Replication and maintenance of the Plasmodium falciparum apicoplast genome

Accepted Manuscript Title: Replication and Maintenance of the Plasmodium falciparum Apicoplast Genome Author: Morgan E. Milton Scott W. Nelson PII: DOI: Reference:

S0166-6851(16)30079-2 http://dx.doi.org/doi:10.1016/j.molbiopara.2016.06.006 MOLBIO 10997

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Molecular & Biochemical Parasitology

Received date: Revised date: Accepted date:

27-4-2016 14-6-2016 19-6-2016

Please cite this article as: Milton Morgan E, Nelson Scott W.Replication and Maintenance of the Plasmodium falciparum Apicoplast Genome.Molecular and Biochemical Parasitology http://dx.doi.org/10.1016/j.molbiopara.2016.06.006 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Replication and Maintenance of the Plasmodium falciparum Apicoplast Genome Morgan E. Miltona,b and Scott W. Nelsona,1 a

Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University,

Ames IA 50011, USA b

Current Address: Department of Molecular and Structural Biochemistry, North Carolina

State University, Raleigh, NC, 27695, USA 1

To whom correspondence should be addressed.

Email: [email protected] Address: 4112 Molecular Biology Building, 2437 Pammel Dr., Ames, IA 50011 Telephone: (515) 294-3434

Keywords: apicoplast, DNA replication, malaria, drug discovery

Graphical abstract

Highlights 

The discovery of the apicoplast and its genome.

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The apicoplast genome is replicated by a polyprotein containing DNA primase, helicase, and polymerase subdomains.

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The apicoplast may contain pathways for mismatch repair and base excision repair.

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The proteins involved in replication and maintenance of the apicoplast genome could be potent and specific antimalarial drug targets.

Abstract Members of the phylum Apicomplexa are responsible for many devastating diseases including malaria (Plasmodium spp.), toxoplasmosis (Toxoplasma gondii), babesiosis (Babesia bovis), and cyclosporiasis (Cyclospora cayetanensis). Most Apicomplexans contain a unique and essential organelle called the apicoplast. Derived from an ancient chloroplast, the apicoplast replicates and maintains a 35 kilobase (kb) circular genome. Due to its essential nature within the parasite, drugs targeted to proteins involved in DNA replication and repair of the apicoplast should be potent and specific. This review summarizes the

current knowledge surrounding the replication and repair of the Plasmodium falciparum apicoplast genome and identifies several putative proteins involved in replication and repair pathways.

Keywords: apicoplast; DNA replication; malaria; drug discovery

1. Malaria Malaria is a devastating disease that affects nearly 200 million people each year [1]. In 2014, approximately 440 thousand children died due to this mosquito-borne disease before reaching their fifth birthday [1]. The spread of drug-resistant parasites in central Africa, India, southeast Asia, and northern South America is a growing problem, with common antimalarial drugs such as chloroquine and sulfadoxin-pyrimethamine becoming ineffective [2]. Currently no drug is available to which some level of resistance has not developed. Even artemisinin has become less effective across southeast Asia, and it is only a matter of time before those strains make their way to Africa [3]. The development of new and potent antimalarial treatments will be critical in our ability to continue to combat malaria in the future [1]. Due to the critical need for new anti-malarial therapeutics, many avenues are being explored for their potential as anti-malarial drug targets. The leading cause of malaria in humans is Plasmodium falciparum, a member of the phylum Apicomplexa, which houses some of the most common and deadly protistan parasites. Many members of the phylum contain an essential organelle called the apicoplast [4]. Due to its exclusivity to Apicomplexa and its strict requirement for parasite survival, drugs targeted against apicoplast function should be potent and specific. 2. Origins of the Apicoplast The history of the discovery and initial investigation of the apicoplast has been convoluted. The presence of an unidentified organelle and an extrachromosomal DNA in malarial parasites bewildered parasitologists for over two decades. A 35 kb circular DNA was originally observed through electron micrographs in 1974, and was thought to originate from the mitochondrial genome of the parasite [5]. Around the same time, an intracellular

organelle of unknown function was detected in electron micrographs of several different apicomplexans [6]. Evidence began to build in favor of a plastidic origin of both this “spherical body” and the 35 kb DNA [7]. Nearly 15 years after the misidentification of the apicoplast genome as originating from the mitochondria, the true mitochondrial genome was found [8], and subsequent work showed that the 35 kb genome belonged instead to the apicoplast [9]. Full sequencing of the apicoplast genome in 1996 established that its organization and gene content are reminiscent of an algae plastid [10]. To account for the presence of a plastidic-like organelle which is surrounded by an average of four membranes (as opposed to the two membranes found in typical plastids), a precursor to apicomplexans must have acquired the plastid through a secondary endosymbiosis event. The identity of the players involved in the creation of the apicoplast was debated for some time, but more recent phylogenetic analysis has provided definitive evidence that the apicoplast originated from a photosynthetic red algae [11–13] that was engulfed and retained by an ancestral dinoflagellate through a secondary endosymbiosis event [14]. The apicoplast has since discarded its photosynthetic abilities, but is still maintained within the parasite, presumably due to its role in isoprenoid biosynthesis, heme and fatty acid biosynthesis, and iron-sulfur cluster maturation [15]. 3. Apicoplast Genome Structure The apicoplast genome is of low complexity and primarily encodes genes involved in its own expression. The apicoplast has one of the most A/T-rich genomes known to date with 86.9% A/T [13]. It contains 68 genes coding for the large and small subunit rRNAs, a minimal but complete set of tRNAs, ribosomal proteins, three subunits of a bacterial-like RNA polymerase, and several protein chaperones [16]. A series of inverted repeats contain duplicate genes for rRNAs and tRNAs [13]. It is believed that the apicoplast genome has been under high selective pressure, consequently reducing the genome from the 150 kb of a typical chloroplast to 35 kb [16]. This has resulted in many deletions and rearrangements to conserve the minimal autonomy of the apicoplast [16]. This review will focus on how the apicoplast genome is replicated and maintained. Each of the known proteins involved in replication, as well as the proteins that are possibly involved in its repair will be examined. There are still many unknowns regarding the replication and repair of the apicoplast genome;

