Does a specific blood group enhance the Plasmodium growth?

Does a specific blood group enhance the Plasmodium growth?

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I am maintaining Plasmodium falciparum cultures for past 6 months. For the blood culture, usually we lab members take turns and donate blood for the culture. I observed that the parasite's normal growth cycle were fast and more healthy in B+ blood, little slow in AB+ blood. But on the whole the culture was maintained good but I really observed fast stage transitions in B+.

Is this really possible? Does a specific blood group have an impact on Plasmodium growth? Did anyone notice the same issue?


It is a well documented observation that Plasmodium (vivax and knowlesi) infection is dependent on the Duffy blood groups [1].

Individuals lacking the Duffy antigens (Fya and Fyb) have lower susceptibility to malaria. Plasmodium expressed Duffy Binding Proteins facilitate in establishing the initial contact between the merozoite and the RBCs.

However, Plasmodium has evolved to break this dependence on Duffy antigens [2, 3].

For, the ABO blood group system, it has been observed that the blood group-O is associated with lower severity of P.falciparum malaria in adults [4, 5], which possibly happens because of reduced rosetting (binding of infected RBCs with uninfected RBCs) [5].


  1. Dean L. Blood Groups and Red Cell Antigens [Internet]. Bethesda (MD): National Center for Biotechnology Information (US); 2005. Chapter 9, The Duffy blood group. Available from:

  2. Ménard, Didier, et al. "Plasmodium vivax clinical malaria is commonly observed in Duffy-negative Malagasy people." Proc Nat Acad Sci, USA 107.13 (2010): 5967-5971.

  3. Mendes, Cristina, et al. "Duffy negative antigen is no longer a barrier to Plasmodium vivax-molecular evidences from the African West Coast (Angola and Equatorial Guinea)." PLoS Negl Trop Dis 5.6 (2011): e1192.

  4. Cserti, Christine M., and Walter H. Dzik. "The ABO blood group system and Plasmodium falciparum malaria." Blood 110.7 (2007): 2250-2258.

  5. Rowe, J. Alexandra, et al. "Blood group O protects against severe Plasmodium falciparum malaria through the mechanism of reduced rosetting." Proc Nat Acad Sci, USA 104.44 (2007): 17471-17476.

Plasmodium vinckei genomes provide insights into the pan-genome and evolution of rodent malaria parasites

Rodent malaria parasites (RMPs) serve as tractable tools to study malaria parasite biology and host-parasite-vector interactions. Among the four RMPs originally collected from wild thicket rats in sub-Saharan Central Africa and adapted to laboratory mice, Plasmodium vinckei is the most geographically widespread with isolates collected from five separate locations. However, there is a lack of extensive phenotype and genotype data associated with this species, thus hindering its use in experimental studies.


We have generated a comprehensive genetic resource for P. vinckei comprising of five reference-quality genomes, one for each of its subspecies, blood-stage RNA sequencing data for five P. vinckei isolates, and genotypes and growth phenotypes for ten isolates. Additionally, we sequenced seven isolates of the RMP species Plasmodium chabaudi and Plasmodium yoelii, thus extending genotypic information for four additional subspecies enabling a re-evaluation of the genotypic diversity and evolutionary history of RMPs.

The five subspecies of P. vinckei have diverged widely from their common ancestor and have undergone large-scale genome rearrangements. Comparing P. vinckei genotypes reveals region-specific selection pressures particularly on genes involved in mosquito transmission. Using phylogenetic analyses, we show that RMP multigene families have evolved differently across the vinckei and berghei groups of RMPs and that family-specific expansions in P. chabaudi and P. vinckei occurred in the common vinckei group ancestor prior to speciation. The erythrocyte membrane antigen 1 and fam-c families in particular show considerable expansions among the lowland forest-dwelling P. vinckei parasites. The subspecies from the highland forests of Katanga, P. v. vinckei, has a uniquely smaller genome, a reduced multigene family repertoire and is also amenable to transfection making it an ideal parasite for reverse genetics. We also show that P. vinckei parasites are amenable to genetic crosses.


Plasmodium vinckei isolates display a large degree of phenotypic and genotypic diversity and could serve as a resource to study parasite virulence and immunogenicity. Inclusion of P. vinckei genomes provide new insights into the evolution of RMPs and their multigene families. Amenability to genetic crossing and transfection make them also suitable for classical and functional genetics to study Plasmodium biology.

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Azithromycin inhibits merozoite invasion

Application of the merozoite purification method of Boyle et al. [18] identified the macrolide antibiotic azithromycin as a candidate inhibitor of P. falciparum asexual blood-stage merozoite invasion of the host erythrocyte (see Fig. 1a for representation of assay setup, Fig. 2 for structure of azithromycin and other drugs used in this study). Initial screens indicated that the invasion inhibitory IC50 differed between azithromycin prepared in ethanol (10 μM) or DMSO (38 μM), suggesting that choice of vehicle can impact azithromycin potency in vitro. Comparative screens of azithromycin potency over different treatment times demonstrated that the IC50 of merozoite invasion assays (1 hour exposure to drug) were surprisingly similar to the in cycle (40 hour incubation, no invasion step, Fig. 1b) and 1 cycle (90 hour, one invasion step, Fig. 1c) assays (IC50 –(drug prepared in ethanol) invasion 10 μm in cycle 6 μM 1 cycle 7 μM, −(drug prepared in DMSO) invasion 38 μM (DMSO) in cycle 12 μM 1 cycle assay 7 μM, Fig. 3a, Table 1). These results suggest that azithromycin can act independently against both merozoite invasion and intracellular parasite growth within one cycle of treatment in vitro.

Drug treatment strategies used in this study. The lifecycle stage of drug treatment is represented in the first box for each panel and the stage of parasitaemia measurements are highlighted by red boxes with yellow background. (a) Merozoite successful invasion of drug-treated purified merozoites was measured at ring-stage immediately after addition of erythrocytes and merozoite invasion (<1 hour rings) or after washing out the drug (denoted by green dashed line) and growing parasites through to late trophozoite stage (40 hours post-invasion). (b) In cycle early ring-stage parasites (<4 hours post-invasion) were drug-treated and the resulting growth inhibition was assessed at late trophozoite stage (40 hours post-invasion). (c) 1 cycle early ring-stage parasites (<4 hours post-invasion) were drug-treated and the resulting growth inhibition was assessed 90 hours later, after 1 cycle of reinvasion, at late trophozoite stage. These assays are a longer duration, but nonetheless equivalent in terms of including 1 cycle of reinvasion and development, to 1 cycle assays reported in other studies [41–44]. (d) 2 cycle (delayed death) early ring-stage parasites (<4 hours post-invasion) were drug-treated and the drug-treated parasites were grown for 80 hours prior to washing out the drug in fresh media (denoted by green dashed line). Growth inhibition was assessed approximately 40 hours later after a second cycle of reinvasion (120 hours post-invasion). (e) Live filming mature schizont-stage parasites were incubated with azithromycin and the success of merozoite invasion was recorded and quantified by live filming (green arrow, successful invasion blue arrow, deformation of the erythrocyte but no invasion red arrow, attachment and release or attachment with failure to deform erythrocyte and no release) (see Additional file 1: Video S1 and Additional file 2: Video S2). White box-drug treatment yellow box-analysis of parasitaemia green dashed line-drug washout

Structure of macrolide antibiotics. (a) Structure of the 15-membered macrolide, azithromycin, and its modified analogues (names used in the text underlined, in brackets). (b) Structure of the 14-membered macrolide, erythromycin A, and its modified analogues. (c) Structure of the 16-membered macrolide, spiramycin. (d) Structure of the non-macrolide antibiotic, clindamycin

Azithromycin inhibits P. falciparum merozoite invasion. (a) The potency of azithromycin in ethanol or DMSO as vehicle was compared for invasion inhibition (unbroken line, 10 minute merozoite treatment, parasitaemia measured 40 hours later) and 1 cycle growth inhibition assays (broken line, treatment rings to trophozoites’ next cycle). The invasion inhibitory IC50 of azithromycin prepared in ethanol (blue, IC50 10 μM) was similar to that for growth inhibition assays (IC50 7 μM P = 0.0743, Log IC50 same between data sets, extra sum of squares F-test). The invasion inhibitory activity of azithromycin in DMSO (red, IC50 38 μM) was 5-fold higher than 1 cycle growth assays (IC50 7 μM P <0.0001, Log IC50 different between data sets). (b) Inhibition profiles for pretreated erythrocytes (RBC Pre), merozoite treatment (T = 0 drug added at time zero) and rings treated for <1 hour (T = 20 drug added 20 minutes post-invasion) were identical between azithromycin (in DMSO) and the invasion inhibitor heparin (IC80 concentration). (c) Increasing the concentration of azithromycin (in ethanol) to 10 × IC80 (380 μM) did not result in substantial inhibition of invasion into pretreated cells compared to treatment of merozoites. (d) Flow cytometry and microscopy assessments confirmed that azithromycin (IC80 in ethanol) and heparin, but not the trophozoite-targeting antimalarial halofantrine (2 × IC80 ring-stage treatment (46 nM) [13]), inhibit merozoite invasion and establishment of ring stages in erythrocytes. Representative (e) flow cytometry plots (GFP high and EtBr low ring-stage parasites represented by square gate) and (f) microscopy thin smears (rings highlighted by green arrows) show absence of ring-stage parasites for azithromycin and heparin compared to non-invasion inhibitory controls (labeling as per Fig. 3d). Experiments represent the mean and SEM of three or more experiments. Significance was tested using an unpaired t-test (*P = 0.01–0.05, **P ≤0.01, ***P ≤0.001)

