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Microsporidia: Why Make Nucleotides if You Can Steal Them?


Microsporidia are strict obligate intracellular parasites that infect a wide range of eukaryotes including humans and economically important fish and insects. Surviving and flourishing inside another eukaryotic cell is a very specialised lifestyle that requires evolutionary innovation. Genome sequence analyses show that microsporidia have lost most of the genes needed for making primary metabolites, such as amino acids and nucleotides, and also that they have only a limited capacity for making adenosine triphosphate (ATP). Since microsporidia cannot grow and replicate without the enormous amounts of energy and nucleotide building blocks needed for protein, DNA, and RNA biosynthesis, they must have evolved ways of stealing these substrates from the infected host cell. Providing they can do this, genome analyses suggest that microsporidia have the enzyme repertoire needed to use and regenerate the imported nucleotides efficiently. Recent functional studies suggest that a critical innovation for adapting to intracellular life was the acquisition by lateral gene transfer of nucleotide transport (NTT) proteins that are now present in multiple copies in all microsporidian genomes. These proteins are expressed on the parasite surface and allow microsporidia to steal ATP and other purine nucleotides for energy and biosynthesis from their host. However, it remains unclear how other essential metabolites, such as pyrimidine nucleotides, are acquired. Transcriptomic and experimental studies suggest that microsporidia might manipulate host cell metabolism and cell biological processes to promote nucleotide synthesis and to maximise the potential for ATP and nucleotide import. In this review, we summarise recent genomic and functional data relating to how microsporidia exploit their hosts for energy and building blocks needed for growth and nucleic acid metabolism and we identify some remaining outstanding questions.


Microsporidia are fungi-related eukaryotic parasites with over 1,400 reported species that infect a wide range of hosts including humans, mammals, and insects [1,2]. They are all strict obligate intracellular parasites and can only complete their life cycle within an infected eukaryotic host cell. The life cycle (S1 Fig) of a typical microsporidia begins with the germination of a resistant spore that physically injects the sporoplasm into the host cell through a polar tube [1]. The parasite cell (meront) grows and multiplies within the host cell cytoplasm through several rounds of division and differentiates into a spore, which exits the host cell, typically through host cell lysis, to complete the cycle (S1 Fig).

Microsporidian genomes have some of the smallest coding capacities among eukaryotes [3] and analyses reveal that they have lost many of the biosynthetic genes needed for making basic metabolites, such as the nucleotides required for making DNA and RNA, amino acids for making proteins, and lipids for making membranes [4]. Genome analyses also show that all microsporidia have lost the pathways for oxidative phosphorylation and the tricarboxylic acid (TCA) cycle [5,6] and Enterocytozoon bieneusi, a major pathogen of immunocompromised patients, has also lost glycolysis [7]. So the capacity for independent biosynthesis of ATP appears to be very limited in microsporidia. Since a typical cell requires enormous amounts of ATP (107 ATP molecules per second) to grow and divide [8], actively growing microsporidia must impose a very high demand for ATP on infected host cells. In this review, we summarise what is currently known about the transport proteins and mechanisms (summarised in Fig 1) that are used to acquire the ATP and other nucleotides needed to support the intracellular growth and replication of this enormously successful group of eukaryotic parasites.

Fig 1. Nucleotide acquisition and metabolism in host cells and microsporidia.

Schematic illustration showing nucleotide metabolism in a typical microsporidian parasite within an infected host cell. Host cells can make nucleotides via de novo biosynthesis and regenerate ATP by oxidative phosphorylation—these pathways are absent in microsporidia [5]. Host purine nucleotides can be stolen using microsporidia NTT transporters and then efficiently used and recycled by the parasites [18]. Key: (a) E. cuniculi physically tethers mitochondria using an unidentified protein [49]. (b) Only EcNTT3 of E. cuniculi has been found in the mitosome [38]. (c) Nematocida may secrete a hexokinase into the host cell to stimulate host nucleotide production [6]. (d) Nucleoside kinases are apparently absent from some microsporidian genomes but are present in Trachipleistophora hominis [18]. (e) Thymidine kinase is present in some microsporidia but not all [18]. (f) The microsporidian RNA degradation pathway is shown in S2 Fig.

Nucleotide metabolism and availability within the host cell

Nucleotides are the building blocks of DNA and RNA that are essential to all life. In free-living species, the eight major purine or pyrimidine nucleoside triphosphates (Box 1) can either be synthesised de novo from amino acids or recycled (salvaged) from the rapid turnover of RNA using pathways that are conserved among prokaryotes and eukaryotes [9,10]. While nucleotide synthesis is located in the cytoplasm, nucleotides can freely diffuse into the eukaryotic nucleus [11], possibly explaining how some microsporidia can complete their lifecycle in the host nucleus [12,13].