however, it is clear that some of the proteins responsible for these processes have a high potential as new antimalarial drug targets. 4. Method of Replication Understanding the means by which the P. falciparum apicoplast genome is replicated could lead to the development of new antimalarial compounds with novel mechanisms of action for which resistance has not yet arisen. Replication initiates within a large inverted repeat region through twin single-stranded displacement loops (D-loops) [17]. This method of replication has been shown to be sensitive to the topoisomerase inhibitor ciprofloxacin (CIP) [18]. At other locations on the genome, replication takes place through a rolling-circle mechanism, which is less sensitive to CIP [17]. Replication of the apicoplast genome initiates in the late trophozoite stage of the parasite life cycle within the red blood cell. Replication begins slightly before that of the nuclear genome and no more than 3 copies of plastid DNA are observed per cell [17]. The relationship between D-loop and rolling circle is unclear. When parasites are subjected to non-lethal doses of CIP, the amount of 35 kb linear DNA increases but the ratio of replication bubbles to replication forks decreases. This suggests the two mechanisms are truly theta and sigma varieties and are independent from each other as has been observed in the chloroplasts of other species [17]. 5. Proteins Involved in DNA Replication Typical genomic replication requires a helicase to unwind the double-stranded DNA, a primase to synthesize RNA primers for Okazaki fragment initiation, and a polymerase to replicate the leading and lagging stand templates. Single-stranded binding (SSB) proteins are also essential and aid in replication by protecting exposed single stranded DNA, melting DNA secondary structure, and interacting with proteins in the replisome. Topoisomerases are required to relieve positive supercoils that are generated ahead of the replication fork and are used to separate interlocked strands of replicated circular DNA. To date, all of these proteins have been identified and, to some extent, biochemically characterized. This review examines what is known about each of these proteins and their involvement in the replication and maintenance of the P. falciparum apicoplast genome.

5.1 Prex (transcription, transport, and processing) The P. falciparum gene PF14_0112 is 6051 bp long, contains no introns, and resides on the 14th chromosome of the nuclear genome. It was originally annotated as pom1, for “polymerase of malaria,” when a portion of the open reading frame was identified in P. chabaudi during a screen for antigenic variation in the murine parasite [19]. The gene has since been renamed Prex for Plastidic DNA Replication/repair Enzyme Complex [20]. The Prex gene has a single open reading frame encoding DNA primase, helicase, and polymerase (Fig. 1). A 20 amino acid long targeting sequence ensures that the gene is co-translated as a single polyprotein into the ER lumen, where it is then transported as a polyprotein into the apicoplast [21–23]. The Prex gene is thought to contain the only DNA polymerase targeted to the apicoplast, and thus it is likely responsible for both DNA replication and repair [24]. The parasite maintains low levels of Prex expression throughout its lifecycle with the maximum amount occurring during the late trophozoite stage and subsequently declining in the schizont stages [20]. Increased expression of Prex is correlated with an increase in the amount of apicoplast DNA [17], consistent with its proposed role in the replication of the apicoplast genome (Fig. 2). Lindner et al. were unable to generate a Prex knockout in P. yoelli (a mouse model for malaria), strongly suggesting Prex is an essential protein in the blood stage of parasite infection [25]. Several research groups have demonstrated that the targeting sequence of Prex is specific to the apicoplast. Fusing the Prex targeting sequence to GFP causes it to be transported into the apicoplast without any transport into other subcellular compartments [20]. Subcellular localization of Prex was further confirmed when GFP was fused to the Cterminus of the primase domain and its localization inside the apicoplast was observed throughout the parasite’s life cycle in the red blood cell [25]. Using immunofluorescence, antibodies to the endogenously expressed Prex helicase and apicoplast SSB were found to co-localize to the apicoplast in both early and late stages of the parasite’s growth within a red blood cell [26]. Upon entering the apicoplast, Prex is post-translationally modified by unknown protease(s) [21]. Bioinformatic analysis suggests there are 21 proteases predicted to be targeted to the apicoplast, but their protein substrates and recognition sequences are unknown [27]. Western blot analysis has identified numerous cleavage products of Prex, although it is