To confirm that azithromycin was inhibiting merozoite invasion directly in our assays and was not acting downstream of invasion during intracellular parasite growth, the effect of exposure of merozoites or early ring-stage parasites to the drug was examined azithromycin was washed out of the cultures within 1 hour and parasitaemia assessed by flow cytometry 40 hours post-assay setup for all treatments (prior to the next round of parasite rupture and invasion of new host cells). Treatment of merozoites (10 minute incubation) and early ring stages for less than 1 hour (T = 0) with azithromycin (1 × IC80 mero DMSO 151 μM) and the invasion inhibitory control compound, heparin, was significantly more inhibitory to parasite growth than the same treatment of early ring stages at the time-point of 20 minutes post-invasion (T = 20 (exposed for 40 minutes) azithromycin P <0.001 heparin P <0.01). Additionally, pretreatment of erythrocytes, followed by washout of drug, had little or no inhibitory effect on invasion (Fig. 3b azithromycin P <0.001 heparin P <0.001). This indicated that exposure of merozoites, but not very early ring stages or uninfected erythrocytes, was inhibitory to parasite growth.

In order to confirm that the inhibitory activity was against purified merozoites and not the uninfected erythrocyte prior to merozoite invasion, erythrocytes were pretreated with azithromycin at a 10 × IC80 mero (380 μM (drug prepared in ethanol to limit nonspecific effects of vehicle)) prior to washing and addition of merozoites. There was minimal loss of invasion into erythrocytes pretreated with a 10 × IC80 mero concentration of azithromycin for 1 hour relative to control (Fig. 3c). In contrast, merozoites treated for 10 minutes with the same concentration of azithromycin were unable to establish erythrocyte infections (P <0.001), supporting the conclusion that it is the merozoite and its interaction with the erythrocyte, and not the erythrocyte itself, that is the target of inhibition.

Next we confirmed that azithromycin was inhibiting merozoite invasion directly, rather than acting on growth downstream of invasion, through the complementary methods of microscopy and flow cytometric quantification of ring-stage parasites at the time-point of 1 hour post-invasion. There was an almost complete loss of ring-stage parasites seen by both microscopy and flow cytometry for azithromycin (1 × IC80 mero ), confirming that azithromycin is indeed inhibitory to merozoite invasion (Fig. 3d,e,f). The merozoite invasion inhibitory activity of azithromycin was unaffected by limiting drug exposure to <5 seconds prior to erythrocyte addition, by the presence of serum in the culture medium or the presence of haemazoin crystals (that can remain after purification of merozoites) in the invasion assay. These results demonstrate that azithromycin is a rapid inhibitor of merozoite invasion in vitro.

Azithromycin inhibits merozoite invasion prior to tight junction formation

Live cell imaging of invading Plasmodium merozoites indicates that erythrocyte invasion is a multi-step process that typically takes less than 30 seconds to complete, after contact occurs between the merozoite and the erythrocyte, in primate species and slightly longer in rodent species [35–37]. After contacting its target erythrocyte, the merozoite reorients its apical end onto the erythrocyte surface in a process that deforms the erythrocyte. This reorientation lasts about 10 seconds and is followed by the formation of a ring-like region of close and irreversible contact, called the tight (or moving) junction, between the merozoite apex and the erythrocyte. The merozoite enters the host cell through this junction using the power of its actin-myosin motor. After invasion, the erythrocyte surface reseals and the merozoite differentiates into a ring-stage parasite over several minutes [35]. We used live cell imaging to identify invasion step(s) that are inhibited by azithromycin (Fig. 1e). Late schizont-stage parasites were allowed to rupture in the presence of 75 and 134 μM azithromycin (in ethanol), which represent 2 × IC80 and 3.5 × IC80 of merozoite invasion inhibition concentrations, respectively. These slightly higher drug concentrations were used to clearly define the inhibitory phenotype. These concentrations were not found to inhibit schizont rupture. Five schizont ruptures were observed for each treatment. In the untreated control, 22 merozoites were observed to contact erythrocytes. After a contact period ranging from a few seconds to 2 minutes, 18 % of the merozoites released or detached. A further 32 % of the merozoites progressed to the erythrocyte deformation stage before releasing over a few minutes, and the remaining 50 % advanced to complete invasion (Fig. 4). Azithromycin treatment at 75 μM changed this profile with fewer merozoites invading (32 %) and a greater proportion failing to progress beyond the initial contact and deformation stages (68 %, Fig. 4). At 134 μM there was a significant and dramatic change in the invasion profile compared to the untreated control and azithromycin treatment at 75 μM with no merozoites invading and most releasing after initial contact (81 %, Fig. 4). The observation that merozoites failed to make sustained contact with erythrocytes in the presence of azithromycin indicates that azithromycin acts early in invasion to prevent tight junction formation [23, 35, 38]. The periods of time that each of the invasion steps took to occur was also measured, but in nearly all cases showed no significant difference between the treatments. Representative video files showing the effects of azithromycin are shown in Additional file 1: Video S1 and Additional file 2: Video S2.

Azithromycin inhibits the early steps of invasion. Video microscopy of merozoite invasion of erythrocytes was performed in the presence of 75 and 134 μM azithromycin (in ethanol) compared to a no drug control (0 μM). Five schizont ruptures were observed for each treatment. Of the merozoites that contacted erythrocytes, some were observed to deform erythrocytes and then successfully invade their host cells (contact–invade), while others did not progress beyond initial attachment (contact–detach) or progressed to deformation but did not invade (contact–deform). From several rupturing schizonts, the number of merozoites exhibiting each of these steps was counted for each drug treatment and the percentages are shown along with the number of events in the column boxes. A Chi-squared test was performed to indicate significant differences at the following levels (**P ≤0.01, ***P ≤0.001)

Macrolides related to azithromycin also inhibit merozoite invasion

After identifying that azithromycin was inhibitory to merozoite invasion, we examined whether related drugs with a history of clinical use as antibiotics also had this property. Macrolides with a 14-membered macrolactone ring (erythromycin A, roxithromycin, dirithromycin) and 16-membered ring (spiramycin) were tested using the same methods as azithromycin (prepared in ethanol, Fig. 2b,c). Determination of the inhibitory concentration (Fig. 5a,b,c,d Table 1) indicated that azithromycin (ethanol, 10 μM) had >8-fold lower IC50 for merozoite invasion when compared to roxithromycin (83 μM) and spiramycin (123 μM), while erythromycin (420 μM) was 42-fold less potent than azithromycin. Of interest was the very poor invasion inhibitory activity of dirithromycin (521 μM), which was 52-fold less potent than azithromycin, even though the IC50 values determined using 1 cycle growth assays were very similar (dirithromycin, IC50 90hr 8 μM azithromycin, IC50 90hr 7 μM Table 1). Invasion inhibition was confirmed by measurement of ring-stage parasites 1 hour after invasion at 1 × IC80 mero for erythromycin A, roxithromycin and spiramycin. The lack of inhibitory activity when erythrocytes were pretreated with drug (and then washed) further confirmed specific inhibition of merozoite invasion (Fig. 5e P <0.01).

Related macrolides inhibit merozoite invasion. The 14-membered macrolides (a) erythromycin A (IC50 mero 420 μM), (b) roxithromycin (83 μM), (c) dirithromycin (521 μM) and (d) the 16-membered macrolide spiramycin (123 μM) had variable levels of invasion inhibitory activity (green) and a higher IC50 than that achieved for 1 cycle assays (red). (e) The invasion inhibitory activity of erythromycin A, roxithromycin and spiramycin at an IC80 concentration was confirmed by flow cytometry assessment of ring stages with minimal inhibition evident for pretreated erythrocytes. All experiments represent the mean and SEM of three or more experiments. Significance of differences was compared using an unpaired t-test (**P ≤0.01, ***P ≤0.001)

Merozoite invasion inhibition is independent of macrolide activity against apicoplast ribosomes

Since macrolide antibiotics are known to target the 50S ribosomal subunit of the apicoplast to inhibit subsequent intra-erythrocytic development, we tested the invasion inhibitory activity of clindamycin, which has a similar mechanism of action and exhibits overlapping binding of the 50S ribosomal subunit of the apicoplast to azithromycin [39] (Table 1 Fig. 2c). Clindamycin had very weak invasion inhibitory activity there was evidence for invasion inhibitory activity with a 74-fold higher IC50 observed for clindamycin (IC50 mero 743 μM) than seen for azithromycin (10 μM). However, pre-treatment of erythrocytes with 2,972 μM of clindamycin (IC80 mero ) resulted in a 65 % reduction in merozoite invasion relative to untreated controls and there was evidence of erythrocyte lysis using these high levels of clindamycin (Fig. 6a). This suggests that at such high concentrations clindamycin is having a largely non-specific effect on merozoite invasion through damaging erythrocytes.