Box 1. Facts and Figures: Nucleotides, nucleosides, and their cellular concentrations and bacterial and microsporidian NTTs

ATP demand in cells: It is estimated that 50 ATP are used to make one nucleotide from scratch when the ATP needed to make all of the required co-factors and substrates is also included [7,14]. Around 107 ATP per second is used in a typical cell [7,14]

Nucleotides: There are four major types of nucleotide triphosphates that make up RNA (ATP, GTP, CTP, UTP) and four that make DNA (dATP, dGTP, dCTP, dTTP). When DNA and RNA are degraded, the nucleoside monophosphate is released which can be recycled.

Nucleotides are made of 3 components:

  • Purine (adenine, guanine) or pyrimidine (cytosine, uracil or thymine) base
  • A ribose or deoxyribose sugar
  • One to three phosphate groups

Nucleosides (adenosine, guanosine, cytidine, uridine, thymidine): Similar to nucleotides but do not have the phosphate groups.

Intracellular concentrations: Mean nucleotide concentrations in a mammalian cell [10]: ATP (3000 μM), GTP (500 μM), UTP (600 μM), CTP (300 μM).

Deoxyribonucleotide (dNTP) and nucleoside concentrations: 4–40 μM.

Nucleotide transporter (NTTs) affinities: Apparent Km (ATP) of NTTs from the microsporidian E. cuniculi NTTs [37] are: EcNTT1 = 11.4 μM, EcNTT1 = 19.8 μM, EcNTT1 = 24.2 μM, EcNTT1 = 2.6 μM. Apparent Km (ATP) of bacterial NTTs are: Protochlamydia amoeabophila *PamNTT1 = 95 μM, PamNTT2 = 437 μM, PamNTT5 = 360 μM; Chlamydia trachomatis: Npt1Ct = 48 μM, Npt2Ct = 1158 μM; RpTLC = 100 μM [4043].

* Note, the apparent Km (ATP) for PamNTT1 in liposomes was 17–100 μM, depending on intra-liposomal nucleotide concentration [43]).

Nucleotide concentrations in mammalian cells have been well documented [11], with ribonucleoside triphosphates (NTPs), particularly ATP (Box 1) [11], at highest concentrations. The concentrations of di- and monophosphate forms of nucleotides, of nucleosides, and of nucleobases are all less than <5μM compared to ~3000 μM for ATP [11] (Box 1). The deoxyribonucleotides (dNTP) that are the building blocks of DNA are also at considerably lower concentrations in a cell than the corresponding NTPs, possibly explaining why microsporidia have retained the ability to synthesise their own dNTPs [4] providing they have a source of NTPs from the host cell. Tight control over dNTP synthesis is also critical to cell viability [14] as discussed below.

Nucleotide biosynthesis pathways and the microsporidia

Making nucleotides de novo from amino acids is energy expensive and is estimated at around 50 ATP per nucleotide when the costs of co-factors and substrates are included [15]. Microsporidian genomes lack the enzymes needed for the de novo synthesis of nucleotides [4,6] (Figs 2 and 3) including phosphoribosyl pyrophosphate (PRPP) synthase [4], which makes the substrate PRPP that is required for the activation of ribose-5-phosphate for both purine and pyrimidine de novo pathways [16,17]. The loss of this biosynthetic function alone means de novo synthesis is not possible. Other key enzymes are also absent [18] (Fig 3), including IMP cyclohydrolase that makes inosine monophosphate (IMP)—the first purine nucleotide in the de novo pathway—and UMP synthase [18], which makes the first pyrimidine nucleotide, uridine monophosphate (UMP). These enzymes are also missing in obligate intracellular bacteria [18] (Figs 2 and 3), indicating that de novo nucleotide synthesis is also not possible in these organisms. However, upon obtaining nucleotides from the host, microsporidia and these bacterial pathogens have retained the core suite of enzymes for metabolising and recycling nucleotides (Fig 2B) [18].

Fig 2. Nucleotide biosynthesis in microsporidia.

(A) Nucleotide biosynthetic enzymes found in the genomes of yeast, Escherichia coli, Microsporidia, and intracellular pathogenic bacteria for purine (red) and pyrimidine (green) metabolism according to the KEGG database [60] are shown along with respective EC numbers (see S3 Fig). Sc = Saccharomyces cerevisiae; Th = Trachipleistophora hominis; Ec = Encephalitozoon cuniculi; Ct = Chlamydia trachomatis; Rr = Rickettsia rickettsii. (B) Purine and pyrimidine pathways retained in microsporidia genomes (black arrows) enable recycling of all of the major nucleotides based on available genome data [18]. Some exceptions to the general rule can be found in (a) T. hominis or (b) N. ceranae, as described in the text.

Fig 3. Key metabolic steps lost by microsporidia and intracellular bacteria.

Several key enzymes and the associated pathways are shown that have been lost during microsporidian evolution (green square). The implications for the parasites are shown to the right. With the exception of PRPP synthase, all enzymes have been lost by obligate intracellular bacterial pathogens (green square). Exceptions: (a) nucleoside kinases are retained in T. hominis

Inter-converting purine nucleotides.