not clear which represent functional proteins. It is possible there are several forms of the mature active Prex proteins, as multiple isoforms of the bacteriophage T7 primase-helicase have been observed [28]. Identification of the protease(s) responsible for separating the Prex primase, helicase, and polymerase domains would lead to a better understanding of posttranslational modifications of apicoplast proteins and identify potential new drug targets. 5.1.1 Primase The Prex primase domain clusters phylogenetically with bacteriophage T7-like DNA primases [20]. Bacteriophage T7 contains a bi-functional primase-helicase protein which forms a hexamer with distinct, but attached, primase and helicase domains [29,30]. It is not clear if, in the active form of the protein, the Prex primase protein remains covalently attached to the helicase, or if the primase and helicase domains are proteolytically separated. Bacterial primases, which are not fused to their partner helicase, generally have a three domain

organization: zinc-binding domain (ZBD),

topoisomerase-primase domain

(TOPRIM), and a helicase-binding domain (HBD). Bacteriophage T7 primase does not have an HBD, presumably because it is fused directly to the helicase, and therefore complex formation is not required. Similarly, Prex primase contains ZBD and TOPRIM domains but no HBD is detected [25], indicating that if the primase is indeed separated from the helicase the two proteins must interact in a novel fashion. Seow et al. used heterologous expression in E. coli to produce the primase and helicase domains of Prex fused as a single polypeptide (Fig. 1). The purified protein contained both Mg2+-dependent primase and helicase activities [20]. This indicates that the native primase and helicase domains of Prex are capable of remaining as a single bifunctional protein in the apicoplast, but there is strong evidence indicating that they primarily exist as separated proteins. Lindner et al. expressed a protein containing just the primase ZBD and TOPRIM domains without the helicase domain (residues 115 to 465, Fig. 1) [25]. The PfPrex primase construct bound Zn2+ in a 1:1 molar ratio and catalyzed DNA dependent RNA synthesis, creating > 20 nt long RNA primers in the absence of other replisomal components [25]. Most importantly, Western blot analysis of parasite extract using antibodies to the TOPRIM domain revealed protein with a molecular weight of ~55 kDa, suggesting that the primase is cleaved from the helicase. Western blots reveal other minor protein bands with

molecular weights of ~80 and ~160 kDa, which could be uncleaved primase-helicase domains and larger fragment Prex, respectively [25]. 5.1.2 Helicase Like the Prex primase, the helicase is homologous to bacteriophage T7 phage helicase, which is a member of the helicase superfamily 4 [20]. All helicases bind and hydrolyze nucleotides, and the Prex helicase has the Walker A and B motifs required for ATPase activity. Based on its homology to T7 helicase, it is likely that Prex helicase forms a hexamer ring and translocates in a 5ʹ to 3 ʹ direction along the single-stranded DNA that is threaded through the center of its ring [31]. Bhowmick et al. expressed the Prex helicase (residues 669-1000, Fig. 1) fused to a maltose-binding protein (MBP) solubility tag [26]. The purified helicase-MBP fusion protein was functional and its unwinding activity increased in the presence of P. falciparum SSB. The enhancement of unwinding activity is specific to Plasmodium SSB, suggesting a direct physical interaction between the proteins. The potential protein-protein interaction was confirmed in pulldown assays where the MBP-tagged Prex helicase pulled-down recombinant and endogenous SSB. To identify the domain that was primarily responsible for the interaction, SSB was separated into two domains and it was found that an N-terminal construct of SSB (residues 77-200), but not a C-terminal construct (residues 200-284) interacted with the helicase [26]. Interestingly, although the C-terminal domain of SSB does not form a stable interaction with the helicase, it was found to be necessary for the stimulation of DNA unwinding activity [26]. It is common to observe helicases interacting with their SSB counterparts [32,33]. E. coli PriA, a protein responsible for restarting a stalled replication fork, has been shown to be loaded onto the DNA by SSB [34,35]. To date, no homolog to a helicase loader has been identified for the apicoplast. It is conceivable that the interaction between the Prex helicase and SSB could be reminiscent of the PriA-SSB interaction, resulting in the apicoplast SSB acting as a helicase loader for the Prex helicase. Alternatively, the Prex primase domain may act as a helicase loader, as it does in the bacteriophage T7 fused primase-helicase protein [36,37]. Western blots employing polyclonal antibodies raised to the helicase identified two prominent bands at 70 kDa and 55 kDa, which were attributed to different helicase isoforms,

