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

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

Activity-based protein profiling of human and plasmodium serine hydrolases and interrogation of potential antimalarial targets.

Malaria remains a global health issue requiring the identification of novel therapeutic targets to combat drug resistance. Metabolic serine hydrolases are druggable enzymes playing essential roles in lipid metabolism. However, very few have been investigated in malaria-causing parasites. Here, we used fluorophosphonate broad-spectrum activity-based probes and quantitative chemical proteomics to annotate and profile the activity of more than half of predicted serine hydrolases in P. falciparum across the erythrocytic cycle. Using conditional genetics, we demonstrate that the activities of four serine hydrolases, previously annotated as essential (or important) in genetic screens, are actually dispensable for parasite replication. Of importance, we also identified eight human serine hydrolases that are specifically activated at different developmental stages. Chemical inhibition of two of them blocks parasite replication. This strongly suggests that parasites co-opt the activity of host enzymes and that this opens a new drug development strategy against which the parasites are less likely to develop resistance

Development of a DPAP1-specific HTS assay.

<p><b>A.</b> Continuous assay. The assay was carried out in 384-well plates using 1% of parasite lysates. Substrate turnover was continuously measured for 5 min. JCP410 (10 µM) was used as a positive inhibition control. Z’ factor, S/N, and % CV of the negative control are shown. <b>B.</b> End-point assay for HTS. The reaction described in A was quenched after 10 min by addition of 0.5 M acetic acid. The final concentration of rhodamine product was quantified by fluorescence.</p

Use of an ABP to identify a DPAP1-selective substrate in parasite lysates.

<p><b>A.</b> Structure and reaction mechanism of the (Pro-Arg)₂-Rho substrate. <b>B.</b> Measurement of (Pro-Arg)₂-Rho apparent <i>K</i>_m in trophozoite lysates (circles) and with recombinant DPAP1 (triangle). Turnover rates at increasing concentrations of substrate were fitted to a Michaelis-Menten equation as described in the <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0011985#s4" target="_blank">methods</a> section. <b>C.</b> Labeling of DPAP1 activity in parasite lysates with FY01. Trophozoite lysates were incubated for 1 h with increasing concentrations of FY01. Labeling was stopped by boiling the sample in SDS-PAGE loading buffer. DPAP1 activity was measured using a flatbed fluorescent scanner. <b>D.</b> DPAP1 labeling correlates with substrate turnover inhibition. An aliquot of the samples treated for 1 h with FY01 was diluted in assay buffer containing 10 µM of (Pro-Arg)₂-Rho, and the initial turnover rate was measured in a 96-well plate (circles). This turnover rate is plotted with the labeling quantified in C.</p

Cat C-specific fluorogenic assay in rat liver lysates.

<p><b>A.</b> Labeling of Cat C with FY01. Rat liver extract extracts were treated with increasing concentrations of FY01 for 1 h and labeled proteins analyzed by SDS-PAGE followed by scanning of the gel using a flatbed laser scanner. The location of labeled Cat C is indicated. <b>B.</b> Inhibition of substrate turnover specifically correlates with Cat C labeling. The cleavage of (Pro-Arg)₂-Rho substrate was measured prior to analysis of FY01 labeling shown in part A. Quantification of the indicated labeled proteins relative to DMSO control is shown. <b>C.</b> Cat C-specific HTS assay in rat liver extracts. Rat liver lysates were treated for 30 min with either DMSO or JCP410 (10 µM) followed by the addition of 10 µM of (Pro-Arg)₂-Rho. The turnover rate was continuously measured for 5 min in a 384-well plate. Z’ factor, S/N, and % CV of the negative control are shown.</p

A Coupled Protein and Probe Engineering Approach for Selective Inhibition and Activity-Based Probe Labeling of the Caspases