The mechanism of invasion inhibition is unlikely to target the apicoplast ribosome. (a) Clindamycin targets the same subunit of the apicoplast ribosome but was found to have a much higher IC80 for apparent merozoite invasion inhibition (2,972 μM). There was evidence of non-specific inhibition of invasion as pretreatment of erythrocytes with clindamycin gave significant inhibition, which was not seen for azithromycin (prepared in ethanol mean and SEM of four or more experiments significance of differences tested with an unpaired t-test **P ≤0.01) (38 μM). (b) The D10-AZR r line showed up to a 57-fold higher tolerance of azithromycin in 2 cycle (delayed death) apicoplast-targeting drug inhibition assays compared to D10 parental line. In contrast, the IC50 for purified merozoite invasion inhibitory activity differed by less than 2.5-fold between the D10-AZR r line and the D10-PfPHG line for (c) azithromycin (IC50: PfPHG 10 μM D10-AZR r 25 μM), (d) erythromycin A (IC50: PfPHG 420 μM D10-AZR r 732 μM) and (e) clindamycin (IC50: PfPHG 743 μM D10-AZR r 557 μM). Data represent the mean of two or more experiments in at least duplicate. D10-AZR r , D10 azithromycin-resistant

We tested further whether apicoplast ribosomal protein translation was the target of the inhibitory activity at the merozoite stage by comparison of the invasion inhibitory activity of azithromycin, erythromycin and clindamycin between the D10-PfPHG (azithromycin-sensitive) parasite used throughout this study and an azithromycin-resistant D10 derivative (D10-AZR r ) selected for reduced sensitivity to both azithromycin and erythromycin in 2 cycle (delayed death) assays. Sequencing of apicoplast ribosome genes revealed a G91D mutation in the rpl4 gene product, corresponding to the G112D mutation in Chlamydomonas reinhardtii, known to confer resistance to erythromycin [40]. This mutation is associated with a 57-fold loss of sensitivity to azithromycin over 2 cycles of parasite growth (IC50 120hr D10-AZR r , 9 μM D10 parental, 0.16 μM P <0.0001 Fig. 6b Table 1). Comparison of the IC50 mero (Fig. 6c,d,e) suggested that there was very little difference in merozoite invasion inhibitory activity between the resistant versus sensitive lines for azithromycin (in ethanol, 2.5-fold difference IC50 D10-AZR r , 25 μM D10-PfPHG, 10 μM P <0.0001), erythromycin (1.6-fold difference IC50 D10-AZR r , 732 μM D10-PfPHG, 442 μM P = 0.0247) or clindamycin (1.3-fold difference IC50 D10-AZR r , 556 μM D10-PfPHG, 736 μM P = 0.26). Together, these data suggest that the invasion inhibitory activity of azithromycin is largely independent of apicoplast ribosomal protein synthesis, consistent with the current view that the apicoplast does not play a role in invasion.

Modification of macrolides enhances invasion inhibitory activity

We tested a panel of macrolide analogues to determine whether the IC50 of merozoite invasion inhibition could be reduced. An erythromycin A L-megosamine sugar derivative (Meg-erythromycin, 6-O-megosaminyl erythromycin A) that has a lower IC50 in 1 cycle growth inhibition assays [41] was tested and found to reduce the IC50 of merozoite invasion inhibition (13 μM) 32-fold compared to the parent compound (420 μM Fig. 7a Table 1 Fig. 2b). Furthermore, addition of an oxime group (N-OH) to erythromycin A (erythromycin oxime) lowered the invasion inhibitory IC50 (150 μM) almost 3-fold compared to erythromycin A. The increased potency of Meg-erythromycin and erythromycin oxime compared to the parent drug indicates that various modifications to macrolides can lead to more potent invasion inhibitory activity.

Macrolide modification lowers invasion inhibitory IC50, but not apicoplast-targeting ‘delayed death’ inhibition. (a) Addition of an L-megosamine sugar [41] to form Meg-erythromycin (6-O-megosaminyl erythromycin A, IC50 13 μM) or an oxime group (150 μM) lowered the invasion inhibitory IC50 activity compared to the parent drug erythromycin A (IC50 420 μM). (b) Screening of an azithromycin analogue panel identified three compounds (12e, 15 μM 1j, 7 μM 11c, 28 μM) with up to 5-fold lower invasion inhibitory IC50 compared to the parent azithromycin (in DMSO, 38 μM). (c) Treatment of parasites during in cycle (40 hours, rings to schizonts), 1 cycle (90 hours, 1 cycle of replication) and 2 cycle (120 hours, 2 cycles of replication) assays with azithromycin (in DMSO) and analogues (12e, 1j), indicated that the IC50 of 1 cycle (40 hour and 90 hour, high drug concentration) inhibition was greatly reduced for the analogues compared to azithromycin. In contrast, the IC50 of the delayed death phenotype was almost identical for azithromycin and its analogues. d) Video microscopy of merozoite invasion was performed in the presence of a no drug control (0 μM), 122 μM of analogue 12e (2 × IC80) and azithromycin (AZR both in DMSO). Merozoites that contacted the erythrocyte and i) invaded (cont–invade), ii) deformed but did not invade (cont–deform), or iii) released without deforming or invading (cont–detach) were tallied and analyzed as per Fig. 4. (e) Removal of the cladinosyl sugar (azithromycin descladinosyl, 50 μM) from azithromycin increased the invasion inhibitory IC50 compared to azithromycin (10 μM). The additional removal of the desosaminyl sugar (azithromycin desglycan, IC50 >1,600 μM) resulted in loss of invasion inhibitory activity compared to azithromycin. Data represent the mean and SEM of three or more experiments (**P ≤0.01, ***P ≤0.001)

Next we obtained a small panel of azithromycin analogues (in DMSO) from GlaxoSmithKline (Tres Cantos, Spain) [42–44] with reported IC50 values lower than azithromycin in a 72 hour growth inhibition assay (roughly equivalent to 1 cycle assays in this study Fig. 1c). In our studies, we found that azithromycin dissolved in DMSO was less effective so the following comparisons use the IC50 values for both azithromycin and the azithromycin analogues with DMSO as the solubilization vehicle. Two compounds (12e, 15 μM 1j, 7 μM) had a substantially lower IC50 for merozoite invasion inhibition than azithromycin (38 μM (when prepared in DMSO) Fig. 7b Table 1 Fig. 2a), indicating that azithromycin can also be modified to lower its invasion inhibitory activity in vitro.

When early ring-stage parasites were treated with the compounds 1j and 12e for 40 hours (in cycle 1j, 0.9 μM 12e, 1 μM) and 90 hours (1 cycle 1j, 0.7 μM 12e, 0.6 μM) the IC50 of the modified analogues was between 10- and 14-fold lower than that of azithromycin (DMSO in cycle, 12 μM 1 cycle, 7 μM Fig. 7c Table 1). In contrast, parasites treated for 120 hours (2 cycle delayed death, Fig. 1d) showed a small increase in the potency of azithromycin (DMSO, 0.1 μM) over 1j (0.16 μM) and 12e (0.16 μM) for the ‘delayed death’ inhibition typical of apicoplast ribosome targeting (Table 1). This would suggest that the apicoplast ribosome targeting is not affected by the modifications of 1j and 12e, and is a further indication that invasion inhibition or parasite growth inhibition over 1 cycle of parasite growth or less is not a result of inhibition of apicoplast ribosome activity. Further, the ability of azithromycin and its analogues to inhibit intracellular parasite growth during in cycle assays (from early rings to late trophozoites) at IC50 values lower than the invasion inhibition assays suggests that the compounds can inhibit parasite growth independent of merozoite invasion.

In order to assess whether the invasion inhibitory phenotype of a more potent analogue was similar to our observations for azithromycin in ethanol (see Fig. 4b), live filming of merozoite invasion in the presence of analogue 12e was examined using live video microscopy. As was observed in Fig. 4, for the no drug treatment control 53 % of merozoites that contacted an erythrocyte invaded, 29 % deformed but failed to invade the erythrocyte after contacting and 18 % released the erythrocyte membrane without invading or deforming the erythrocyte (Fig. 7d). In contrast, addition of 12e at 122 μM (2 × IC80) reduced invasion to only 8 % of total contact events, reduced contacts leading to erythrocyte deformation to 27 % of contact events and increased the number of merozoites releasing without invading or deforming the erythrocyte to 65 % of all contact events, resulting in failure to invade for 92 % of all contact events. These results support the live filming observations for azithromycin in ethanol (Fig. 4b) and again indicate that azithromycin acts early in the invasion process and inhibits tight junction formation. We compared the invasion phenotype of 12e to that of azithromycin in DMSO at an equal concentration (approximately 1 × IC80 of azithromycin in DMSO). As expected, at the same concentration the less potent parent azithromycin was less inhibitory to invasion (25 % contact events invaded) and as a result had fewer failed invasion events (75 %). Together, these findings establish a proof-of-concept that macrolides with more potent invasion inhibitory activity can be developed through chemical modification and that the dual activities of invasion inhibition and other inhibitory activities can be developed in single compounds and are not mutually exclusive.