similar to what is seen in T7 bacteriophage [26,38]. The 70 kDa band is present throughout the blood stage of parasite infection, whereas the 55 kDa band appears only during and after the trophozoite stage [26]. Neither of these bands correspond in size to the Bhowmick et al. recombinant protein, which has a theoretical molecular weight of 38 kDa, suggesting that the starting and/or ending points chosen for the Prex helicase construct are not those found in the native protein. This highlights the importance of identifying the Prex proteases and their cleavage sites, as the functional properties of the native proteins could differ from the truncated counterparts. 5.1.3 Polymerase The apicoplast DNA polymerase is the most thoroughly investigated Prex domain. Genetic analysis classifies the polymerase as a prokaryotic A-family DNA polymerase, which are divided into five clades: prototypical bacterial PolAs, thermophilic viruses, Aquificaceae and Hydrogenothermaceae, Apicomplexa, and other viral-like bacterial polymerases that are found in bacteria that also contain typical PolAs [39]. There is evidence that the atypical A-family polymerase originated through lateral gene transfer when a thermophilic virus infected a cyanobacterium, transferring its polA gene to the host. It is likely that the transfer occurred prior to the second endosymbiosis event that gave rise to the apicoplast [39]. Typical PolAs contain a 5’ to 3’ DNA polymerase domain, 3ʹ-5ʹ exonuclease proofreading domain, and a 5ʹ-3ʹ exonuclease domain for removing RNA primers from Okazaki fragments [40]. Members of Apicomplexa have lost the 5ʹ-3ʹ exonuclease domain, and instead are fused to the primase and helicase domains. DNA polymerases often contain accessory proteins that act as processivity factors. Currently there is no known processivity factor associated with the apicoplast DNA polymerase. If a processivity factor does exist, it is likely to be substantially different than known processivity factors since no obvious homologs with apicoplast targeting sequences have been identified in the Plasmodium genome. However, based on the relatively small size of the apicoplast genome and the time required for its replication, it may be possible for apPOL to fully replicate the 35 kb genome without the assistance of a processivity factor. Several labs have biochemically characterized the apicoplast DNA polymerase. One complication has been that the native start site of the polymerase domain is unknown. Thus far, three different apicoplast DNA polymerase constructs have been investigated. Seow et al.

first expressed a protein construct encompassing residues 1107 to 2016 (Fig. 1), which was named PfPREXpol [20]. Kennedy et al. employed a construct expressing the polymerase domain beginning at residue 1426 (Fig. 1), which was referred to as KPom1 based on sequence similarity to the Klenow fragment of E. coli Pol I [24]. Most recently, the Nelson laboratory designed a construct based on sequence alignments (referred to as apPOL) and identified a possible polymerase boundary spanning residues 1389 to 2016 that is conserved across the Plasmodium genus [41,42] (Fig. 1). Below, the biochemical properties of each of these polymerase constructs is summarized. Protein expression levels vary greatly between protein constructs. PfPREXpol was reported to yield 4 mg of purified recombinant protein per liter of culture [20], whereas the yield of apPOL is 50 mg of pure protein per liter (although, these constructs were expressed in different E. coli strains) [43]. The total yield of KPom1 was not reported, and the protein was expressed with a maltose-binding-protein (MBP) fusion tag as opposed to the hexahistidine tags used for PfPREXpol and apPOL [24]. The MBP fusion tag may serve to increase the solubility of KPom1, as we have found that the expression and stability of the apicoplast polymerase is very sensitive to modifications at the N-terminal region. Upon solving the crystal structure of apPOL (PDB 5DKU), it became clear why removal of the Nterminal region impacts stability. The polymerase N-terminal region forms a long β-hairpin that spans the bottom of the exonuclease domain (distal to the polymerase domain). The βhairpin and flanking loops are involved in numerous hydrophobic and hydrogen bonding interactions with the rest of the exonuclease domain. The start site of KPom1 disrupts the βhairpin and excludes a majority of these interactions, which could result in a large unstructured loop at the N-terminus of the construct. It is possible that the differences in construct start sites are responsible for the differences in polymerase activity discussed below. Biochemical assays examining the polymerase and exonuclease activities, as well as the optimal pH, Mg2+ concentrations, and temperature have been carried out. PfPREXpol and KPom1 have maximal DNA polymerase activity at Mg2+ concentrations of 4 and 10 mM, respectively [20,24]. KPom1 has pH and temperature optimums of 9.0 and 40°C, respectively, whereas PfPREXpol optimum values are 7.0 and 75° C, respectively [20,24]. It is unclear if differences in construct alone can account for these discrepancies. The disparity

in temperature tolerated by the polymerase is particularly interesting. The temperature where PfPREXpol shows the highest activity (75° C) is consistent with the thermophilic virus ancestry of the polymerase. On the other hand, the temperature optimum of 40°C for KPom1 is more physiologically relevant, as the replication of the apicoplast genome takes place in infected red blood cells at 37oC [17]. Our findings also suggest that apPOL operates at an optimal temperature that is close to 37°C. We find that apPOL has a thermal denaturation temperature of about 45°C, and at temperatures higher than 45°C the activity significantly decreases and cannot be recovered upon returning to lower temperatures (unpublished observations). Since the native apicoplast DNA polymerase boundaries are unknown, it is difficult to tell what structural features may be causing the differences in temperature tolerance observed between PfPREXpol and KPom1/apPOL. Kennedy et al. determined the fidelity of KPom1, and found that it was surprisingly error-prone, but that the exonuclease domain corrected a majority of its mistakes [24]. It was concluded that without an active exonuclease domain, KPom1 has a unique error signature with a mutation spectrum more closely resembling lesion bypass polymerases than replicative polymerases [39]. KPom1 was found to have an especially strong bias towards misincorporating a dGTP across from a dTMP [24]. The apPOL construct behaves more like the E. coli Klenow fragment, and its fidelity is reminiscent of other replicative DNA polymerases [41]. Differences in the reported fidelity of KPom1 and apPOL may not be solely due to the structural variances between the constructs, as the methods of analysis differed between the two proteins [41]. Mutation rates are not exclusively determined by polymerase fidelity, as the ability of a polymerase to extend a mismatch and removal of misincorporated nucleotides also plays a role. Mismatch extension by apPOL is highly sequence dependent. The incoming nucleotide must compensate for an altered templating base position caused by upstream mismatches. By combining misincorporation, exonuclease, and mismatch extension rate data, the most likely mutations caused by apPOL were ranked. apPOL is most likely to mutate a G to an A if the downstream base is a T, followed by C replacing a T if a T, C, or G is downstream [41]. Of course, these observations do not take into account potential DNA repair enzymes which could impact mutation probabilities in the apicoplast. The high fidelity of apPOL is not reminiscent of a lesion bypass polymerase, and we find that apPOL stalls at common DNA

