cyclase/P450 oxidase clusters. Thus bioinformatic analysis of the recently sequenced genomes hint at metabolite diversity in dinoflagellates and chromerids as well as unique genetic architectures that may be conserved within apicomplexans. Future experiments may connect complex protistan natural products with their corresponding BGCs, potentially disclosing novel enzymatic mechanisms and new secondary metabolites. Neither a terpene cyclase nor terpene synthase has been identified in apicomplexans, yet there is compelling evidence of terpene biosynthesis in Plasmodium. Besides having intriguing bioactivities, terpenes are also known to have significant ecological roles.50 For example, these metabolites often attract pollinators or fend off prey in plants.50 Several studies noted that Plasmodium-infected hosts are more attractive to Anopheles mosquitoes, raising the possibility that like plants, these parasites could attract insects with specific chemical cues.51–54 Indeed, Odom and co-workers have shown that red blood cells infected with P. falciparum release a mixture of terpenes and other organic volatiles.55 The terpenes identified include pinene (16), 4,5,9,10-dehydro-isolongifolene (15), and 8,9-dehydro-9-formyl cycloisolongifolene (17). Remarkably, these molecules are produced in various types of plants to attract pollinators,56 offering an interesting example of how divergent organisms use similar structures for common


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purposes. Additionally, recent work by Emami et al. has shown that mosquitoes prefer to feed on blood supplemented with (E)-4-hydroxy-3-methyl-but-2-enyl pyrophosphate (HMBPP, 5), a metabolite in the MEP pathway, further confirming that this pathway facilitates transmission in the Plasmodium parasite.57 Biosynthesis of larger isoprenoids has also been demonstrated in Plasmodium. Carotenoids, which are 40-carbon isoprenoids, are canonically associated with photosynthetic organisms, such as the chromerids. Interestingly, Plasmodium has been shown to synthesize the carotenoids phytoene, β-carotene (8), and lutein in red blood cell culture.58 Although there is little work completed on their physiological relevance in Plasmodium, small molecule inhibitors of carotenoid biosynthesis inhibit parasite growth, suggesting an essential function for the metabolites.58,59 Additionally, heightened oxidative stress increases carotenoid production, suggesting these molecules likely protect the parasites against free radicals.58 Again, likely due to the low interspecies gene and protein sequence homology, few enzymes downstream of DMAPP and IPP biosynthesis have been identified in Plasmodium. In Toxoplasma, evidence of carotenoid biosynthesis is indicated by the production of the phytohormone abscisic acid (ABA, 9), a catabolic product of β-carotene.60 In plants, hydra, and sponges, ABA promotes intracellular calcium secretion, which is correlated with multiple different phenotypes, including seed dormancy and stress responses, among others.61 In T. gondii, Sibley and co-workers determined that ABA stimulates parasite egress from the host cell in a dose-dependent manner and speculate that the genes for its production were retained from the endosymbiotic event that resulted in the apicoplast.60 The genes required for the biosynthesis of ABA in Toxoplasma are currently unknown, but Toxoplasma ABA-response genes have been


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identified.60 While these genes are conserved in other apicomplexan parasites like Plasmodium, concrete evidence of ABA production within Plasmodium has yet to be reported. Despite the lack of genetic evidence surrounding terpene biosynthesis within apicomplexans, we highlighted some compelling reports that hint at their presence within these parasites. These cases illustrate a larger ecological context for secondary metabolites and inform us on how they may be used by parasites as environmental cues. Terpenes enhance transmission within Plasmodium and stimulate egress in Toxoplasma, however the complete potential of terpene diversity within apicomplexans is largely unexplored.

The Shikimate Pathway: Aromatic Amino Acids and Vitamins Another pathway responsible for the biosynthesis of secondary metabolite building blocks present within apicomplexan parasites is the seven-step shikimate pathway (Figure 3A). Classically known for utilizing glycolytic fragments to synthesize aromatic metabolites, the shikimate pathway is at the crossroads of primary and secondary metabolism. De novo biosynthesis of the aromatic amino acids tyrosine, phenylalanine, and tryptophan is an essential process for protein synthesis, and these as well as other shikimate-derived metabolites can serve as key precursors for numerous secondary metabolites. In bacteria, the role of shikimate-derived metabolites in natural product biosynthesis is exemplified by catechol siderophores, which utilize 2,3-dihydroxybenzoic acid, a downstream metabolite of chorismate (2), as a substrate for nonribosomal peptide synthetases (NRPSs) to synthesize metal chelators like enterobactin from Escherichia coli.62 Plants use chorismate for a plethora of molecules including lignins, polypropenes, coumarins, and others with important biotechnological and pharmacological properties. Some of the medically relevant shikimate-derived plant metabolites include the