Identification of structural groups on azithromycin important for invasion inhibitory activity

We sought to identify structural groups of azithromycin that were most important for invasion inhibitory activity by determining the effect of removing one or both of the glycosylated groups. The two glycan groups of azithromycin, in particular the desosamine group, have been identified as critical to proper drug binding and inhibition of bacterial ribosome translation [39]. Removal of the cladinosyl group (descladinosyl) for both azithromycin and erythromycin A led to an increase of the drugs IC50 for both in cycle (azithromycin, 6 μM, AZR-descladinosyl, 39 μM erythromycin A, 230 μM, ERY-descladinosyl, 288 μM) and 1 cycle (azithromycin, 7 μM AZR-descladinosyl, 31 μM erythromycin A, 52 μM ERY-descladinosyl, 185 μM) drug assays (Table 1 Fig. 2a,b). Similarly, removal of the cladinosyl group from azithromycin (AZR-descladinosyl, 50 μM) led to 5-fold reduction in the invasion inhibitory activity of the compound compared to azithromycin (azithromycin, 10 μM, Fig. 7e).

We were further able to remove both the cladinosyl and desosamine groups from azithromycin (AZR-desglycan Fig. 7e Table 1 Fig. 2a), which led to a dramatic loss of invasion inhibitory activity and an increase in the IC50 beyond the limits of the assay (>1,600 μM). Comparison of these compounds indicates that the cladinosyl group has a role in lowering the IC50 of invasion inhibition for azithromycin, but the presence of both the cladinosyl and desosamine groups is critical for azithromycin’s invasion inhibitory activity. While these data suggest that the desosamine group plays the critical role during invasion inhibition, technical limitations prevented the selective removal of the desosamine group.

Azithromycin inhibits P. berghei merozoite and Toxoplasma gondii tachyzoite invasion, but not P. berghei sporozoite invasion

To test whether azithromycin could inhibit invasion of P. berghei, widely used as a murine model of malaria, we purified merozoites [45] and allowed invasion to proceed in the presence of azithromycin. Azithromycin inhibited merozoite invasion of P. berghei at very similar concentrations to that seen for P. falciparum, with no evidence of inhibition for parallel treatments of newly invaded rings (Fig. 8a,b).

Azithromycin inhibits P. berghei merozoite and Toxoplasma gondii tachyzoite, but not sporozoite, invasion of host cells. (a) Azithromycin inhibited purified P. berghei merozoite invasion at similar concentrations compared to P. falciparum. (b) Treatment of P. berghei-infected erythrocytes immediately after invasion with azithromycin for 30 minutes did not result in a loss of late-stage parasites detected by flow cytometry, confirming that azithromycin inhibited invasion and not parasite growth. (c) Azithromycin was found to have a small but significant effect on the number of P. berghei sporozoites that had entered or traversed host cells. However, using a more specific assay that measures successfully invaded hepatocytes containing a developing parasite (d), there was no significant inhibition in sporozoites that had invaded and formed a parasitophorous vacuole. (e) Azithromycin treatment of T. gondii tachyzoites at concentrations of 250, 125 and 50 μM resulted in a dose-dependent inhibition of host cell invasion. The invasion inhibitory control cytochalasin D (1 μM) was a considerably more potent inhibitor of tachyzoite invasion, while erythromycin A, as was found for P. falciparum, showed no evidence of invasion inhibitory activity at concentrations up to 500 μM. (f) Azithromycin analogue 12e (42 μM) was significantly more inhibitory to Toxoplasma tachyzoite invasion than azithromycin (250 μM, both solubilized in DMSO) even when tested at a 6-fold lower concentration. Data represent the mean and SEM of three or more experiments, significance of differences was tested using an unpaired t-test (*P = 0.01 to 0.05, **P ≤0.01, ***P ≤0.001)

Since azithromycin was found to be an effective inhibitor of merozoite invasion, we tested whether this drug could also inhibit invasion of P. berghei sporozoites into liver cells in vitro. The effects of azithromycin on P. berghei in vitro liver-stage infections following invasion have been previously reported to be limited to retardation of apicoplast development with no effect on overall parasite growth (up to 65 hours) [46] the effect of azithromycin on sporozoite invasion has not directly been examined. Initially, we assayed the ability of P. berghei sporozoites to enter liver cells in vitro using an established assay. GFP-expressing sporozoites were allowed to settle on and invade HepG2 cells. After free sporozoites were removed by washing, cells were fixed and then antibody-labeled without cell permeabilization, thereby preventing labeling of sporozoites that are inside host cells. We then compared the number of labeled cells (external) to the total number of cells (internal and external, as measured by GFP expression) to determine invasion efficiency. There was a statistically significant, but modest, effect indicating that azithromycin may inhibit sporozoite invasion of hepatocytes at concentrations up to 100 μM azithromycin (100 μM, 42 % reduction in cell entry (azithromycin prepared in ethanol) Fig. 8c). However, this assay is unable to robustly distinguish between sporozoites traversing liver cells from those establishing an infection. Therefore, to assess the impact of azithromycin treatment on the establishment of successful infection of liver cells, we repeated the treatment and then allowed any successfully invaded sporozoites to develop for 24 hours this is a point at which parasites developing within the liver cell can be distinguished from any remaining sporozoites that have not established infection. The number of parasites successfully establishing an infection and forming a vacuole at 24 hours did not differ between azithromycin-treated and control (ethanol-treated) cultures. This suggests that while there was some evidence of reduced sporozoite invasion of liver cells, this did not result in measurably reduced hepatocyte infections by P. berghei sporozoites (Fig. 8d).

We next explored whether azithromycin could inhibit invasion of the related apicomplexan parasite, Toxoplasma gondii, which has orthologues of many Plasmodium invasion proteins and a similar invasion process [47]. There was a significant dose-dependent inhibition of tachyzoite invasion at azithromycin concentrations (in ethanol) between 250 μM and 50 μM (Fig. 8e). Azithromycin inhibited 47 % of tachyzoite invasions at the highest concentration tested (250 μM) compared to 90 % inhibition for the invasion inhibitory control cytochalasin D (1 μM) [48] and the invasion inhibitory IC80 of azithromycin against P. falciparum merozoite (38 μM in ethanol), clearly indicating that azithromycin is a less potent inhibitor of T. gondii invasion in vitro. Importantly, there was no evidence of tachyzoite invasion inhibition in the presence of 500 μM erythromycin A, reflecting the results of the P. falciparum studies which showed that erythromycin A is a poor inhibitor of merozoite invasion compared to azithromycin. We next explored whether the azithromycin analogue 12e, which was found to be a more potent inhibitor of P. falciparum merozoite invasion than azithromycin (DMSO as vehicle), had improved tachyzoite invasion inhibitory activity compared to azithromycin. Analogue 12e (tested at 42 μM due to limited compound availability) was found to be a consistently more potent inhibitor of T. gondii tachyzoite invasion (48 % inhibition compared to non-inhibitory control) than much higher concentrations of azithromycin (250 μM, 24 % inhibition, Fig. 8e), suggesting that analogues of azithromycin could be developed to have much greater potency against T. gondii host cell invasion than the parent compound. These data suggest that azithromycin and analogues are effective inhibitors of Plasmodium spp. merozoite and T. gondii tachyzoite invasion, raising the possibility that azithromycin may target proteins or events conserved between organisms. However, there are clearly differences in drug potency between parasites hence drugs will have to be optimized to target invasion for each organism.



Human insulin was purchased from Sigma-Aldrich (St Louis, MO, USA) and recombinant human IGF1 from R&D Systems (Minneapolis, MN, USA). Monoclonal anti-diphosphorylated ERK1/2 (Thr183, Tyr185) was obtained from Sigma-Aldrich. Anti-phospho-forkhead box O1 (FoxO1 Thr24)/FoxO3a (Thr32) antibody and anti-phospho-p70S6K (Thr412) were purchased from Millipore (Billerica, MA, USA). Anti-GAPDH antibody was purchased from Abcam (Cambridge, MA, USA). Anti-phospho Akt/PkB antibody (Ser473) was purchased from Cell Signaling Technology (Danvers, MA, USA). Horseradish peroxidase-conjugated polyclonal rabbit anti-mouse IgG was purchased from Sigma-Aldrich. Horseradish peroxidase-conjugated goat anti-rabbit F(ab')2 fragment and peroxidase-conjugated goat anti-rabbit IgG (H + L) were purchased from Invitrogen/Life Technologies (Grand Island, NY, USA) and Pierce/Thermo Scientific (Rockford, IL, USA), respectively. The SuperSignal West Pico chemiluminescent detection kit was purchased from Pierce. All other chemicals and reagents were obtained from Sigma-Aldrich or ThermoFisher Scientific (Waltham, MA, USA). Human serum and red blood cells (RBCs) were obtained from Interstate Blood Bank (Memphis, TN, USA).

Mosquito cell culture, mosquito rearing and experimental treatments

The immortalized A. stephensi embryo-derived (ASE) cell line was maintained as previously described (Surachetpong et al., 2009). For in vivo studies, A. stephensi (Indian wild-type strain) were reared and maintained at 27°C and 75% humidity. All mosquito rearing and feeding protocols were approved by and in accordance with regulatory guidelines and standards set by the Institutional Animal Care and Use Committees of the University of California, Davis, and the University of Georgia.