In free-living species, the first purine nucleotide produced by de novo synthesis is IMP, which requires hydrolysis of six ATP, and can then be converted to either AMP or guanosine monophosphate (GMP), which requires hydrolysis of an additional ATP. Genome analyses [4,6,18] suggest that microsporidia lack the enzymes for inter-converting IMP to AMP or GMP, or for inter-converting between guanosine and adenosine nucleotides [4,6,18], suggesting that they must import both types of purine nucleotide. In addition, the apparent lack of a GMP synthase or adenylosuccinate synthase in the genome sequences of microsporidia [4] also suggests that import of inosine nucleotides (IMP/IDP/ITP) would be fruitless as they cannot be utilised. Microsporidia have, however, retained all of the necessary enzymes for converting between the three phosphorylation states of guanosine and adenosine nucleotides, including the broad-spectrum nucleoside diphosphate kinase (NDK/YNK in yeast), which is highly expressed during infection and in the spore stage of the microsporidian Trachipleistophora hominis [19,20], suggesting an important role in parasite metabolism.

Inter-converting pyrimidine nucleotides.

The first pyrimidine nucleotide produced by the de novo pathway in free-living species is UMP, which can be stepwise converted into the pyrimidine triphosphates dTTP, UTP, or CTP (Fig 1). Analysis of microsporidian genomes suggests that some species are unable to convert CTP and its derivatives to UTP as they lack the relevant deaminase enzymes [18], suggesting that they need to import UTP from the host. Some microsporidians have retained the enzyme CTP synthase enabling them to make CTP from imported UTP as a route towards synthesising cytidine derivatives [4,6]. However, this step is energy expensive and, therefore, an ability to steal host CTP would be beneficial; hence, some intracellular Chlamydia possess a CTP synthase and can also import CTP from the host cell [21]. Microsporidia have generally retained all of the enzymes [18] needed to make dTTP from UTP, as described below. In summary, if UTP can be stolen from the host, microsporidia appear to have the necessary enzymes to synthesise the other pyrimidines.

Synthesis of dNTPs.

Maintaining the correct cellular concentrations of the deoxyribonucleotides (dNTPs) that are needed to make DNA is critical as an imbalance can be highly mutagenic [14,2224]. Control over cellular dNTP concentrations is exerted at the level of the broad-spectrum enzyme ribonucleotide reductase, which is tightly regulated by the dNTPs themselves [12]. The need to exert tight control over dNTP concentrations may explain why microsporidia [4,18], along with the obligate intracellular bacteria Chlamydia [25] and Rickettsia [18], have retained their own ribonucleotide reductase and, hence, do not depend on the relatively low levels of dNTPs in the host cytoplasm [11].

Unlike the other dNTPs, dTTP must be synthesised from dUMP and requires the enzyme thymidylate synthase that is found in some, but not all, microsporidia [18]. The apparent absence of thymidylate synthase in Nosema ceranae (and potentially other microsporidians that lack this enzyme), suggests they must make or acquire dTTP by other means. Indeed, several species of microsporidia, including N. ceranae and Encephalitozoon cuniculi [26], have acquired the enzyme thymidine kinase by lateral gene transfer from bacteria and this would allow them to phosphorylate thymidine imported from the host cell [26].

Microsporidia and the recycling of nucleotides

The continual turnover of RNA, which releases the nucleoside monophosphates, represents a major, ready-made source of nucleotides [27]. The enzymes needed to regenerate nucleoside triphosphates following nucleic acid degradation have been retained by microsporidia [4] and include dedicated monophosphate kinases and the broad-spectrum nucleoside diphosphate kinase (NDK/YNK in yeast) (Fig 1) [4,18]. While nucleoside triphosphate regeneration involves some energy-expensive steps, it still represents a more economical source of nucleotides compared to de novo nucleotide synthesis.

In eukaryotic cells, RNA degradation occurs by two main pathways [28] (S2 Fig), both of which are initiated by the deadenylation of RNA by enzymes including the Ccr4 complex. The machinery for both RNA degradation pathways is present in microsporidia (S2 Fig) including the Ccr4 complex [4]; the decapping enzymes Dcp1 and Dcp2 [29]; the exonuclease Rat1, which is involved in 5’ to 3’ degradation; and the exosomal complex needed for 3’ to 5’ degradation, including the exosomal protein Dis3, which acts as the main catalytic component of the exosome complex in yeast [27,30].

In comparison to other organisms, there are some important salvage enzymes missing from microsporidian genomes (Fig 2) [31]. For example, microsporidia appear to lack [18] the various enzymes for conversion of nucleobases to nucleotides, suggesting that they cannot use nucleobases as starting points for nucleotide synthesis. In addition, the ribose-phosphate moiety of nucleotides that represents a potential carbon or energy source [31] is unlikely to be recycled in microsporidia as it must be converted to intermediates for glycolysis [16]—a pathway that appears to be most active in the spore stage of microsporidia [19,32] or has been lost altogether [7].

Genome analysis suggests that microsporidia have retained the broad-spectrum enzyme 5'-nucleotidase that converts nucleotides to nucleosides, a known regulatory function of this enzyme, thus maintaining an optimal nucleotide balance required for normal cell physiology [24,33]. By contrast, some microsporidia are unable to convert nucleosides to nucleotides [18] as they lack the necessary kinases, and this raises doubt about whether these species could utilise nucleosides imported from the host. By contrast, T. hominis has retained several nucleoside kinases [18] that would enable it to utilise nucleosides stolen from the host or generated internally by the activity of the 5'-nucleotidase.