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salicylate analgesics, the podohyllotoxins-derived antitumor agents etoposide and teniposide, and the anticoagulant drug warfarin.14

The shikimate pathway was first recognized in apicomplexans in 1998 after the discovery

that glyphosate, an herbicide inhibitor of 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS), inhibits growth of P. falciparum, T. gondii, and C. parvum.63 Growth inhibition could be reversed in both P. falciparum and T. gondii by the addition of p-aminobenzoic acid (11) or folate (vitamin B9), suggesting that the only essential function of the shikimate pathway in red blood cell (RBC) culture is to produce precursors for folate biosynthesis.64 The discovery of the shikimate pathway spurred optimism in the malaria community, since this parasite-specific pathway could provide new, much-needed drug targets. Since its discovery nearly 20 years ago, studies examining the shikimate pathway in apicomplexans have been sparse. Particularly, it is still unknown whether the shikimate pathway serves distinct roles during other stages of the complex life cycles of these parasites or in the ecologically richer context of host-pathogen interactions. To date, bioinformatics have putatively identified all seven of the shikimate enzymes in Toxoplasma, five of which (steps 2–6) reside on a putative pentafunctional AroM complex, along with a type II 3-deoxy-D-arabino-heptulosonate 7-phosphate synthase (DHS-II) and chorismate synthase (CS) as stand-alone proteins (Figure 3B).63,65 Despite these efforts, only the shikimate dehydrogenase (SDH) domain of the AroM complex has been biochemically characterized.66 In P. falciparum, the shikimate pathway is more enigmatic. Only the last three enzymes in this pathway have been putatively assigned through bioinformatics [EPSPS, shikimate kinase (SK), and CS]65 and only CS has been partially biochemically characterized.67 The EPSPS and SK enzymes appear on a large gene encoding for a putative 2539 amino acid


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AroM complex, however the only domains that have homology to any known enzymes are EPSPS and SK (Figure 3C). With the currently available bioinformatic and biochemical information, it is difficult to estimate the evolutionary origins of the shikimate pathway in Plasmodium.68 One possibility is that the shikimate pathway is similar in architecture to Toxoplasma, meaning that DHS and CS were acquired from a prokaryotic source and the AroM complex was retained from an early eukaryotic ancestor. This would imply that the putative EPSPS-SK enzyme is encoding additional enzymatic domains for 3-dehydroquinate synthase (DHQS), dehydroquinate dehydrogenase (DHQD), and SDH that cannot be identified bioinformatically due to high sequence divergence. Clearly, there is still much to learn about the evolution and biochemistry of the shikimate pathway in Plasmodium, which remains an enticing parasite-specific pathway that may be targeted to treat malaria. The shikimate pathway is also present in the genomes of dinoflagellates69,70 and chromerids,28 providing an exciting opportunity for comparative genomic analyses.71 Our analysis of the dinoflagellate S. microadriaticum genome72 identified a putative AroM complex, yet the only predicted domain is DHQS (Figure 3D). Within this genome, there are also putative CS and DHS-I genes, however biochemical experiments to validate the function of the shikimate pathway has yet to be reported. Thus there is still much to learn about the shikimate pathway in dinoflagellates. On the other hand, genetic analysis of the shikimate pathway in chromerids predicts an AroM complex with all five enzymatic domains along with a putative CS and multiple copies of DHS-I (Figure 3E). Future genomic and biochemical investigations in chromerids may help illuminate the evolution and enzymology of the shikimate pathway in closely related apicomplexans like Plasmodium.