Western blotting

For in vivo studies, female mosquitoes (3–5 days old) were maintained on water for 24–48 h and then allowed to feed for 30 min on reconstituted blood provided through a Hemotek Insect Feeding System (IFS Discovery Workshops, Accrington, UK). This blood meal contained washed human RBCs and saline (10 mmol l −1 NaHCO3, 15 mmol l −1 NaCl, pH 7.0) with or without recombinant human IGF1 or insulin. Midguts were dissected from 30 mosquitoes in each treatment group and processed as previously described (Surachetpong et al., 2009). Control mosquitoes were provided blood meals supplemented with an equivalent volume of IGF1 diluent [0.1% bovine serum albumin (BSA) in phosphate-buffered saline (PBS)].

Detection of IIS proteins followed the protocol of Surachetpong et al. (Surachetpong et al., 2009). In brief, protein lysates from cells or mosquito midguts were separated by gel electrophoresis on 10% sodium dodecyl sulfate-polyacrylamide gels, transferred to nitrocellulose membranes (BioRad, Hercules, CA, USA) and probed for proteins of interest with target-specific antibodies. Membranes were blocked in 5% dry milk/Tris-buffered saline with 0.1% Tween 20 for 1 h at room temperature, and then incubated overnight in each antibody solution. Primary and secondary antibodies were used at the following dilutions: 1:10,000 phospho-ERK (1)/1:20,000 rabbit anti-mouse IgG (2) 1:1000 phospho-FOXO (1)/1:2000 goat anti-rabbit IgG (2) 1:500 phospho-Akt (1)/1:2000 goat anti-rabbit IgG (2) 1:1000 phospho-p70S6K (1)/1:5000 goat anti-rabbit IgG (2) and 1:10,000 GAPDH (1)/1:20,000 goat anti-rabbit IgG (2).

Lifespan studies

A control and two treatment groups of A. stephensi females (N=300 per group, 3 days post-emergence) were fed a weekly artificial blood meal (washed human RBCs and saline, as described above) via a Hemotek IFS (Discovery Workshops) with: (1) 0.013 μmol l −1 (0.1 μg ml −1 ) IGF1, (2) 0.133 μmol l −1 (1.0 μg ml −1 ) IGF1 or (3) an equivalent volume of IGF1 diluent (0.1%BSA/PBS). Dead mosquitoes were counted three times per week, and oviposition cups were provided once a week after blood feeding. The experiment was replicated three times with separate cohorts of mosquitoes.

Preparation of radiolabeled peptides

Lyophilized human insulin (Sigma-Aldrich, 32 μg) was dissolved in 160 μl phosphate buffer (PB, pH 7.4), iodinated with 125 I (MP Biomedical) using chloramine T (Sigma-Aldrich), and separated from reaction byproducts by HPLC, as described in Crim et al. (Crim et al., 2002). The final 125 I-insulin stock concentration was 64.1 nmol l −1 , after adding BSA (Sigma-Aldrich, 10%, 50 μl). Lyophilized IGF1 (Novozymes, 1 mg) was dissolved in 100 μl 10 mmol l −1 HCl as a stock solution. For the reaction, IGF1 stock (20 μg) was diluted 1:10 in PB, radioiodinated using lactoperoxidase (Sigma-Aldrich) and separated as above. The final 125 I-IGF1 stock concentration was 96.4 nmol l −1 after addition of BSA. Radiolabeled peptides were stored at −20°C and used within 27 days after receiving the isotope.

Feeding of radiolabeled insulin and IGF1

Radiolabeled insulin (9.1 μl from stock) was added to aliquots of saline (491 μl 30 mmol l −1 NaCl, 20 mmol l −1 NaHCO3, 2 mmol l −1 ATP, pH 7.0) and stored at −20°C for up to 30 h before use. Radiolabeled IGF1 (20.7 μl from stock) and unlabeled IGF1 (37.7 μl, 2.6 μmol l −1 ) were added to aliquots of saline (441.6 μl) and similarly stored. Washed human RBCs (1.5 ml) were centrifuged at 1000 g for 10 min at 4°C, and 0.5 ml of the cell pellet was transferred from the bottom of the tube and added to the radiolabeled insulin or IGF1 solution immediately before feeding.

Anopheles stephensi females (50–60 individuals, 7–10 days old) kept without sucrose for 18–30 h in a humidified chamber were transferred to individual 500 ml polypropylene feeding chambers fitted with nylon mesh. Glass-jacketed feeders with a Hemotek IFS membrane (Discovery Workshops) were warmed to 37°C, set onto the nylon mesh of the feeding chambers and then filled with a mixture of saline (15 mmol l −1 NaCl, 10 mmol l −1 NaHCO3, 1 mmol l −1 ATP, pH 7.0) and RBC solution with a final concentration of 5.9×10 −4 μmol l −1 insulin or 0.133 μmol l −1 IGF1. Females had access to the feeder for 45 min, after which engorged mosquitoes were sorted from non-engorged mosquitoes at 4°C. Feedings were performed at 11:00, 16:00 and 21:00 h to facilitate 6 h interval collections.

Electrophoresis and autoradiography

Up to 48 h post blood meal (PBM), abdomens and heads + thoraces were dissected every 6 h and separately stored in extraction solution (six heads + thoraces or six abdomens per 100 μl 40% CH3CN, 0.1% trifluoroacetic acid in water) at −80°C. Samples were sonicated for 10 s, lyophilized and resuspended in 20 μl deionized water and 20 μl Tris-tricine gel SDS sample buffer (NuSep). They were then heated (100°C) for 5 min, centrifuged and loaded (three body part equivalents per lane) onto a Criterion 16.5% Tris-tricine gel (BioRad) along with a protein molecular weight marker mix (Kaleidoscope Polypeptide Standards, BioRad). After electrophoresis in Tris-tricine buffer for 1.5 h at 110 V, gels were dried between cellulose sheets and exposed to autoradiography film (Blue Basic Autorad Film BioExpress, Kaysville, UT, USA) at −70°C for up to 28 days.

Radioactivity in female body parts prepared as above was quantified as counts per minute (cpm) per three body part equivalents (20 μl aliquots) on a Cobra II AutoGamma counter (Packard Instrument Company, Meriden, CT, USA). Relative amounts of insulin or IGF1 in the samples were calculated from a regression line obtained for dilutions of the radiolabeled insulin or IGF1 in RBC solutions. Feeding experiments and the above steps were replicated with three different cohorts of female mosquitoes for each of the radiolabeled peptides.

Midgut insulin receptor phosphorylation

After emergence, female A. stephensi were maintained on 10% sucrose for 3 days followed by water for 2 days and then given access to washed RBCs in saline alone or with human insulin (1×10 −4 μmol l −1 ) or human IGF1 (0.013 or 0.133 μmol l −1 ) as above. At 0.5, 1, 3, 12 and 24 h PBM, midguts (20 per sample) were dissected into PhosphoSafe (Novagen/EMD Millipore, Billerica, MA, USA) with 4× protease inhibitor (Complete Mini Roche Applied Science, Indianapolis, IN, USA) and transferred to 1.5 ml centrifuge tubes on ice. As a control, midguts were similarly collected from non-blood-fed mosquitoes. Following centrifugation (4°C, 5000 g, 1 min), supernatant was removed and homogenization buffer (PhosphoSafe 4.75 ml 0.25 ml of 1.0 mol l −1 Tris pH 7.0, 0.43 g sucrose and two tablets of Roche complete mini protease inhibitor) was added to the pelleted midguts (150 μl per sample), which were then sonicated and centrifuged (2000 g, 5 min, 4°C). Supernatants were each transferred to a 1.5 ml high G-force tube on ice. Pellets were resuspended in homogenization buffer (150 μl per sample) and processed as before. This second supernatant was added to the first supernatant and centrifuged (4°C, 48,000 g, 1 h). The supernatant was immediately removed, and the membrane pellet was resuspended with 20 μl of homogenization buffer (one midgut per μl). Samples were stored at −80°C.

Midgut membrane samples were resuspended in Laemmli buffer [20 μl per sample 0.125 mol l −1 Tris (pH 6.8), 50% glycerol, 4% SDS, 0.02% Bromophenol Blue] and sonicated. Following incubation (30°C, 5 min) and brief centrifugation, samples (20 μl per lane) were loaded and separated on a 4–20% Tris-HCl glycine gel (BioRad Criterion 100 V for 3 h at 4°C). Membrane proteins were transferred onto a nitrocellulose membrane (0.1 μm Protran, Whatman/GE Healthcare, Piscataway, NJ, USA 30 V for 2 h at 4°C), which was then dried overnight. The membrane was covered with Tris-buffered saline containing 0.1% Tween 20 (TBS-T) for 2 min and blocked with 2% ECL Advance blocking agent (GE Healthcare) and 2% goat serum for 2 h at 25°C. Thereafter, rabbit anti-phospho-tyrosine antibody conjugated to horseradish peroxidase (Invitrogen) was added to the blocking solution (1:20,000 dilution) and incubated overnight at 4°C. Blots were rinsed for 3×20 min with TBS-T, and immunoreactive proteins were visualized with the ECL Advance kit (GE Healthcare) for image capture (GeneGnome Syngene, Frederick, MD, USA). A total of five immunoblots were obtained from different female cohorts subjected to the same RBC feeding as described above. Each blot was processed and exposed (1–3 min) with identical conditions, and the density of immunoreactive bands corresponding to the native MIR was qualitatively assessed with GeneTools Software (Syngene). The MIR bands in the lanes with midgut membranes from non-fed females were assigned a value of one.