How do microsporidia acquire the energy and nucleotides they need?

Nucleotides cannot be transported physiologically across plasma membranes without specific transporters [34, 35]. In mammalian cells, extracellular nucleotides are generally converted to nucleosides that can then be imported by members of the equilibrative nucleoside transporter (ENT) family [34]. Members of the ENT family are found in some parasites, but, so far, are not found in microsporidia [4,35]. Instead, microsporidia use a family of nucleotide transporters (NTTs) that are also found in phylogenetically diverse intracellular bacterial pathogens, such as Chlamydia and Rickettsia [36], to import nucleotides directly from the host cell cytoplasm.

Phylogenetic analyses [18,37,38] suggest that a single NTT gene was probably acquired by horizontal transfer from bacteria into the microsporidian common ancestor. This was followed by lineage-specific gene duplications to generate the multiple copies of NTT genes found in contemporary microsporidian genomes [4,6,18]. For example, the microsporidia T. hominis, E. cuniculi, and E. bieneusi have four NTTs while Spraguea lophii has six [39]. NTT gene duplications may provide the parasites with the starting materials for NTT functional diversification, differential NTT expression throughout the life cycle, or a gene dosage effect to increase the amount of NTTs being made.

Microsporidian NTTs have been found to be highly and differentially expressed during the different stages of the microsporidia life cycle [6,20] including spores [19]. In E. cuniculi, one of its four NTTs is localised to its highly reduced mitochondrion (called a mitosome [40]), whereas the other three NTTs are located at the parasite cell surface [38]. By contrast, all four NTTs of T. hominis are located at the cell surface [18], suggesting that the location of NTT transporters at the host–parasite interface is a general strategy used by microsporidia to exploit host cells [18,38].

The lack of axenic culture systems and the strict obligate intracellular lifestyle of microsporidia has impeded attempts to genetically manipulate these parasites [41], and all of the published functional work with microsporidian NTTs has employed heterologous expression in engineered E. coli strains (Box 1) [18,38]. This work has shown that the four NTTs in E. cuniculi and T. hominis can all transport ATP [18,38] and, hence, they can, in principal, be used to steal vital energy from the infected host cell. The T. hominis NTTs can also transport other purine nucleotides (ADP, GTP, and guanosine diphosphate [GDP]) that are needed for DNA and RNA biosynthesis, but not pyrimidine nucleotides [18]. Dose response data suggests that the four E. cuniculi NTTs [38] have a high affinity for ATP, with apparent Km values considerably lower than that of bacterial NTTs [4244] (Box 1), and well below host cytosolic ATP concentrations [11]—implying a high level of ATP transport could occur during infection.

The lack of transport of pyrimidine nucleotides by T. hominis NTTs expressed in E. coli raises the question of how T. hominis obtains the pyrimidines needed to make DNA and RNA. As discussed above, it appears that all major pyrimidine nucleotides can generally be synthesised if UTP is available, but no UTP transport was detected by the T. hominis NTTs [18]. It is not yet clear if an inability to transport pyrimidine nucleotides is a general feature of microsporidian NTTs because transport of radiolabelled pyrimidine nucleotides has only been tested for T. hominis [18,38]. Substrate competition experiments with the E. cuniculi NTTs suggest that ATP transport is not reduced by cold competitor pyrimidine nucleotides and, hence, pyrimidine transport appears unlikely [38]. However, since some bacterial NTTs can transport both purine and pyrimidine nucleotides [45] as well as NAD [46], it would not be too surprising if microsporidian NTTs in different species have also evolved to transport different substrates, including pyrimidine nucleotides. The extraordinary possibilities of NTT-mediated transport were recently demonstrated when a diatom NTT was used to import unnatural nucleotides into an engineered E. coli strain to create a semisynthetic organism with an expanded genetic alphabet [47].

It has been suggested [6] that homologues of the bacterial-like NupG transporters that are conserved on all microsporidian genomes [4] might, as in E. coli, transport purine and pyrimidine nucleosides including adenosine and uridine [48]. However, there is currently no functional data for microsporidia NupG-like transporters and genome analysis suggest that some microsporidia would also be unable to convert imported nucleosides into nucleotides as they lack the necessary nucleoside kinases [18] (Figs 2 and 3). In common with other parasites, microsporidia possess genes for a number of transporter families and hypothetical transporters [4,19,35] whose locations and functions are currently unknown. Given the minimal nature of microsporidian primary metabolism, it would not be surprising if some of these putative transport proteins also played roles in providing the metabolites needed for parasite growth, DNA biosynthesis, and RNA biosynthesis.

Manipulating the host cell to support nucleotide acquisition

While NTTs confer an ability to steal nucleotides, the absolute dependence on host nucleotides could potentially act as an Achilles’ heel for microsporidia if the host was able to limit their availability. Recent work now suggests that microsporidia have evolved strategies to manipulate host cells and to ensure that a ready supply of host nucleotides is maintained.