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Molecules likely derived from chorismate have been identified in Plasmodium-infected red blood cells (iRBC) in culture, including tyrosine (13), benzoic acid (10), salicylic acid (12), paminobenzoic acid, folate cycle metabolites (14),73 and vitamin E (18).74 Vitamin E metabolites (tocopherols and tocotrienols) are normally found within plants and include a head group derived from chorismate and an isoprenyl-derived tail.75 Within Plasmodium, vitamin E metabolites function as antioxidants and membrane stabilizers.74 Radiolabeling studies have shown that isoprenoid precursors are incorporated into vitamin E in P. falciparum, suggesting de novo synthesis by the parasite.74 Additionally, the vitamin E biosynthesis inhibitor, usnic acid, inhibits parasite growth and this phenotype can be rescued by exogenous supplementation of αtocopherol, suggesting that vitamin E biosynthesis is an essential function in this apicomplexan.74 The head group of vitamin E metabolites is usually derived from homogentisic acid, a catabolite of phenylalanine and tyrosine, suggesting an additional role of the shikimate pathway.75 As homogentisic acid has never been reported in Plasmodium metabolomics studies, and there are no clear homogentisate prenyltransferases in its genome, future work is necessary to fully understand the biosynthesis of vitamin E. Within Toxoplasma there have been no studies investigating vitamin E biosynthesis, but parasite growth inhibition in vitro and in vivo by usnic acid indicating the potential presence of this pathway.76 Biosynthesis of tyrosine and other aromatic amino acids has not been demonstrated in Plasmodium, despite the fact that supplementation of aromatic amino acids is not essential for parasite growth in RBC culture.77 This is consistent with the parasite’s ability to salvage every amino acid by breaking down host hemoglobin (expect isoleucine, which is absent from hemoglobin),78 however this scavenging mechanism is not possible in other stages of the parasite’s life cycle. Thus, an outstanding question remains about how the parasite satisfies its


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amino acid requirements without a large complement of amino acid transporters.79 Taken together, it is still unclear whether Plasmodium synthesizes aromatic amino acids de novo, procures them from the host by a unique mechanism, or can acquire these metabolites by both means.

The Acetate Pathway: Fatty Acids and Polyketides Fatty acids (FA) are ubiquitous molecules that serve diverse metabolic, structural, and signaling roles. FA synthesis occurs through a eukaryotic fatty acid synthase (FASI) and/or a prokaryotic FA synthesis pathway (FASII), which is plastid-associated in plants and algae.80,81 The discovery of the apicoplast and an increase in genomic resources was concomitant to the realization that apicomplexans possessed plastid-specific pathways, such as FASII.37 The initial excitement about yet-another parasite pathway with no mammalian counterpart was later dampened by the discovery that Plasmodium parasites preferably scavenge FA from the host during blood stage infection,82 discrediting the pathway as a drug-target for malaria treatment.35,83 Nonetheless, more recent studies have demonstrated that the FASII pathway is essential in Plasmodium sporozoites34 and late liver-stage development,35,83 once again highlighting the complexity of parasite metabolism throughout its life cycle. Driven by early drug discovery efforts, most FASII enzymes in Plasmodium have been thoroughly characterized.84 On the other hand, not much work has focused on characterizing the identity and fate of its fatty acid products. In plants, fatty acids modulate several responses to biotic and abiotic stresses. Surprisingly, common plant lipid catabolites of α-linolenic acid (19) like traumatic acid (20), jasmonic acid (21), and methyl-jasmonate (22) have been detected both in the blood plasma of Plasmodium-infected patients and in infected RBC culture.85 Importantly,


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13

C-labeled α-linolenic acid precursors were shown to incorporate into these three metabolites in

vitro, suggesting these molecules are bona fide metabolic products.85 Soon after this discovery, an untargeted metabolomics study of iRBCs detected both methyl-jasmonate and traumatic acid.73 In plants, these metabolites are hormones involved in stress response/defense, seed germination, root growth, flowering, and wound healing.86 However, in Plasmodium the function of these metabolites remains elusive. In Toxoplasma, inhibition of the FASII system does not affect acetate radiolabeling into fatty acids, suggesting the existence of FASI in this organism.87 FASI are multifunctional protein systems that share an ancestry with bacterial and fungal polyketide synthases (PKSs).88 FASI systems are often iterative and fully reducing, frequently synthesizing saturated fatty acids instead of the diversity of natural product scaffolds seen from polyketide synthases. Interestingly, there is evidence for both modular FASI and modular type-I PKS89 genes in many apicomplexans, including Toxoplasma,90 Cryptosporidium,91,92 and Eimeria,93,94 among others. In Cryptosporidium, some FASI domains have been partially characterized in vitro,95,96 however, since genetic manipulation is challenging and the predicted size of these proteins is relatively large (~20–45 kb), their functions are difficult to examine. Although there have been hypotheses regarding the function of the FASI/PKS within these parasites,90,94,97 no studies have directly connected FASI/PKS with specific molecules or phenotypes. The closest related homologs to the putative Toxoplasma and Cryptosporidium FASI/PKS genes based on BLASTP searches are putative PKSs in dinoflagellates and the closely related free-living chromerids. Dinoflagellates notably produce a plethora of fascinating secondary metabolites including the cytotoxins brevatoxin B (23), okadaic acid (24), and zooxanthellamide D (25), to name a few (Figure 4A).98 These molecules have been shown to be