Malaria parasite culture and mosquito infection

Cultures of P. falciparum strain NF54 were grown in 10% heat-inactivated human serum and 6% washed human RBCs in RPMI 1640 with HEPES (Gibco/Invitrogen) and hypoxanthine for 15 days, or until stage V gametocytes were evident. Exflagellation rates of mature gametocytes were evaluated on the day prior to and the day of mosquito infection. Mosquitoes were fed on mature gametocyte cultures diluted with human RBCs and heat-inactivated human serum. All IGF1 treatments were added to the diluted culture just before blood feeding. Human IGF1 in 10% heat-inactivated human serum used for parasite culture ranged from 117 to 210 ng ml −1 (0.015–0.027 μmol l −1 ). This serum is diluted 1:10 for parasite culture in our standard protocol, so concentrations of IGF1 in parasite culture medium prior to recombinant IGF1 supplementation ranged from 0.0015 to 0.0027 μmol l −1 . Thus, IGF1 in the parasite culture medium only minimally increased the total levels of IGF1 in the experiments (see below) and the absolute concentrations of IGF1 used still bracketed the low and high ends of the normal physiological range in humans. A single source of human serum was used for all groups (controls and IGF1 treated) within each parasite infection experiment, and this serum came from non-infected individuals. Protocols involving the culture and handling of P. falciparum for mosquito feeding were approved and in accordance with regulatory guidelines and standards set by the Biological Safety Administrative Advisory Committee of the University of California, Davis.

For mosquito feedings, laboratory-reared 3- to 5-day-old female A. stephensi were maintained on water for 24–48 h prior to blood feeding. The experiment was repeated four times with separate cohorts of mosquitoes. Mosquitoes (N=125 per treatment group) were provided blood meals containing P. falciparum NF54-infected RBCs and treatments of 0.013 μmol l −1 (100 ng ml −1 ) IGF1, 0.133 μmol l −1 (1.0 μg ml −1 ) IGF1 or an equivalent volume of IGF1 diluent (0.1% BSA in PBS) via a Hemotek IFS (Discovery Workshops) and allowed to feed for 30 min. After 10 days, midguts from fully gravid females were dissected and stained with 0.1% mercurochrome to visualize P. falciparum oocysts. The mean number of oocysts per midgut (infection intensity) and the percentage of infected mosquitoes (infection prevalence infection defined as at least one oocyst on a dissected midgut) were calculated for all dissected mosquitoes.

Plasmodium falciparum growth assays

Aliquots of P. falciparum NF54 culture were synchronized for 48 h as previously described (Lambros et al., 1979) and then plated in 96-well flat-bottom plates in complete RPMI 1640 with HEPES, hypoxanthine and 10% heat inactivated human serum. Parasites were treated for 48 h at 37°C with equivalent volumes of PBS and human IGF1 at concentrations ranging from 0.13 nmol l −1 to 1.33 μmol l −1 . Assays were terminated by replacing culture medium with RPMI 1640/1% formalin. Erythrocytes were stained with 10 μg ml −1 propidium iodide (Sigma-Aldrich) in PBS for 1 h at room temperature. Infected RBCs were counted with a FACSCalibur flow cytometer (BD Biosciences, San Jose, CA, USA). Relative levels of parasite growth in response to treatment were normalized to PBS-treated controls, which were set to 100%.

Statistical analyses

Data were tested for normality using Kolmogorov–Smirnov, D'Agostino–Pearson omnibus and Shapiro-Wilk methods (GraphPad Prism 5.02, La Jolla, CA, USA). Normally distributed data were analyzed by ANOVA for overall significance and Bonferroni multiple comparison tests for pairwise comparisons. Non-normally distributed data were analyzed by Friedman's test for overall significance followed by Dunn's multiple comparison tests for pairwise comparisons. Parasite infection intensity and prevalence were analyzed to determine whether IGF1-treated mosquitoes were more resistant than controls. Data were analyzed by ANOVA to determine whether oocyst intensity in the controls differed among replicates. When no significant differences were evident, the data were pooled across replicates and analyzed by Kruskal–Wallis ANOVA to test for overall significance and Dunn's post-test for pairwise comparisons. Parasite prevalence was analyzed by Fisher's exact test to determine whether infection status differed between treatment conditions. Survival analyses were performed using the Kaplan–Meier method (Kaplan and Meier, 1958), and differences between survival curves were calculated using the Wilcoxon test (GraphPad Prism 5.02). All differences were considered to be significant at P<0.05.

Definitions and Methodology: How Does Metabolomics Fit in With System Biology

Malaria has been recognized as a significant global health burden, resulting in approximately 228 million new cases and 405 thousand deaths in 2018 worldwide (Sills et al., 2018 World Health Organization, 2019). Malaria is a vector-borne disease caused by the eukaryotic protozoan parasite of genus Plasmodium which has a complicate life cycle, transmitting from Anopheles mosquitoes to human hosts (Cowman et al., 2016 Haldar et al., 2018 Figure 1). With the urgent demands for better understanding of the molecular biology of the malaria parasite (Miller et al., 2013), system biology has been proven to be a versatile and robust strategy to explore the complex Plasmodium biological process (Hasin et al., 2017 Karczewski and Snyder, 2018 Box 1). Advances in these omics-based approaches have shed light on parasite biology and further revolutionized the research for parasitic diseases (Figure 2 Florens et al., 2002 Gardner et al., 2002 Zhu et al., 2018).

Figure 1. Life cycle of representative plasmodium (P. falciparum). (A) Malaria is initiated through the bite of Anopheles mosquitoes. (B) Injected sporozoites are transported to liver through vasculature and then matured in hepatocytes. (C) After schizogony by producing hundreds of daughter merozoites, released merozoites then invade into erythrocytes for asexual life cycles. (D) A portion of asexually reproducing merozoites undergo reprogramming sexual differentiation to form gametocytes. (E) Matured gametocytes are then released into peripheral circulation for ingestion by a mosquito. (F) Formed zygotes transform into ookinete in the midgut of mosquito.

Figure 2. Overview of the relationship between each omics approach and ground-breaking applications in Plasmodium spp. Figures of significant achievements are adapted from Gardner et al. (2002), Florens et al. (2002), Olszewski et al. (2009), Zhu et al. (2018) and the copyright of each figure has been achieved.

BOX 1. Systems biology: A comprehensive approach for malarial research.

Since 2002, the genome draft of laboratory-adapted and field-isolated strains of P. falciparum have been sequenced which enables the investigation of epidemiological and transmission dynamics in malaria endemic areas. This excellent work has opened a window for promoting and accelerating drug and vaccine development. However, the large evolutionary distance from model organisms has hindered the genome annotation and the exact functions of specific genes are hypothetical, putative, or even unknown, making it extremely challenging in identifying functional genes in cellular processes. With the growing demands to overcome this challenge, downstream functional omics strategies including transcriptomics, proteomics, and metabolomics have been and continue to be put forward to characterize the gene functions and to obtain better understanding of the molecular biology of malaria parasites (Hall et al., 2005 Salinas et al., 2014 Lindner et al., 2019). Since the first attempt of genome annotation, proteomics research has also been performed to identify stage-specific proteomes or to reveal the critical cellar regulatory mechanisms, suggesting its potential in identification of drug targets or developing transmission block strategies. Meanwhile, the successful construction of various databases, such as VarDB, PlasmoDB, and MalVac, also makes it possible for researchers to share findings, providing broader applications of acquired genomic data to accelerate the development of malaria biology (Chaudhuri et al., 2008 Cristina et al., 2009 Lakshmanan et al., 2011).

It has been well characterized that the epigenetic mechanism or the physiological effect of post-translational modifications can be regulated by metabolites. Therefore, changes in the Plasmodium development stages can be captured by omics approaches from a genome to metabolome level (Skretas and Wood, 2005 Saito and Matsuda, 2010 Sperber et al., 2015). Metabolomics is defined as an approach that aims at simultaneously detecting small molecules (ρ,500 Da) to understand the systemic changes in a different state and allows global metabolic profiling in various bio-samples. Metabolomics allows the investigation of metabolic phenotypes associated with gene functions and protein expressions by amplifying changes in transcriptomes and proteomes (Saito and Matsuda, 2010 Horgan and Kenny, 2011 Klassen et al., 2017 Giera et al., 2018). Therefore, metabolome can be visually described as metabolic gears intertwined with genomes and proteomes, which acts as an integral component in the overall system without hypothesizing the effect of any single element. Opposed to conventional 𠇍own-top” strategies, metabolomics works well in system biology because it is capable of providing one “top-down” view of biochemical profiles in complex organisms. A surge in metabolomics studies were seen over the past decades with the incredible development of high-resolution platforms (Dunn and Ellis, 2005 Markley et al., 2017 Misra et al., 2017 Box 2).