lesions (unpublished observations). This suggests that DNA repair pathways may be functional within the apicoplast. Although there is some contradictory data surrounding the details, the results of these three groups are in strong agreement that the Prex polymerase domain is the polymerase responsible for replication of the entire apicoplast genome. Determination of the native construct would aid in understanding the discrepancies in polymerase activities between PfPREXpol, KPom1, and apPOL. Until then, the structure of apPOL will act as a starting point for further understanding of this atypical A-family polymerase and aid in the development of new antimalarial compounds (PDBs: 5DKU and 5DKT). 5.2 SSB Single stranded binding proteins (SSB, gene ID PF3D7_0508800) are responsible for protecting and stabilizing ssDNA during replication and repair, and for melting secondary structures in the DNA. Additionally, SSBs from other organisms have been shown to interact with numerous proteins involved with replication and repair, and may act to coordinate these processes [33]. The gene encoding the P. falciparum apicoplast SSB (PfSSB) resides on chromosome five of the nuclear genome and is of bacterial origin [44]. Although E. coli SSB shares only 39% and 66% sequence identity and similarity, respectively, with PfSSB, the two proteins share a high degree of structural similarity. The N-terminal region of E.coli SSB is composed of an oligonucleotide-binding domain (OBD) that contains three tryptophan residues that base stack with ssDNA (45). PfSSB maintains these protein-ssDNA interactions by positioning these residues in an identical manner [44,45]. The OBD of PfSSB also contains a conserved histidine that facilitates oligomerization [44,46]. Prokaryotic SSBs have an acidic C-terminal tail that is involved in mediating protein-protein interactions [33]. PfSSB lacks this acidic sequence, suggesting that its binding partners are different and likely specific to apicoplast function [44,47]. In addition to having a distinct tail sequence, PfSSB has a 28 residue extension on its C-terminus that has no sequence homology to its E. coli counterpart [44]. Removal of these 28 residues results in an increase in helicase unwinding activity in the presence of SSB, suggesting that the extreme C-terminus of SSB could be involved in helicase regulatory functions [26]. Alternatively, Antony et al. found that removal of these residues shifted the ssDNA binding equilibrium to favor a partially wrapped mode [47]. This indicates that the increased stimulation of helicase unwinding activity may

be due to the altered DNA binding mode, rather than the loss of a regulatory interaction between SSB and the helicase. Unfortunately, the crystal structure does not shed light on the possible mechanism of helicase activation, as only the residues in the OBD (residues 77-194) were observed, with no visible electron density for the C-terminal region. Prusty et al. demonstrated that PfSSB localizes to the apicoplast and that it is expressed during all three stages of the parasite’s life cycle in the red blood cell (Fig. 2) [44]. Native and recombinantly expressed protein form a stable tetramer in solution and in the crystal [44,47,48]. Gel shift assays along with a co-crystal structure with DNA, confirm that PfSSB specifically binds ssDNA [44,48]. Despite being nearly identical in structure to the E. coli SSB [48], PfSSB is unable to complement E. coli SSB in vivo, and removal of the 28 residue C-terminal extension does not induce complementation [44]. The inability to complement is likely due to the loss of specific protein-protein interactions between SSB and other DNA replication/repair proteins. Additionally, the PfSSB crystal structure revealed that the mechanism of ssDNA binding differs from that of E. coli. While PfSSB wraps ssDNA with the same topology as E. coli, the DNA backbone polarity is reversed [48]. There are other notable differences between the two proteins: protein-DNA contacts, symmetry of inter-subunit contact sites, and protein-protein interactions between adjacent tetramers. PfSSB only binds ssDNA in a tight binding, fully wrapped mode, while E. coli SSB is able to bind in full and partially wrapped modes [47]. Even in a fully wrapped mode, PfSSB is able to fully diffuse along the ssDNA and melt DNA secondary structures [47]. A different interface is observed for tetramer-tetramer interactions for PfSSB than those seen in E. coli. In E. coli, loops from neighboring tetramers pack against each other, but this interface is not seen in PfSSB. All of these differences combined could result in an SSB that is functionally different from that of E. coli. 5.3 Topoisomerase II Normally only found in bacteria, P. falciparum has a DNA gyrase that is imported into the apicoplast. The P. falciparum gyrase (PfGyr) is a bacterial type IIA topoisomerase composed of an A2B2 complex that catalyzes DNA supercoiling in an ATP-dependent manner [49]. The A subunit, GyrA (gene ID PFIT_1223000), is composed of an N-terminal DNA breakage and union domain, and a C-terminal domain involved in DNA wrapping [50]. GyrB (gene ID PFIT_1239200), the B subunit, contains an N-terminal ATPase domain, and a