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acetate-derived and have been predicted to be PKS products for decades.98 However, no PKS has been linked to a dinoflagellate polyketide product. With the recent release of multiple dinoflagellate genomes, it is apparent that they contain a multitude of PKS genes based on BGC prediction algorithms.99–101 Our fungiSMASH results predict an interesting PKS gene from S. microadriaticum (Figure 4B) and this is only one of many from this dinoflagellate. Our results also highlight some of the interesting domain architectures seen in T. gondii PKSs. For example, instead of terminal thioesterase domains, two Toxoplasma PKSs (TGME49_204560 and TGME49_294820) are predicted to have terminal reductase domains, which have only been characterized in select NRPSs and PKSs.102–105 Additionally, the predicted domain architecture in TGME49_204560 is irregular compared to typical type I modular PKSs.89 Specifically, this enzyme appears to have six ketosynthase domains, but only two acyl-transferase domains and three ACP domains. This noncanonical domain order in Toxoplasma PKSs hints at the exciting possibility for novel enzymatic mechanisms within apicomplexans (Figure 4B). While the presence of PKS genes within apicomplexans is compelling, the lack of peptidic natural products is equally as intriguing. Dinoflagellates and chromerids appear to have genes encoding for NRPS clusters, but ribosomally synthesized and posttranslationally modified peptides or RiPPs106 appear to be absent.107 Within apicomplexans, genes encoding for NRPS and RiPP clusters are absent. This could imply that apicomplexans have lost the genes for making peptidic natural products, which could be attributed to the importance of amino acid acquisition for primary metabolism and protein incorporation in intracellular pathogens. On the other hand, it remains possible that these clusters exist in apicomplexan parasites, but that they are undetectable with our current bioinformatic tools due to highly divergent sequences.


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In 2015, multiple genomes of chromerids were reported, including C. velia and V. brassicaformis, which revealed they are the closest related free-living organism to apicomplexan parasites.28 Since the discovery of the chromerids there has not been a natural product isolation reported. However, genome analysis suggests they contain multiple type I modular PKS genes that resemble both dinoflagellate and apicomplexan PKS genes. Thus chromerids may serve as an amenable model system to understand protistan enzymology, metabolism, gene architecture, and evolution. Outlook Despite tremendous advances towards understanding apicomplexan biology over the past century, we are still largely unaware of how parasites adapt to the diverse environments they encounter during their life cycles. The discovery and comprehensive study of unique, parasitespecific metabolic pathways, such as the MEP, shikimate, and acetate pathways, have been crucial for revealing much-needed drug targets and uncovering how parasites utilize secondary metabolites to influence their behavior. Since these pathways provide the building blocks for secondary metabolites, outstanding questions regarding the identity and function of potential downstream products have arisen. Although studying parasite metabolism is extraordinarily difficult, the increase in genomic resources and the advancement of genetic and metabolomics tools could pave the way to dissecting the biosynthetic potential of apicomplexans. A hurdle to these studies remains the challenges of acquiring sufficient materials from the best available model systems, which can be laborious and yield low cell numbers compared microbial and fungal systems. However, with the recent advances in methodologies to measure and analyze complex samples, metabolomics


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constitutes a promising tool for the discovery of unique secondary metabolites. Indeed, genetic disruption of candidate biosynthetic genes followed by comparative metabolomics analyses has successfully identified novel metabolites in plants,108 fungi,109 and bacteria.110 The application of CRISPR/Cas9 technology for genetic disruption of parasite genomes will now make this approach more accessible in apicomplexan parasites.111–113 Nonetheless, analogous to ‘cryptic’ or ‘silent’ BGCs in bacteria, the expression of biosynthetic pathways in apicomplexans and related species may be low in certain culture conditions. Therefore, it will be important to explore secondary metabolism in distinct parasite life stages, as well as in co-culture with other interacting microorganisms, like those found in vector microbiomes. Finally, the increase in genomic resources, including chromerid and dinoflagellate genomes, may lead to insightful comparative genomic and transcriptomic analyses for the discovery of biosynthetic enzymes. In this review, we have highlighted many of the unknowns surrounding apicomplexan enzymology and demonstrated how the discovery of secondary metabolites can facilitate our understanding of parasite transmission and pathogenesis.