Stimulating nucleotide metabolism in the host is one obvious strategy to increase the available pools of energy and nucleotides for import and there is some preliminary data that hints at how microsporidia could possibly do this. RNAseq analysis of the microsporidian Nematocida parisii during infection of the nematode Caenorhabditis elegans demonstrated the upregulation of a microsporidian hexokinase during early infection, despite low expression of other glycolytic enzymes, suggesting it may have an alternative role [6]. The presence of a signal peptide in this hexokinase, which was not found in the other glycolytic enzymes in this species [6], raised the possibility that it might be secreted into the host. The presence of a functional secretion signal was supported when the protein was expressed in yeast [6]. Hexokinase catalyses the phosphorylation of glucose to glucose-6-phosphate that can be used to synthesise PRPP and other nucleotide biosynthetic precursors. Thus, secretion of a microsporidian hexokinase could, in principal, stimulate host nucleotide production [6]

Mitochondria are rich sources of ATP and the microsporidian E. cuniculi forms an intimate association with host mitochondria [49,50], possibly to maximise the surface area in contact between parasite and organelle. Mitochondria appear to be physically attached to the E. cuniculi parasitophorous vacuole [49] suggesting that it may be porous to ATP or has associated transport proteins to permit ATP passage [51]. Association with host mitochondria has also been reported for intracellular bacterial parasites including Legionella [52] and Chlamydia [53], as well as the microbial eukaryote Toxoplasma [54], which secretes a protein called MAF1 to tether mitochondria to the parasite surface [54]. Homologues of MAF1 are not present in the E. cuniculi genome [49], but electron microscopy images suggest that E. cuniculi does use electron-dense proteinaceous structures to tether mitochondria and that it also influences the location of mitochondrial ATP-gating channels in the outer mitochondrial membrane [49]. Whether microsporidia can also increase ATP production by host mitochondria is not known, but no changes in mitochondrial activity were detected during infection by E. cuniculi [49,50]. Interestingly, however, comparing the transcriptome of T. hominis-infected mammalian host cells with non-infected controls did suggest that host-energy metabolism and mitochondrial biogenesis were both induced upon infection [20].

Future perspectives

Comparative genomics data suggests that microsporidians cannot make their own nucleotides and must, therefore, import them from infected host cells. Given the central and non-redundant role of nucleotide import and metabolism in the parasite life cycle, this makes it a logical therapeutic target, especially against enzymes and transporters that are not found in host species. These would include the bacterial-derived NTT nucleotide transporters that appear to be so critical for microsporidia growth and replication, which are not found in vertebrates. Since NTT transporters are also used by important bacterial pathogens like Chlamydia and Rickettsia to exploit the eukaryotic host cells that they infect, finding effective therapeutic agents that act against NTTs would have applied interest beyond microsporidia, although the presence of multiple copies of such transporters in Microsporidia and their overlapping substrate ranges may complicate or hinder such approaches. Functional work on the other transporters present in microsporidian genomes to establish which are surface-located and which substrates they transport will also be important for filling the many gaps in our understanding of microsporidia–host interdependencies, including the acquisition of pyrimidine nucleotides needed for parasite DNA and RNA biosynthesis. Experimental work is essential to identify substrate specificities given the highly derived nature of microsporidia DNA and protein sequences and the resultant difficulty in reliably predicting protein function based upon low sequence similarity to characterised proteins from model organisms.

The mechanism of transport used by microsporidian NTTs has not been determined. NTT transporters in bacteria can function as ATP/ADP exchangers [42,45] to provide energy, or as proton-driven symporters to provide net import of nucleotides for DNA and RNA synthesis [42,45]. It is currently unclear if microsporidian NTTs also use different transport mechanisms, but given the demands imposed by parasite growth and replication, we think it very likely that both symporters and exchangers have also evolved during microsporidia NTT evolution. Understanding how NTTs function in detail would obviously be aided by the availability of a high-resolution structure for one or more NTT proteins, but unfortunately none are yet available. In particular, the levels of expression of microsporidian NTT proteins in E. coli are very low so providing enough protein for structural and comprehensive liposome studies [45] will require systematic investigation of different strategies to improve protein yields and the evaluation of eukaryotic [55], as well as prokaryotic expression systems.

Bacterial intracellular pathogens are known to utilise diverse effector proteins to manipulate the metabolism of the host cells that they infect [56,57]. It is very likely that microsporidia also use a variety of strategies and secreted proteins to manipulate host cellular processes, including energy and nucleotide metabolism. At present, it is difficult to investigate these phenomena effectively because of the lack of tools for reproducible genetic manipulation of well described microsporidian model species. However, some progress has recently been made using RNAi on the honeybee microsporidian parasite N. ceranae [58,59]. If the promise of these initial experiments can be confirmed and extended to more tractable model species, it might finally be possible to test hypotheses of microsporidian protein function and their potential role(s) in microsporidia–host interactions at the molecular level.