C-terminal domain that binds DNA and likely interacts with GyrA [51]. Both PfGyrA and GyrB have apicoplast targeting sequences at their N-terminus and have been shown to localize to the apicoplast through immuno-colocalization assays [52]. Both proteins are expressed during the trophozoite and schizont stages when the apicoplast genome is being replicated, suggesting involvement with replication of the apicoplast DNA (Error! Reference source not found.) [17,52]. Dar et al. demonstrated that PfGyrB could complement its E. coli equivalent in vivo using a temperature sensitive E. coli strain. On the other hand, E. coli could not express full length PfGyrA leading to an inability to test for complementation of that subunit [52]. Expression of recombinant PfGyrA and PfGyrB has proven difficult, but full length PfGyrB and separate domains of PfGyrA, PfGyrAN and PfGyrACC have been obtained [52]. Since gyrase forms a dimer of dimers, Dar et al. examined the dimerization potential of the two subunits. GyrB proteins usually dimerizes in an ATP-dependent manner which facilitates the opening and closing of the dimer around DNA [53]. On the contrary, gel filtration and glutaraldehyde cross-linking indicate that full length PfGyrB and several N-terminal truncation mutants do not require ATP to form stable dimers [52]. A conserved isoleucine has been shown to be essential in dimer formation in E. coli, but mutation of the isoleucine in P. falciparum has no impact on dimerization [54]. It is likely that the protein-protein interface between the GyrB subunits of E. coli and P. falciparum are different, allowing PfGyrB to maintain a dimeric form in solution independent of ATP [52]. The C-terminus of GyrA in E. coli has been shown to be responsible for dimerization, but P. falciparum has no homology to this region [55]. Instead, PfGyrA has a leucine heptad repeat that is predicted to form a coiled-coil. The C-terminal GyrA construct, GyrACC, is able to form a dimer, suggesting that this region is responsible for dimerization of full length GyrA [52]. While Dar et al. did not test for hetero-tetramer formation, cleavage was dependent on the presence of both PfGyrAN and PfGyrB, suggesting that a complex does form [52]. Gyrases utilize ATPase activity to convert the chemical energy of ATP into the mechanical force needed to introduce superhelical turns into DNA [56]. PfGyrB shows intrinsic ATPase activity that is linearly-dependent on protein concentration and is further stimulated by the presence of DNA or PfGyrA. This differs from E. coli GyrB, which displays a nonlinear pattern of ATP hydrolysis [52]. It is probable that the differences in

activity are caused by the modification in dimerization potential, as E. coli GyrB forms dimers in an ATP dependent fashion, whereas PfGyrB does not [52]. A functional gyrase must cleave dsDNA to introduce or alleviate supercoils, and PfGyrAN and PfGyrB together are able to cleave supercoiled DNA. PfGyrB is also able to form a functional cleavage complex with E. coli GyrA [52]. Dar et al. found that the topoisomerase II inhibitor, ciprofloxacin, also inhibits the apicoplast gyrase. CIP traps topoisomerase on the DNA, resulting in broken DNA, and can induce further cleavage of DNA [57]. P. falciparum gyrase has increased cleavage activity in a dose-dependent manner, supporting the fact that PfGyrAN and PfGyrB form a functional enzyme [52]. Finally, a gyrase needs to be able to reintroduce supercoils into circular DNA. PfGyrAN and PfGyrB could not induce negative supercoiling. Adding E. coli GyrA to P. falciparum GyrB led to partial activity with limited supercoiling activity [52]. These results suggest that full-length GyrA is required for the formation of supercoils and that, although E. coli and P. falciparum gyrases perform similar functions and to some degree complement each other, their relative activities are very different [52]. Subsequent work is consistent with this conclusion. The C-terminus of GyrA is involved in wrapping the DNA substrate to ensure the correct “handedness” of negative supercoiling, therefore it is not surprising that removal of this domain reduces supercoiling activity [58]. There are many differences between the C-terminal sequence of PfGyrA and other known gyrases. Specifically, the DNA wrapping domain is generally composed of six β-pinwheel motifs that form blades; P. falciparum lacks 3 of these blades in favor of a Plasmodium-specific insertion [58]. Nagano et al. cloned a PfGyrA C-terminal construct, PfGyrA-CTD, in order to further investigate the importance of the C-terminal domain and its role in the holoenzyme. Using fluorescence anisotropy, they determined that the Kd-DNA of GyrA-CTD was significantly lower than the E. coli equivalent (0.2 and 1 µM, respectively). They also demonstrated that GyrA-CTD was able to introduce writhe into nicked circular plasmid [58]. Between Dar et al. and Nagano et al., all necessary topoisomerase activity has been detected for the P. falciparum GyrA and GyrB.