BOX 2. Overview of metabolomics.

Application of various analytical platforms is of extreme importance in which the fast development will greatly aid in our capabilities to understand the biology from an overall perspective. Technological advances will hugely contribute to the detection of thousands of compounds when facing complex biofluids. Historically, nuclear magnetic resonance (NMR) spectroscopy and mass spectrometry (MS) have been the two most widely applied platforms with highly complementary attributes in various metabolomics researches.

Although limited by the sensitivity, NMR still provides a window in the qualitative and quantitative analysis for compounds with high abundance present in almost all biofluids, tissues, or cell extractions. Excellent reproducibility alone with non-destructive property for NMR allows all kinds of sample analysis in a short period, making it an ideal candidate in large-scale metabolomics screening. With the growing demand for capturing low-abundance metabolites to meet the requirement of obtaining metabolic patterns as detailed as possible, the MS platform coupled with chromatography could be another powerful and complementary platform for simultaneous analysis of hundreds of compounds in complex biosamples with sufficient resolution, sensitivity, and reproducibility. Until now, the development trend of analytical platforms still mainly focuses on acquiring more metabolic information in a minimized sample, developing high throughput approaches with improved qualitative and quantitative accuracy.

As a hypothesis generating strategy, it mainly focuses on the global assessment of a broad range of both known and unknown compounds to offer either comprehensive or unbiased approaches to investigate unanticipated perturbations. Using this unbiased strategy, changes of metabolic profiles subjected to different conditions can be revealed, leading to the identification of novel metabolites or metabolic pathways. Then subsequent analysis can be performed to structural or functional characterization of these candidates.

As a hypothesis driven strategy, it mainly emphasizes the detection of clearly defined compounds which holds the advantages in obtaining a comprehensive understanding of the kinetic profile in specific pathways.

Functional metabolomics, as one kind of emerging concept, spans from the discovery of differential metabolites to investigate the actual function of specific compounds. Based on untargeted and/or targeted approaches, it further characterizes the functional role of selected metabolites as well as the enzymes in related pathways through both in vitro and in vivo assays including cell biology and clinical platform. It is able to increase the reliability for metabolomics study and contributes novel knowledge about previous findings.


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Plasmodium, a genus of parasitic protozoans of the sporozoan subclass Coccidia that are the causative organisms of malaria. Plasmodium, which infects red blood cells in mammals (including humans), birds, and reptiles, occurs worldwide, especially in tropical and temperate zones. The organism is transmitted by the bite of the female Anopheles mosquito. Other insects and some mites may also transmit forms of malaria to animals.

Five species cause human malaria: P. vivax (producing the most widespread form), P. ovale (relatively uncommon), P. falciparum (producing the most severe symptoms), P. malariae, and P. knowlesi. There are several species that have been isolated from chimpanzees, including P. reichenowi and P. gaboni. P. falciparum, P. gaboni, and other species have been isolated from gorillas. Examples of parasites found in reptiles include P. mexicanum and P. floridense, and those in birds include P. relictum and P. juxtanucleare.

Plasmodium species exhibit three life-cycle stages— gametocytes, sporozoites, and merozoites. Gametocytes within a mosquito develop into sporozoites. The sporozoites are transmitted via the saliva of a feeding mosquito to the human bloodstream. From there they enter liver parenchyma cells, where they divide and form merozoites. The merozoites are released into the bloodstream and infect red blood cells. Rapid division of the merozoites results in the destruction of the red blood cells, and the newly multiplied merozoites then infect new red blood cells. Some merozoites may develop into gametocytes, which can be ingested by a feeding mosquito, starting the life cycle over again. The red blood cells destroyed by the merozoites liberate toxins that cause the periodic chill-and-fever cycles that are the typical symptoms of malaria. P. vivax, P. ovale, and P. falciparum repeat this chill-fever cycle every 48 hours (tertian malaria), and P. malariae repeats it every 72 hours (quartan malaria). P. knowlesi has a 24-hour life cycle and thus can cause daily spikes in fever.

This article was most recently revised and updated by Kara Rogers, Senior Editor.


The World malaria report 2015 reported the reduction of malaria mortality rates by an impressive 48 % between 2000 and 2015 as a result of a major scale-up of vector control interventions, diagnostic testing, and treatment with artemisinin�sed combination therapy [1]. Despite these tremendous achievements, an estimated 214 million cases of malaria occurred globally in 2015, and the disease led to 438,000 deaths, mostly those of children under 5 years of age in Africa [1]. Limited efficacy achieved by subunit vaccine candidates, emerging anti-malarial drug resistances, along with reported insecticide resistances, underline the need of new tools to control and prevent malaria [2, 3]. In this perspective, the development of an effective malaria vaccine is recognized as one of the most promising approaches to conquer the disease. Despite decades of research, an effective vaccine against malaria has remained elusive. Anti-malarial vaccines can break the parasite life cycle at different stages: infection-blocking vaccines targeting hepatic stages, anti-morbidity vaccines targeting the asexual blood stages, and transmission-blocking vaccines targeting the sexual stages. To achieve effective protection, the ideal malaria vaccine is thought to target several steps of the parasite life-cycle in a multistage combination vaccine [4].

Clinical and experimental data support the feasibility of developing an effective malaria vaccine. Adults living in malaria endemic areas rarely experience malaria episodes: partial protection of adults is mediated by naturally acquired immunity, and protects against symptomatic disease and high�nsity parasitaemia, but is not effective in offering sterile immunity [5]. Also, passive transfer of γ-globulin from semi-immune adults to malaria patients conferred a significant reduction of parasitaemia and recovery from clinical symptoms [6]. Those studies showed that immunity can be naturally acquired with exposure and indicated antibodies as crucial components of the protective immune response against asexual blood stage parasites. In this perspective, a multi-stage malaria vaccine should contain as one component antigen(s) that elicit antibody responses upon parasite presentation, leading to clearance of asexual blood stage parasites, and thus reducing the clinical symptoms.

Currently, with a total 25 projects in the pipeline [7], three candidate vaccines are in phase 2B clinical trials and one, the pre-erythrocytic subunit vaccine RTS,S/AS01, has completed phase 3 [8]. In infants aged 6� weeks at first vaccination with four doses of RTS,S reduced the number of cases of clinical malaria by 26 % to the end of the study over an follow-up of 38 months. Blood-stage vaccines, designed to elicit anti-invasion and anti-disease responses [9], are traditionally mainly based on a few protein candidate antigens: apical membrane antigen 1 (AMA1) [10�], erythrocyte-binding antigen-175 (EBA-175) [13], glutamate-rich protein (GLURP) [14, 15], merozoite surface protein (MSP) 1 [16], MSP2 [17, 18] and MSP3 [19, 20] and serine repeat antigen 5 (SERA5) [21, 22]. These immunodominant antigens, highly expressed on merozoites surface or within apical organelles, are involved in the invasion process. Unfortunately, AMA1 and MSP1, the most advanced blood-stage vaccines, have not demonstrated effective protection in African children [10, 16, 23], probably due to their highly polymorphic nature [24]. Genetic variability, extensive polymorphism and antigenic complexity in immunodominant antigens represent major obstacles in the development of an effective blood-stage malaria vaccine [25�]. Identifying and designing antigens able to induce strain-transcending immune responses, which cover antigenic diversity remains a critical issue to be addressed. Since the Plasmodium falciparum genome was sequenced and annotated in 2002 [28], reverse vaccinology represents the most attractive strategy to rationally identify novel malaria vaccine candidates [29, 30]. On the basis of the large-scale genomic, transcriptomic, proteomic and comparative data from Plasmodium spp. that have become available, new antigens with great potential as blood-stage vaccine candidates have been discovered [31].

Among the newly characterized proteins, the cysteine rich protective antigen (PfCyRPA) exhibited remarkable properties: PfCyRPA (1) elicits Abs that inhibit parasite growth in vitro and in vivo [32], (2) is highly conserved among P. falciparum isolates [32], (3) has limited natural immunogenicity, and (4) forms together with the reticulocyte-binding homolog 5 (PfRH5) and the PfRH5-interacting protein (PfRipr) a multiprotein complex crucial for P. falciparum erythrocyte invasion [33]. PfRH5 is currently regarded another leading blood-stage malaria vaccine candidate: it has been shown to induce invasion-inhibitory antibodies that are effective across common PfRH5 genetic variants and PfRH5-based vaccines can protect Aotus monkeys against virulent vaccine-heterologous P. falciparum challenges [34�]. The PfCyRPA encoding gene PFD1130w is located in the subtelomeric region of chromosome 4 in close proximity to other genes playing a crucial role in the erythrocytes invasion, including PFD1145c that encodes for PfRH5 [36]. PfCyRPA is a 362-aa-long protein with a predicted molecular mass of 42.8 kDa, an N-terminal signal peptide, a C-terminal GPI-anchor motif and twelve cysteine residues, potentially involved in the formation of six disulfide bridges. PfCyRPA was identified as a promising blood-stage malaria vaccine candidate exploiting a cell-based approach that utilizes antigens expressed on the surface of mammalian cells for mouse immunization [38]. Since antigen-loaded cells are not suitable for human immunization, the study investigated whether invasion inhibitory anti-PfCyRPA antibodies could be raised by active immunization with purified recombinant PfCyRPA protein. In the present study, PfCyRPA was recombinantly-expressed in mammalian cells and adjuvanted vaccine formulations of purified PfCyRPA were tested for their potential to elicit antibodies that inhibit P. falciparum parasite growth in vitro and in vivo.