Supporting Information

S1 Fig. The lifecycle of a microsporidian.

A typical life cycle begins with the germination of a spore, which discharges a polar tube that pierces the host cell plasma membrane enabling transfer of the parasite sporoplasm into the host cytoplasm. The parasite cell (meront) grows and divides, sometimes within a parasitophorous vacuole (not shown), and after several rounds of division, differentiates back into spores, which are released following host cell lysis. The life cycle of T. hominis during infection of cultured cells, is around 3–4 days.


S2 Fig. RNA degradation pathways in microsporidia enable recycling of nucleotides.

RNA (mRNA and rRNA) is a rich source of nucleotides that can be continually recycled in the cell. Genome analysis [4] suggest microsporidia have retained components needed for RNA degradation via 2 main pathways (a) 3’>5’ degradation involving the exosomal complex (b) 5’>3’ degradation involving decapping enzymes and Rat1. All enzymes depicted here are conserved in at least 9 microsporidian genomes [4].


S3 Fig. Enzymes involved in purine and pyrimidine nucleotide biosynthesis and their EC numbers as given in Fig 2A in the main text.



We are grateful to Dr. Kacper Sendra for critical comments on the manuscript.


  1. 1. Williams BA (2009) Unique physiology of host-parasite interactions in microsporidia infections. Cell Microbiol 11: 1551–1560. pmid:19673893
  2. 2. Weiss LM, Becnel JJ (2014) Microsporidia: Pathogens of Opportunity: Wiley-Blackwell.
  3. 3. Corradi N, Pombert JF, Farinelli L, Didier ES, Keeling PJ (2010) The complete sequence of the smallest known nuclear genome from the microsporidian Encephalitozoon intestinalis. Nat Commun 1: 77. pmid:20865802
  4. 4. Nakjang S, Williams TA, Heinz E, Watson AK, Foster PG, et al. (2013) Reduction and expansion in microsporidian genome evolution: new insights from comparative genomics. Genome Biol Evol.
  5. 5. Keeling PJ, Corradi N (2011) Shrink it or lose it: balancing loss of function with shrinking genomes in the microsporidia. Virulence 2: 67–70. pmid:21217203
  6. 6. Cuomo CA, Desjardins CA, Bakowski MA, Goldberg J, Ma AT, et al. (2012) Microsporidian genome analysis reveals evolutionary strategies for obligate intracellular growth. Genome Res 22: 2478–2488. pmid:22813931
  7. 7. Keeling PJ, Corradi N, Morrison HG, Haag KL, Ebert D, et al. (2010) The reduced genome of the parasitic microsporidian Enterocytozoon bieneusi lacks genes for core carbon metabolism. Genome Biol Evol 2: 304–309. pmid:20624735
  8. 8. Phillips R, Milo R (2009) A feeling for the numbers in biology. Proc Natl Acad Sci U S A 106: 21465–21471. pmid:20018695
  9. 9. Witz S, Jung B, Furst S, Mohlmann T (2012) De novo pyrimidine nucleotide synthesis mainly occurs outside of plastids, but a previously undiscovered nucleobase importer provides substrates for the essential salvage pathway in Arabidopsis. Plant Cell 24: 1549–1559. pmid:22474184
  10. 10. Evans DR, Guy HI (2004) Mammalian pyrimidine biosynthesis: fresh insights into an ancient pathway. J Biol Chem 279: 33035–33038. pmid:15096496
  11. 11. Traut TW (1994) Physiological concentrations of purines and pyrimidines. Mol Cell Biochem 140: 1–22. pmid:7877593
  12. 12. Scheid P (2007) Mechanism of intrusion of a microspordian-like organism into the nucleus of host amoebae (Vannella sp.) isolated from a keratitis patient. Parasitol Res 101: 1097–1102. pmid:17579885
  13. 13. Stentiford GD, Feist SW, Stone DM, Bateman KS, Dunn AM (2013) Microsporidia: diverse, dynamic, and emergent pathogens in aquatic systems. Trends Parasitol 29: 567–578. pmid:24091244
  14. 14. Hofer A, Crona M, Logan DT, Sjoberg BM (2012) DNA building blocks: keeping control of manufacture. Crit Rev Biochem Mol Biol 47: 50–63. pmid:22050358
  15. 15. Lynch M, Marinov GK (2015) The bioenergetic costs of a gene. Proc Natl Acad Sci U S A 112: 15690–15695. pmid:26575626
  16. 16. Tozzi MG, Camici M, Mascia L, Sgarrella F, Ipata PL (2006) Pentose phosphates in nucleoside interconversion and catabolism. FEBS J 273: 1089–1101. pmid:16519676
  17. 17. Moffatt BA, Ashihara H (2002) Purine and pyrimidine nucleotide synthesis and metabolism. Arabidopsis Book 1: e0018. pmid:22303196