Caspases are cysteine proteases that play essential roles in apoptosis and inflammation. Unfortunately, their highly conserved active sites and overlapping substrate specificities make it difficult to use inhibitors or activity-based probes to study the function, activation, localization, and regulation of individual members of this family. Here we describe a strategy to engineer a caspase to contain a latent nucleophile that can be targeted by a probe containing a suitably placed electrophile, thereby allowing specific, irreversible inhibition and labeling of only the engineered protease. To accomplish this, we have identified a non-conserved residue on the small subunit of all caspases that is near the substrate-binding pocket and that can be mutated to a non-catalytic cysteine residue. We demonstrate that an active-site probe containing an irreversible binding acrylamide electrophile can specifically target this cysteine residue. Here we validate the approach using the apoptotic mediator, caspase-8, and the inflammasome effector, caspase-1. We show that the engineered enzymes are functionally identical to the wild-type enzymes and that the approach allows specific inhibition and direct imaging of the engineered targets in cells. Therefore, this method can be used to image localization and activation as well as the functional contributions of individual caspase proteases to the process of cell death or inflammation

Formation of plaques in static <i>P</i>. <i>falciparum</i> asexual blood stage microplate cultures.

(A) Time-dependent evolution of plaques. Scanned images (RGB) of a single microplate well (diameter 6.38 mm) taken at the indicated times following the introduction of a low parasitaemia P. falciparum 3D7 culture (200 μl per well, 0.75% haematocrit, starting parasitaemia ~0.00004%, corresponding to ~6 infected cells per well). Plaques first became detectable by light microscopy or high resolution scanning at day 8. (B) Plaque density correlates with starting parasitaemia. Shown are microplate wells containing 10-fold serial dilutions (indicated) of a P. falciparum culture (0.75% haematocrit, starting parasitaemia 13%). The wells were imaged on day 13 following initiation of the culture. By this point, at the highest parasite densities (neat and 10−1 dilution) the erythrocyte layer was completely destroyed, whilst discrete plaques were visible at the lower parasite densities. Total numbers of plaques in each set of 6 replicate wells of the 10−3–10−8 dilutions (replicate wells are not shown) was 2,034 (10−3), 245 (10−4), 26 (10−5), 4 (10−6), and 0 plaques (10−7 and 10−8). No plaques were ever detected in the complete absence of parasites (none). (C) Linear regression by analysis of covariance (ANCOVA) indicating a strong linear inverse correlation between dilution and plaque density in the wells containing the 10−3–10−6 parasite dilutions. A plot of observed mean plaque frequency against dilution (blue) from the experiment shown in (B) is shown alongside a plot of the plaque frequencies expected if there is a linear inverse correlation between plaque number and dilution (red). Values of the statistical data (R2, F statistic, number of degrees of freedom and p value) are shown. (D) Optimisation of haematocrit conditions. Microplate wells containing cultures at ~0.00004% parasitaemia at the indicated haematocrits, imaged on day 15. Whilst plaque formation was easily detected in the 0.75% haematocrit wells, they were much more difficult to detect at higher haematocrit, whilst at lower haematocrit values the plaques were typically more diffuse with signs of erythrocyte lysis. Inset, zoomed region of the 0.75% haematocrit erythrocyte layer, showing the discrete nature of the plaques.</p

Role of DPAP3 in parasite egress.