ASSOCIATED CONTENT

AUTHOR INFORMATION Corresponding Author ** E-mail: [email protected] ORCID Jack G. Ganley: 0000-0002-1396-439X


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Maria Toro-Moreno: 0000-0002-0497-2259 Emily R. Derbyshire: 0000-0001-6664-8844

Author Contributions * J.G.G. and M.T-M. contributed equally to this work. Funding Sources We are grateful to Duke University for sponsoring this research. Notes The authors declare no financial interests. ABBREVIATIONS MEP Methylerythritol Phosphate Pathway IPP Isopentenyl Pyrophosphate DMAPP Dimethylallyl Pyrophosphate BGC Biosynthetic Gene Cluster HMBPP (E)-4-Hydroxy-3-Methyl-But-2-Enyl Pyrophosphate ABA Abscisic Acid NRPS Nonribosomal Peptide Synthetase RBC Red Blood Cells EPSPS 5-Enolpyruvylshikimate-3-Phosphate Synthase DHS-I 3-Deoxy-D-Arabino-Heptulosonate 7-Phosphate Synthase CS Chorismate Synthase SDH Shikimate Dehydrogenase SK Shikimate Kinase DHQS 3-Dehydroquinate Synthase DHQD Dehydroquinate Dehydrogenase FA Fatty acids FASI Eukaryotic Fatty Acid Synthase FASII Prokaryotic FA Synthesis Pathway ACP Acyl Carrier Protein iRBC Infected Red Blood Cells PKS Polyketide Synthase PPP Pentose Phosphate Pathway TCA Tricarboxylic acid cycle GPS Geranyl pyrophosphate synthase


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GGPPS Geranylgeranyl pyrophosphate synthase DHFR Dihydrofolate Reductase

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Biochemistry

Asexual in Host

A

B

Sporozoite into Host

Merozoite/ Tachyzoites Liv er

An Ce y ll

Ho sts

ALVEOLATA Apicomplexans

Plasmodium

Chromerids

Dinoflagellates

Toxoplasma In ep testi ith na eli l um

le/ sc s Mu uron Ne

Blo

od

Ciliates

RHIZARIA

Cryptosporidium

EXCAVATA

Mosquito

Sexual in Host or Vector

Vectors

Cat

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 24 of 28

STRAMENOPILES Asexual in Host

OPISTHOKONTA PLANTAE

Gametocytes

AMOEBOZOA

Figure 1. (a) Overview of apicomplexan life cycles. (b) Representation of eukatyotic diversity depicting the phylogenetic relationship between apicomplexans, chromerids, and dinoflagellates to each other and to other classifications of eukaryotes.


!

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O

CO2H

OH HO HO

Glycolysisa

CO2H

OH O

(1)a

CO2H

(13)†‡§

DHFR H N 2

(12)†§

NH2

(11)†*

PPP or TCAa

O

O

P

CO2H

N H

CO2H O O

O

HO2C

O

O OH

O

O P

CO2H

Pyruvate Kinasea

O

OHC

O

CO2H OH

(4)a

P

OH

HO

O

OH

Shikimate Pathway

O

DSX, IspC-IspG

O

O OH

O

O

P

P

O

O

CO2H

Plasmo

OH

a

(2)†*

O O

O

P O

O

O O

P O

† Product ion detected ‡ MS/MS match § Retention time match ∥ Isotope labeling * Pathway inhibitor

Enzyme, metabolite, or pathway may be present in both host and parasite

O

(6)a

IspH

(5)

Isoprenoid Biosynthesis

Both

(10)†‡§

Toxo

O

O

SK, EPSP synthase, & CS

O

O

(3)a

OH

DHS II

OH P

(14)†*

N H

Neither CO2H

DHQS, DHQD, & SDH

OH

CHO

O

CO2H

N H

Key

OH O

N H

H N

N

NH2

HO

O

OH

CO2H

OH

P O

O

GPS

O O

(7)a

P O

CHO O

(15)†‡

(17)†‡

(16)†‡§

GPS GGPPS

O Pyruvate Dehydrogenasea

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biochemistry

O

PYS 2

OH FabG

O CO2H OH

(9)†‡§* O

P O

HO O O

(18)†‡§∥*

O

R

FabZ

SACP

CO2H

O

O

R

SACP

Type II FA Biosynthesis

FabB/F or FabH

O R

O SCoA

O

O O

2

2

(8)†‡§*

O

P

R

5

FabI

SACP

Acetyl-CoA carboxylasea + FabD

HO2C

O

SACP

SACP

(19) HO2C

6

CO2H

(20)†‡§∥

O O R

R = H: (21)†‡∥ R = Me: (22)†‡∥

!