5.4 DNA ligase (Lig1) Ligases are responsible for joining the phosphodiester backbone of adjacent, mature DNA fragments during replication and are necessary for DNA replication and repair. DNA ligases perform their phosphoryl transfer reaction in an ATP (eukaryotes, bacteriophage, and archaea) or NAD+-dependent (bacteria) manner to produce a sealed DNA backbone [59]. P. falciparum DNA Ligase I (PfLigI, gene ID PF3D7_1304100) has a catalytic-core displaying high homology to eukaryotic DNA ligase I, and appears to be the only ligase encoded for by the P. falciparum genome [60]. Western blot analysis of cell lysate detects PfLigI only in schizont stage of parasite growth, suggesting that PfLigI is present only when the parasite is actively replicating (Fig. 2). Buguliskis et al. expressed a PfLigI construct (residues 128 to 912) that is able to ligate nicked DNA at a similar rate to T4 DNA ligase, is optimally active at 37°C and pH 8.5, and is sensitive to salt concentration [60]. PfLigI ligation activity was highly stimulated by the addition of ATP but no activity was observed with NAD+, establishing that PfLigI is an ATP-dependent enzyme. Based on kinetic properties, PfLigI is comparable to human ligase I with a kcat of 2.3 x 10-3 s-1. They concluded that PfLigI is a high fidelity ligase that strongly discriminates between correct and incorrect base pairing at the nick site [60]. Interestingly, PfLigI has no PCNA binding domain, but instead has an apicoplast targeting sequence at its N-terminus [60]. PCNA, a eukaryotic DNA clamp that acts as a scaffold and processivity factor for replication and repair proteins, is known to interact with DNA ligase I [61]. Of seven Apicomplexan parasites screened by Buguliskis et al., none contained PCNA binding domains [60]. They propose that either PfLigI has a novel PCNA binding domain and interacts with the Plasmodium PCNA homolog, or that PfLigI does not interact with PCNA at all [60]. Due to its nature as the sole DNA ligase in P. falciparum, it is likely that PfLigI is responsible for replication and repair of both the apicoplast and nuclear genome. As such, PfLigI is required to interact with both prokaryotic (apicoplast) and eukaryotic (nuclear) replication systems, which may account for the lack of a PCNA binding domain.

5.5 5ʹ-3ʹ exonuclease 5ʹ to 3ʹ exonucleases are responsible for removing the RNA primers synthesized by primase so that they can be replaced with DNA and ligated. In typical A-family polymerases, an N-terminal 5ʹ to 3ʹ exonuclease domain performs this activity, but the Prex polymerase lacks such a domain, and instead has an N-terminal primase and helicase domain [39]. The 5ʹ to 3ʹ exonuclease domain can function independently of the 3ʹ to 5ʹ exonuclease domain and polymerase domain [62]. In fact, thermophilic virus PolAs frequently have a separate gene which encodes a 5ʹ to 3ʹ exonuclease [39]. We have identified the putative apicoplast targeted 5ʹ to 3ʹ exonuclease on chromosome 7 of the nuclear genome of P. falciparum (gene ID PF3D7_0203900). BLASTP analysis reveals a DNA Pol I 5ʹ to 3ʹ exonuclease domain with predicted metal binding sites. We propose that this is the 5ʹ to 3ʹ exonuclease involved in DNA replication within the apicoplast and will act in conjunction with the Prex polymerase. 6. Repair of the Apicoplast Genome Proper maintenance of genomic DNA is vital for cellular integrity, and it is likely that the apicoplast genome undergoes DNA repair. The genomic database for Plasmodium, PlasmoDB, suggests several candidate proteins with apicoplast targeting sequences that may be involved in DNA repair [63]. A MutS homolog (gene ID PF3D7_1405400) in the P. falciparum nuclear genome contains a predicted apicoplast targeting sequence. MutS is involved in the mismatch repair pathway, which resolves misincorporated nucleotides as well as some damaged bases [64]. MutS recognizes the damage and initiates the binding of MutL to the mismatch site, which in turn recruits MutH and UvrD [65]. However, there are no obvious homologs of MutL, MutH, or UvrD in the P. falciparum genome that contain apicoplast targeting sequences. It is unclear if the apicoplast equivalent of this prokaryotic mismatch repair pathway have just not been identified within the genome, or if some system is in place to utilize the proteins from the nuclear and mitochondrial genome repair pathways. The apicoplast appears to have a nearly complete base excision repair (BER) pathway. P. falciparum lacks homologs for short-patch BER and it has been shown that

abasic lesions are exclusively repaired through the long-patch DNA BER pathway [66]. Long-patch