(A) L-WSAK is a more selective DPAP3 inhibitor than SAK1. The structures of SAK1, L-WSAK and D-WSAK are shown. Merozoite or schizont lysates were pre-incubated with a dose response of inhibitor for 30 min followed by FY01 labelling. Samples were run on an SDS-PAGE gel, and the gel scanned on a flatbed fluorescence scanner. Bands corresponding to each of the labelled cysteine proteases are indicated by arrows. (B) Effect of inhibitors on egress. DMSO or RAP treated A1cKO parasites were treated at schizont stage with a dose response of inhibitor for 24 h. The accumulation of schizonts upon inhibitor treatment was quantified by FACS. (C) Analysis of egress by video microscopy. C2-arrested schizonts obtained from DMSO- or RAP-treated of A1cKO, F8cKO, F3cKO or E7ctr parasite lines were monitored by time-lapse DIC microscopy for 30 min after C2 washout. Representative still images taken at 0, 15, and 30 min are shown for F8cKO parasites. The full time-lapse video can be seen in S4 Video. The percentage of schizonts that egressed during this 30 min time-lapse (left graph) and the time at which each individual schizont ruptured (right graph) are shown. Bar graphs show mean values ± standard deviation; circles show individual biological replicates (filled for F8cKO, empty for A1cKO, and grey for F3cKO). (D) WB analysis of culture supernatant collected after egress of 3D7 and A1cKO after DMSO or RAP treatment. No differences in the processing of AMA1, MSP1 or SERA5 was observed as a result of DPAP3 truncation.</p

DPAP3 has proteolytic activity.

<p>(<b>A</b>) Analysis of purified rDPAP3. Two main bands are detected by silver stain, both of which are strongly labelled by FY01 and recognized by the anti-Nt-DPAP3 and anti-Ct-DPAP3 antibodies. All other minor bands in the silver stain are also recognized by DPAP3 antibodies and represent degradation products that could not be separated during purification. (<b>B</b>) Measurement of VR-ACC turnover and FY01 labelling for WT and C504S MUT rDPAP3. Silver stain analysis shows equivalent amounts of protein were obtained from the purification of WT and MUT rDPAP3. (<b>C</b>) pH dependence of rDPAP3 activity measured at 10 μM VR-ACC (n = 3).</p

Generation of DPAP3 cKO lines.

<p>(<b>A</b>) Schematic representation and assessment of the <i>dpap3</i> recombinant genetic locus before and after RAP-mediated excision for the F8cKO and A1cKO lines. Wild type exons (EX) and intron (IN) sequences are depicted with grey and pink boxes, respectively. The homology regions used for single crossover recombination are indicated with blue lines, and the recodonized 3’ end of the second exon is shown in green (scEX2). <i>loxP</i> sites (yellow arrows) were introduced downstream of the <i>P</i>. <i>berghei</i> 3’UTR (white circle) and either within the ORF of scEX2 (A1cKO) or upstream of scEX2 (F8cKO) as a <i>loxPint</i> artificial intron (pink striped box). The <i>mCherry</i> coding sequence (red box), the <i>hdhfr</i> resistance cassette (black box), and the displaced endogenous <i>dpap3</i> locus with its 3’UTR (black circle) are also shown. Arrows indicate primers annealing sites used for diagnostic PCR of excised (purple) and non-excised (green) loci. (<b>B</b>) Diagnostic PCR showing excision at the <i>dpap3</i>-locus. PCR was performed on genomic DNA collected from the E7ctr, A1cKO and F8cKO lines 24 h after DMSO or RAP treatment. Genomic DNA from the parental 1G5 line was used as a negative control. Excision and non-excision PCR products are indicated with purple and green arrows, respectively. Excision product was only observed after RAP treatment of the cKO lines. The presence of a non-excised PCR product after RAP treatment indicate that excision is not 100% efficient. (<b>C</b>) WB analysis showing highly efficient loss of DPAP3 upon RAP treatment. Schizonts collected 45 h after DMSO or RAP treatment of E7ctr and F8cKO parasites were saponin lysed, and the parasite pellet analyzed by WB using an anti-mCherry antibody (red arrow). SUB1 was used as a loading control (blue arrow). (<b>D</b>) IFA analysis of mature schizonts showing the loss of DPAP3 signal after RAP treatment. Ring-stage F8cKO parasites were treated with DMSO or RAP for 3 h and fixed for IFA analysis at 48 h.p.i. Slides were stained with anti-mCherry (green) and anti-MSP1 (red) or anti-SUB1 (red) antibodies. DNA was stained with DAPI (blue); scale bar: 5 μm.</p