  18. 18. Heinz E, Hacker C, Dean P, Mifsud J, Goldberg AV, et al. (2014) Plasma membrane-located purine nucleotide transport proteins are key components for host exploitation by microsporidian intracellular parasites. PLoS Pathog 10: e1004547. pmid:25474405
  19. 19. Heinz E, Williams TA, Nakjang S, Noel CJ, Swan DC, et al. (2012) The genome of the obligate intracellular parasite Trachipleistophora hominis: new insights into microsporidian genome dynamics and reductive evolution. PLoS Pathog 8: e1002979. pmid:23133373
  20. 20. Watson AK, Williams TA, Williams BA, Moore KA, Hirt RP, et al. (2015) Transcriptomic profiling of host-parasite interactions in the microsporidian Trachipleistophora hominis. BMC Genomics 16: 983. pmid:26589282
  21. 21. Wylie JL, Wang LL, Tipples G, McClarty G (1996) A single point mutation in CTP synthetase of Chlamydia trachomatis confers resistance to cyclopentenyl cytosine. J Biol Chem 271: 15393–15400. pmid:8663065
  22. 22. Reichard P (1988) Interactions between deoxyribonucleotide and DNA synthesis. Annu Rev Biochem 57: 349–374. pmid:3052277
  23. 23. Kumar D, Abdulovic AL, Viberg J, Nilsson AK, Kunkel TA, et al. (2011) Mechanisms of mutagenesis in vivo due to imbalanced dNTP pools. Nucleic Acids Res 39: 1360–1371. pmid:20961955
  24. 24. Hunsucker SA, Mitchell BS, Spychala J (2005) The 5'-nucleotidases as regulators of nucleotide and drug metabolism. Pharmacol Ther 107: 1–30. pmid:15963349
  25. 25. McClarty G, Tipples G (1991) In situ studies on incorporation of nucleic acid precursors into Chlamydia trachomatis DNA. J Bacteriol 173: 4922–4931. pmid:1907263
  26. 26. Alexander WG, Wisecaver JH, Rokas A, Hittinger CT (2016) Horizontally acquired genes in early-diverging pathogenic fungi enable the use of host nucleosides and nucleotides. Proc Natl Acad Sci U S A.
  27. 27. Houseley J, Tollervey D (2009) The many pathways of RNA degradation. Cell 136: 763–776. pmid:19239894
  28. 28. Garneau NL, Wilusz J, Wilusz CJ (2007) The highways and byways of mRNA decay. Nat Rev Mol Cell Biol 8: 113–126. pmid:17245413
  29. 29. Coller J, Parker R (2004) Eukaryotic mRNA decapping. Annu Rev Biochem 73: 861–890. pmid:15189161
  30. 30. Dziembowski A, Lorentzen E, Conti E, Seraphin B (2007) A single subunit, Dis3, is essentially responsible for yeast exosome core activity. Nat Struct Mol Biol 14: 15–22. pmid:17173052
  31. 31. Lane AN, Fan TW (2015) Regulation of mammalian nucleotide metabolism and biosynthesis. Nucleic Acids Res 43: 2466–2485. pmid:25628363
  32. 32. Dolgikh VV, Sokolova JJ, Issi IV (1997) Activities of enzymes of carbohydrate and energy metabolism of the spores of the microsporidian, Nosema grylli. J Eukaryot Microbiol 44: 246–249. pmid:9183713
  33. 33. Bianchi V, Pontis E, Reichard P (1986) Interrelations between substrate cycles and de novo synthesis of pyrimidine deoxyribonucleoside triphosphates in 3T6 cells. Proc Natl Acad Sci U S A 83: 986–990. pmid:3456577
  34. 34. Cass CE, Young JD, Baldwin SA, Cabrita MA, Graham KA, et al. (1999) Nucleoside transporters of mammalian cells. Pharm Biotechnol 12: 313–352. pmid:10742981
  35. 35. Dean P, Major P, Nakjang S, Hirt RP, Embley TM (2014) Transport proteins of parasitic protists and their role in nutrient salvage. Front Plant Sci 5: 153. pmid:24808897
  36. 36. Greub G, Raoult D (2003) History of the ADP/ATP-translocase-encoding gene, a parasitism gene transferred from a Chlamydiales ancestor to plants 1 billion years ago. Appl Environ Microbiol 69: 5530–5535. pmid:12957942
  37. 37. James TY, Pelin A, Bonen L, Ahrendt S, Sain D, et al. (2013) Shared signatures of parasitism and phylogenomics unite cryptomycota and microsporidia. Curr Biol 23: 1548–1553. pmid:23932404
  38. 38. Tsaousis AD, Kunji ER, Goldberg AV, Lucocq JM, Hirt RP, et al. (2008) A novel route for ATP acquisition by the remnant mitochondria of Encephalitozoon cuniculi. Nature 453: 553–556. pmid:18449191
  39. 39. Campbell SE, Williams TA, Yousuf A, Soanes DM, Paszkiewicz KH, et al. (2013) The genome of Spraguea lophii and the basis of host-microsporidian interactions. PLoS Genet 9: e1003676. pmid:23990793
  40. 40. Williams BA, Hirt RP, Lucocq JM, Embley TM (2002) A mitochondrial remnant in the microsporidian Trachipleistophora hominis. Nature 418: 865–869. pmid:12192407