BER

requires

a

N-glycosylase/DNA

lyase,

an

apurinic/apyrimidinic

endonuclease, a DNA polymerase that can perform strand-displacement synthesis, a 5ʹ to 3ʹ exonuclease to remove the DNA flap, and a ligase to seal the DNA (Fig. 3) [67]. Prex polymerase is able to perform the required strand-displacement synthesis reaction, making it a prime candidate for the long-patch BER pathway [43]. An APN1 homolog (gene ID PF3D7_1332600) appears to be the apurinic/apyrimidinic endonuclease and contains an apicoplast targeting sequence. In addition to the putative APN1 homolog, members of the Afamily of DNA polymerases have been shown to contain AP lyase activity, although because this activity is specific to short-patch BER, the Prex polymerase may not perform this function [68–70]. BER typically involves a variety of different glycosylases depending on the substrate [71]. A DNA glycosylase (gene ID PF3D7_0917100) is predicted to have an apicoplast-targeting peptide, but not a signal sequence [72]. It has been demonstrated that proteins containing both a signal sequence and targeting peptide will enter into the apicoplast, whereas proteins linked only to an apicoplast ACP targeting peptide are targeted to the cytosol [72]. It is possible that sequence variations in specific targeting peptides may be sufficient to drive apicoplast targeting, or a different mechanism for localization may be implemented. Regardless, the subcellular localization of this DNA glycosylase will need to be confirmed. A protein necessary for long-patch BER that appears to be missing is the flap endonuclease. While the P. falciparum genome contains a FEN1 gene, we have found no homologs that are predicted to be transported to the apicoplast. Alternatively the 5ʹ to 3ʹ exonuclease mentioned above or the ExoIII-like AP nuclease (gene ID PF3D7_0305600), which has only an apicoplast targeting peptide, could possibly perform the needed reaction. The final step of the long patch BER pathway is the ligation of the nicked strand, which is presumably carried out by DNA ligase I (section 5.4). In addition to these repair proteins, we have found two putative proteins involved in Holliday junction processing [73,74]: MUS81-like (gene ID PF3D7_1449400) and RuvBlike (gene ID PF3D7_1106000). Due to the inverted repeat region of the apicoplast genome, cruciform structures are frequently observed [17]. The apicoplast may utilize these homologs to resolve the Holliday junctions formed by the inverted repeats. No other proteins involved in double-strand break pathways have been identified.

7.1 Potential for Anti-malarial Drug Target Since its discovery as a unique and essential organelle, the apicoplast has been investigated for its potential as a target for anti-malarial compounds [18]. It was observed that antibiotics that affect protein or nucleic acid synthesis have anti-malarial activity. Interestingly, these compounds induced a “delayed-death” phenotype [18,75]. Delayed-death refers to the phenomenon that upon administration of the drug, the first generation of parasites remain viable, continuing to divide and infect new red blood cells, but the second generation is unable to form functional merozoites [76]. Parasites treated with doxycycline, which inhibits the 70 S ribosome, cannot replicate their apicoplast genomes and were thus nonfunctional. Interestingly, the mitochondria is left unaffected [76]. This strongly suggests that small molecules are able to penetrate the four membranes encompassing the apicoplast and impact proteins involved in apicoplast replication and transcription. 7.2 Current drugs that impact replicative proteins Many antibiotics inhibit proteins involved in transcription and translation of the apicoplast genome: tetracycline, clindamycin, azithromycin, and rifampin [18,77]. Ciprofloxacin and coumermycin inhibit the DNA gyrase of the apicoplast [52,78] and the Prex polymerase is inhibited by chloroquine and suramin [20]. Unfortunately, these drugs are not completely specific to the apicoplast and frequently have high EC50 values. More potent and specific compounds need to be identified. Medicine for Malaria Venture assembled a library of 400 compounds identified through phenotypic screening that inhibit malaria. This library, known as the Malaria Box, is refined from 20,000 compounds identified from four million compounds screened by St. Jude’s Children’s Research Hospital, Novartis, and GlaxoSmithKline [79]. All compounds inhibit malaria in the blood stage and are nontoxic to humans. Bowman et al. identified roughly 40 small molecules from the Malaria Box that have an apicoplast-targeting phenotype based on a parasite growth recovery of at least 60% upon supplementation with the isoprenoid precursor IPP [80]. We have found that one of these 40 compounds potently inhibits the Prex polymerase in vitro with a sub-micromolar IC50 value, strongly suggesting that apPOL is a suitable drug target (unpublished results).

The replication proteins mentioned in this review could be potent and highly specific drug targets. There may be some concern of cross-reactivity between DNA ligase, SSB, and Prex helicase and human cellular functions (as these proteins have distant human homologs), but it is likely that apicoplast-specific features could be exploited. Efforts to identify additional inhibitors of the Prex polymerase are underway [43]. As we gain more biochemical and structural knowledge concerning the proteins involved with replication and maintenance of the apicoplast genome, we begin to elucidate key features that can be employed in our efforts in anti-malarial drug discovery. Acknowledgements

This work was funded by grants from Iowa State University (LAS Signature Research Initiative, 2014) and The Roy J. Carver Charitable Trust (grant No. 10-3603).

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Figure 1.

P. falciparum Prex. Schematic of Prex depicting key motifs as determined by

BLAST. A 20 residue signal sequence (yellow) is followed by an apicoplast targeting sequence of unknown length. The fading of colors between subdomains represents the unknown cleavage sites between the primase, helicase, and polymerase regions. Black bars below the schematic represent the different constructs that have been studied for each region.

Figure 2.

Prex Expression Relative to Plasmodia Life Cycle. Apicoplast DNA is

replicated in late trophozoites. Prex expression is aligned with replication of the apicoplast DNA. Different constructs of the Prex helicase have been detected in the ring stage as well as in trophozoites and schizonts.

Figure 3.

Mechanism for long-patch base excision repair. Each step of the BER

pathway is listed on the left with the possible apicoplast targeted protein responsible for performing that task listed on the right.