Figure 2. Global metabolic map of Toxoplasma and Plasmodium discussed in this perspective. Blue metabolites have been reported in at least one Plasmodium spp., those in red have been reported in at least one species of Toxoplasma and those in black are either host metabolites or reported in both apicomplexans. Dashed arrows indicate an unknown biosynthetic pathway. Plain arrows indicate putative enzymes for biosynthetic pathways. The presence of both dashed and solid arrows indicate a partially known biosynthetic pathway.


Biochemistry

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 26 of 28

A. Canonical Shikimate Pathway CO2H O O

P O

O O

O P

O

CHO

O

1

HO2C

O

O OH

4-P

CO2H

HO CO2H OH

DHS

OH D-Erythrose

PEP

OH

OH

DHAP

O P

DHQS

O

2

DHQD

O

OH

3

O

OH OH

OH

O

3-Dehydroquinate

3-Dehydroshikimate

SDH

CO2H

CO2H O

CS

O

CO2H

7

OH

CO2H

O

P O

O

CO2H

OH

Chorismate

P O

SK

O

OH

5

OH

HO

OH

OH

Shikimate 3-P

Shikimate

D. Symbiodinium microadriaticum DHQS

SDH SDH

EPSPS SK

CS

6

O

EPSP

B. Toxoplasma gondii ME49 DHQS

O

EPSPS

O

4

CO2H

DHQD

0.5 kb

DHS-II

CS

CS

0.5 kb

DHS-I

C. Plasmodium falcipraum 3D7

E. Vitrella brassicaformis DHQS

EPSPS CS

EPSPS

SK

DHQD SDH

SK

0.5 kb

CS

DHS-I

0.5 kb

!

Figure 3. (a) Overview of the canonical seven-step shikimate pathway. Genetic architecture of shikimate pathway genes in the apicomplexans (b) T. gondii ME49 and (c) P. falciparum 3D7, (d) in the dinoflagellate S. microadriaticum, and (e) in the chromerid V. brassicaformis. Predicted introns were deleted (putative mRNA used). Domains were predicted using the NCBI Conserved Domain tool.


Page 27 of 28

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biochemistry

A. Protistan Polyketide Natural Products – Unknown Biosynthetic Orgin HO O O

O

O

HO OH

O H

HO

HO H O

O

O

O

HN

O

OH OH OH

O

(23)

OH H H

O

O

OH

OH

H OH

H

(25)

B. Protistan Polyketide Synthases/Type I Fatty Acid Synthases

1 kb

Symbiodinium microadriaticum strain CCMP2467 – OLQ06785.1

DH ER KR ACP KS AT

DH KR ACP KS KR ACP KS

Domain Key KS ketosynthase

ACP

acyl carrier protein

AT

acyl transferase

DH DH ER KR ACP KS DH KR ACP KS DH

1 kb Vitrella brassicaformis – Vbra_18624

KR ketoreductase ACP

OH OH

(24)

Me

OH

A

H

H

HO OH

H OH

O O

O

OHC

OH

CO2H

O

O

O

O

O

O

KS AT

DH ER KR ACP KS DH ER KR ACP KS AT

ER

1 kb

A KR KS

R

reductase

TE thioesterase

CAL

CoA ligase

KR ACP NAD TE

Toxoplasma gondii ME49 – TGME49_204560

KS ACP KS AT ACP KS ACP KS KS AT

DH dehydrogenase

enoylreductase NAD NAD binding domain adenylation

S/MT

Sulfo or methyl transferase

R

1 kb Toxoplasma gondii ME49 – TGME49_294820

CAL ACP

DH ER KR ACP

AT

DH KS

ER KR ACP

KS

DH

KR ACP ER

KS

AT

DH

ER KR ACP

KS

DH AT

ER KR ACP

KS

R

S/MT

DH AT

!

Figure 4. (a) Isolated natural product polyketides from dinoflagellates. (b) Putative PKS/FAS genes and predicted domain architecture (predicted by fungiSMASH) from a dinoflagellate, chomerid, and apicomplexan parasite.


Biochemistry

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 28 of 28

Table of Contents Figure Apicomplexans

CO2H Me O

O

O

P

P

O-

O

O-

O O-

HO2C

SCoA HO

OH OH

Secondary metabolite building blocks

?


Exploring the Untapped Biosynthetic Potential of Apicomplexan

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