  41. 41. Reinke AW, Troemel ER (2015) The Development of Genetic Modification Techniques in Intracellular Parasites and Potential Applications to Microsporidia. PLoS Pathog 11: e1005283. pmid:26720003
  42. 42. Haferkamp I, Schmitz-Esser S, Wagner M, Neigel N, Horn M, et al. (2006) Tapping the nucleotide pool of the host: novel nucleotide carrier proteins of Protochlamydia amoebophila. Mol Microbiol 60: 1534–1545. pmid:16796686
  43. 43. Knab S, Mushak TM, Schmitz-Esser S, Horn M, Haferkamp I (2011) Nucleotide parasitism by Simkania negevensis (Chlamydiae). J Bacteriol 193: 225–235. pmid:20971898
  44. 44. Tjaden J, Haferkamp I, Boxma B, Tielens AG, Huynen M, et al. (2004) A divergent ADP/ATP carrier in the hydrogenosomes of Trichomonas gallinae argues for an independent origin of these organelles. Mol Microbiol 51: 1439–1446. pmid:14982636
  45. 45. Trentmann O, Horn M, van Scheltinga AC, Neuhaus HE, Haferkamp I (2007) Enlightening energy parasitism by analysis of an ATP/ADP transporter from chlamydiae. PLoS Biol 5: e231. pmid:17760504
  46. 46. Haferkamp I, Schmitz-Esser S, Linka N, Urbany C, Collingro A, et al. (2004) A candidate NAD+ transporter in an intracellular bacterial symbiont related to Chlamydiae. Nature 432: 622–625. pmid:15577910
  47. 47. Malyshev DA, Dhami K, Lavergne T, Chen T, Dai N, et al. (2014) A semi-synthetic organism with an expanded genetic alphabet. Nature 509: 385–388. pmid:24805238
  48. 48. Xie H, Patching SG, Gallagher MP, Litherland GJ, Brough AR, et al. (2004) Purification and properties of the Escherichia coli nucleoside transporter NupG, a paradigm for a major facilitator transporter sub-family. Mol Membr Biol 21: 323–336. pmid:15513740
  49. 49. Hacker C, Howell M, Bhella D, Lucocq J (2014) Strategies for maximizing ATP supply in the microsporidian Encephalitozoon cuniculi: direct binding of mitochondria to the parasitophorous vacuole and clustering of the mitochondrial porin VDAC. Cell Microbiol 16: 565–579. pmid:24245785
  50. 50. Scanlon M, Leitch GJ, Visvesvara GS, Shaw AP (2004) Relationship between the host cell mitochondria and the parasitophorous vacuole in cells infected with Encephalitozoon microsporidia. J Eukaryot Microbiol 51: 81–87. pmid:15068269
  51. 51. Ronnebaumer K, Gross U, Bohne W (2008) The nascent parasitophorous vacuole membrane of Encephalitozoon cuniculi is formed by host cell lipids and contains pores which allow nutrient uptake. Eukaryot Cell 7: 1001–1008. pmid:18408058
  52. 52. Francione L, Smith PK, Accari SL, Taylor PE, Bokko PB, et al. (2009) Legionella pneumophila multiplication is enhanced by chronic AMPK signalling in mitochondrially diseased Dictyostelium cells. Dis Model Mech 2: 479–489. pmid:19638422
  53. 53. Matsumoto A, Bessho H, Uehira K, Suda T (1991) Morphological studies of the association of mitochondria with chlamydial inclusions and the fusion of chlamydial inclusions. J Electron Microsc (Tokyo) 40: 356–363.
  54. 54. Pernas L, Adomako-Ankomah Y, Shastri AJ, Ewald SE, Treeck M, et al. (2014) Toxoplasma effector MAF1 mediates recruitment of host mitochondria and impacts the host response. PLoS Biol 12: e1001845. pmid:24781109
  55. 55. He Y, Wang K, Yan N (2014) The recombinant expression systems for structure determination of eukaryotic membrane proteins. Protein Cell 5: 658–672. pmid:25119489
  56. 56. Dean P (2011) Functional domains and motifs of bacterial type III effector proteins and their roles in infection. FEMS Microbiol Rev 35: 1100–1125. pmid:21517912
  57. 57. Mueller KE, Plano GV, Fields KA (2014) New frontiers in type III secretion biology: the Chlamydia perspective. Infect Immun 82: 2–9. pmid:24126521
  58. 58. Paldi N, Glick E, Oliva M, Zilberberg Y, Aubin L, et al. (2010) Effective gene silencing in a microsporidian parasite associated with honeybee (Apis mellifera) colony declines. Appl Environ Microbiol 76: 5960–5964. pmid:20622131
  59. 59. Huang Q, Evans JD (2016) Identification of microRNA-like small RNAs from fungal parasite Nosema ceranae. J Invertebr Pathol 133: 107–109. pmid:26678507
  60. 60. Kanehisa M, Goto S, Kawashima S, Nakaya A (2002) The KEGG databases at GenomeNet. Nucleic Acids Res 30: 42–46. pmid:11752249