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Review
. 2017 Aug;284(16):2604-2628.
doi: 10.1111/febs.14130. Epub 2017 Jul 3.

Proteases as antimalarial targets: strategies for genetic, chemical, and therapeutic validation

Edgar Deu  1
Affiliations
Review

Proteases as antimalarial targets: strategies for genetic, chemical, and therapeutic validation

Edgar Deu. FEBS J. 2017 Aug.

Abstract

Malaria is a devastating parasitic disease affecting half of the world's population. The rapid emergence of resistance against new antimalarial drugs, including artemisinin-based therapies, has made the development of drugs with novel mechanisms of action extremely urgent. Proteases are enzymes proven to be well suited for target-based drug development due to our knowledge of their enzymatic mechanisms and active site structures. More importantly, Plasmodium proteases have been shown to be involved in a variety of pathways that are essential for parasite survival. However, pharmacological rather than target-based approaches have dominated the field of antimalarial drug development, in part due to the challenge of robustly validating Plasmodium targets at the genetic level. Fortunately, over the last few years there has been significant progress in the development of efficient genetic methods to modify the parasite, including several conditional approaches. This progress is finally allowing us not only to validate essential genes genetically, but also to study their molecular functions. In this review, I present our current understanding of the biological role proteases play in the malaria parasite life cycle. I also discuss how the recent advances in Plasmodium genetics, the improvement of protease-oriented chemical biology approaches, and the development of malaria-focused pharmacological assays, can be combined to achieve a robust biological, chemical and therapeutic validation of Plasmodium proteases as viable drug targets.

Keywords: malaria; protease; target validation.

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Figures

Figure 1
Figure 1
The malaria parasite life cycle. Schematic representation of the insect (A), liver (B) and blood (C) stages of parasite development. The timing of parasite development at each stage is indicated for Plasmodium falciparum. Note that gametocyte development is much faster in other Plasmodium spp., and that P. falciparum does not form hypnozoites.
Figure 2
Figure 2
Role of proteases during the erythrocytic cycle. (A) Role of proteases in parasite egress. (B) Role of proteases in RBC invasion. (C) Core biological functions of malaria proteases illustrated at trophozoite stage. Circles indicate zymogen/inactive protease forms, pacman shapes indicate active proteases. Asp, Cys, Ser, Thr and metalloproteases are shown in red, green, blue, orange and grey, respectively. Nucl, nucleus; Exo, exonemes; Mic, micronemes; Rhop, rhoptries; PV, parasitophorous vacuole; ER, endoplasmic reticulum; Mito, mitochondria; FV, food vacuole; Apic, apicoplast; FLC, falcilysin; CLP, Plasmodium calpain; hCLP1, human calpain‐1; and PAP, serine proline aminopeptidase.
Figure 3
Figure 3
Conditional complementation strategies in Plasmodium falciparum. Targeted protease genes are depicted in blue and complementing copies in green. Promoters are represented by block arrows and LoxP sites by triangles. (A) Domain swap strategy using the DiCre system. Introduction of ‘silent’ LoxP sites within artificial introns (orange rectangles) allows replacement of the catalytic domain with a mutant version upon rapamycin treatment. (B) Combination of the DiCre cKO with cKD strategies. In all examples, rapamycin‐induced conditional truncation of the catalytic domain is coupled with the up‐regulation of the complementing protease (wild‐type or mutant): (i) LoxP sites facing opposite directions can be used to activate the promoter of the complementing protease by adding rapamycin. (ii) Introduction of the glmS ribozyme (in red) between the mRNA stop codon and its 3’‐UTR allows for post‐transcriptional regulation of the target of interest. Glucosamine‐6‐phosphate (GlcN‐6P, blue hexagon) activates the ribozyme resulting in mRNA degradation and downregulation of the complementation copy. Removal of GlcN‐6P after rapamycin treatment will turn‐on the complementing copy after excision of the protease of interest. (iii) RNA aptamers (dark brown) designed to bind the Tet repressor (light brown shape) can be inserted upstream or downstream of the mRNA ORF to prevent translation. Addition of anhydrotetracycline (aTc: black circles) results in dissociation of the Tet repressor, thus allowing translation of the complementation copy. (iv) Fusion of a degradation domain (grey) to the complementing protein leads to its proteasomal degradation. However, addition of shield (blue square) allows folding and stabilisation of the degradation domain, thus preventing degradation of the complementing protease. (C) Example of how the ribozyme and TetR/RNA‐aptamer strategies can be combined to achieve conditional knock down of the target of interest and upregulation of its complementation copy.
Figure 4
Figure 4
Activity‐based probes as tools to study protease function. (A) ABPs are composed of: an electrophile (red triangle) that covalently modifies the catalytic nucleophilic residue of a protease; a recognition element (brown shape) that targets the probe towards specific proteases; and a tag, usually a fluorophore (pink circle), that allows for visualisation of labelled proteases. In the case of Asp or metalloproteases, which lack a nucleophilic residue, covalent interactions between the protease and the ABP can be obtained by using a photo‐crosslinker (represented in grey, FX‐ABPs). Quenched‐ABPs (Q‐ABPs) contain a quencher (black shape) within the leaving group of the electrophilic warhead that renders the probe nonfluorescent. Covalent modification of the protease results in the release of the quencher and an increase in fluorescent signal. Clickable ABPs are small molecule inhibitors containing a clickable handle (usually an alkyne group) that can be used to couple different tags to the probe after treatment of intact cells. (B) Broad‐spectrum ABPs use the conserved mechanism of an enzyme family to covalently modify all members of a protease family. These can be separated by SDS/PAGE and their labelling visualised as fluorescent bands. Specific inhibition of a protease (S INH) will result in the loss of signal for a single specific band (green arrow). Nonspecific inhibitors (NS INH) will block labelling of multiple bands (red arrows). (C) Quenched‐ABPs can be used to visualise protease activity in living cells since the probe only becomes fluorescent after binding to the protease of interest. This results in a localised increased of fluorescence signal within the subcellular compartment where the active protease resides. These probes can be used to measure real‐time target activation or inhibition. (D) Addition of an alkyne group to a lead inhibitor usually does not alter its biological activity. These clickable probes can be used to confirm target inhibition and identify potential off‐targets. After pretreatment of living cells with the clickable probe, cells are lysed, biotin linked to the alkyne group via click chemistry, and the targets of the compounds pulled down and identified by MS.
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