The domain on the Duffy blood group antigen for binding Plasmodium vivax and P. knowlesi malarial parasites to erythrocytes.

Plasmodium vivax and the related simian malarial parasite P. knowlesi use the Duffy blood group antigen as a receptor to invade human erythrocytes and region II of the parasite ligands for binding to this erythrocyte receptor. Here, we identify the peptide within the Duffy blood group antigen of human and rhesus erythrocytes to which the P. vivax and P. knowlesi ligands bind. Peptides from the NH2-terminal extracellular region of the Duffy antigen were tested for their ability to block the binding of erythrocytes to transfected Cos cells expressing on their surface region II of the Duffy-binding ligands. The binding site on the human Duffy antigen used by both the P. vivax and P. knowlesi ligands maps to a 35-amino acid region. A 34-amino acid peptide from the equivalent region of the rhesus Duffy antigen blocked the binding of P. vivax to human erythrocytes, although the P. vivax ligand expressed on Cos cells does not bind rhesus erythrocytes. The binding of the rhesus peptide, but not the rhesus erythrocyte, to the P. vivax ligand was explained by interference of carbohydrate with the binding process. Rhesus erythrocytes, treated with N-glycanase, bound specifically to P. vivax region II. Thus, the interaction of P. vivax ligand with human and rhesus erythrocytes appears to be mediated by a peptide-peptide interaction. Glycosylation of the rhesus Duffy antigen appears to block binding of the P. vivax ligand to rhesus erythrocytes.


The Journal of Experimental Medicine &ndash Rockefeller University Press

3. Results

TNF expression is known to be highly enhanced in P. falciparum infection but its role during the asexual stage was not described. The present work showed that TNF is of physiological importance in Plasmodium, as it was able to modulate Plasmodium intra-erythrocytic invasion, similarly as previously described in the liver stages [25].

Parasites where incubated with TNF (1, 2 or 8 ng/ml at a final parasitemia of around 4% (3.98ਊ.u. ±਀.21, n =ਉ). We analyzed possible changes in parasitemia and intra-erythrocytic development by flow cytometry ( Fig. 1 ). Data indicate invasion reduction in the presence of 1 and 2 ng/ml TNF treatment with parasitemia being 13.11% (±ਅ.98 n =ਈ) and 12.72% (±ਉ.4 n =ਇ) lower than untreated control samples. Intra-erythrocytic parasite development was otherwise similar for all other treatments ( Fig. 1 A and B). Representative histogram showing flow cytometry analysis of total parasitemia and intra-erythrocytic stage distribution can be found at (Supplemental material Fig. S4).

Effects of TNF on P. falciparum (3D7) erythrocyte invasion and Ca 2 + concentration in isolated parasites. (A) Flow cytometry analyses of parasitemia in synchronized P. falciparum (3D7) after 48 h of control or TNF (1, 2 and 8 ng/ml) treatment (* P <਀.05) (B) and intra-erythrocytic stages distribution. Bars represent the number of parasites expressed as percentage of control or ring-trophozoites and schizonts (average) normalized by total parasitemia ± S.E.M. Cells were stained with dihydroethidine (5 μg/ml) and data (10 5 cells) were compared by one way ANOVA and by Newman–Keuls test. (C) Spectrofluorimetric analyses of Ca 2 + concentration in isolated parasites labelled with Fluo4/AM (5 μM) after addition of TNF 0.25 0.5 1 or 10 ng/ml (3.05ਊ.u. ±਀.59, n =ꀑ, P =਀.1536 5.35ਊ.u. ±ਁ.01, n =ꀔ, P =਀.064 9.476ਊ.u. ±ਁ.03 n =ꀕ, P =਀.001 and 7.91ਊ.u. ±ਁ.65, n =ꀐ, P =਀.034, respectively). P values were calculated by comparison with the PBS data (1.505ਊ.u. ±਀.07, n =਄). Bar graph means and SEM of at least three independent experiments. Arb. arbitrary. (D) Representative tracing of Fluo4/AM changes over time by addition of TNF (0.25 0.5 1 and 10 ng/ml and PBS) in P. falciparum isolated parasites.

Next we verified the capacity of TNF to modulate calcium signaling in P. falciparum parasites. For this purpose, saponin-isolated parasites were incubated with Fluo-4/AM and spectrofluorimetric analyses showed that TNF can increase free intracellular calcium concentration ([Ca 2 + ]i) ( Fig. 1 C and D) with the highest transient increase observed at 1 ng/ml. These quantities reflect typical values used in in vitro systems but are towards the higher end of the physiological levels encountered in the human host where TNF concentrations up to 1ng/ml can be found in the plasma of patients with severe disease [30], [31].

The use of kinase inhibitors has been explored for the control of inflammation [15], [32] and the inhibitor KN62 was previously shown to modulate Plasmodium pathways [20], [21]. Interestingly KN62 is a Ca 2 + /calmodulin-dependent protein kinase II inhibitor and P2X7 antagonist that also blocked [Ca 2 + ]i increase after TNF addition ( Fig. 2 A & B), indicating the involvement of the pathways in TNF signaling. Next A438079, a competitive P2X7 receptor antagonist also inhibited the TNF/ IE response ( Fig. 2 A & B). Membrane integrity after TNF treatment was studied following staining isolated parasites with WGA by confocal microscopy imaging, revealing no effects of this factor on membrane integrity, as WGA localization in the membrane was the same for TNF-treated and control samples ( Fig. 2 C and Supplemental data S1 and S2).

TNF intracellular calcium signalling can be blocked by P2X7 antagonist in P. falciparum isolated parasites labelled with Fluo4/AM (5 μM). (A) Analyses of Ca 2 + concentration after incubation with the purinergic inhibitor KN62 (10 μM) for 30 min, A438079 (5 μM) for 2 min or thapsigargin (10 μM) all followed by addition of TNF (1 ng/ml) (2.15ਊ.u. ±਀.19, n =ꀓ, P <਀.0001 1.76ਊ.u. ±਀.24, n =ꀑ, P <਀.0001 or 1.236ਊ.u. ±਀.25 n =ꀑ, P <਀.0001, respectively). P values were calculated by comparison with the TNF (1 ng/ml) data (9.746ਊ.u. ±ਁ.03, n =ꀕ). Bar graph means and SEM of at least three independent experiments. Arb. arbitrary. (B) Representative tracing of Fluo4/AM changes over time by addition of PBS, TNF (1 ng/ml) and TNF (1 ng/ml) after incubation with KN62 (10 μM) or A438079 (5 μM) (C) Imaging of wheat germ agglutinin (WGA) staining in P. falciparum (3D7) saponin isolated parasite after TNF (1 ng/ml) treatment for 1 h at 37 ଌ. Smears are stained with WGA (10 μg/ml for 10 min at 37 ଌ) and DAPI (1:1000 in PBS 15 min at RT) and observed by confocal microscopy.

Plasmodium intracellular calcium homeostasis is known to be regulated by endoplasmic reticulum (ER) among other organelles [33], [34]. Here we showed using thapsigargin (THG) that the organelle is essential for the calcium response elicited by TNF as there is no response after ER calcium depletion by THG ( Fig. 2 A & B).

Extracellular calcium can also be involved in intracellular ion homeostasis so parasites were incubated with EGTA (2 or 5 mM). This was capable of decreasing the calcium ion rise at a concentration of 2 mM, and completely abolished at 5 mM ( Fig. 3 ). The parasite viability during the experiment, in terms of its ability to respond, following THG addition was confirmed at the end of the experiment ( Fig. 3 ).

Extracellular calcium is required for TNF signalling in isolated P. falciparum labelled with Fluo4/AM (5 μM). (A) Analyses of Ca 2 + concentration after addition of TNF (1 ng/ml) (9.746ਊ.u. ±ਁ.03, n =ꀕ) or incubation with the extracellular calcium chelator EGTA (5 mM) for 1 min followed by addition of TNF (1 ng/ml) (1.002ਊ.u. ±਀.005, n =ꀑ, P =਀.371), EGTA (2 mM) (1.304ਊ.u. ±਀.038, n =ਉ, P =਀.001) or PBS treatment (0.989ਊ.u. ±਀.016, n =ਃ). Bar graph means and SEM of at least three independent experiments. Arb. arbitrary. (B) Representative tracing of Fluo4/AM changes over time by addition of TNF (1 ng/ml) or TNF (1 ng/ml) followed by thapsigargin (10 μM) after incubation with EGTA (2 and 5 mM, respectively).

As TNF reduces parasitemia we have searched for a target that might be involved in the control of cell cycle proliferation. A possible target involved in the TNF response could be P. falciparum Proliferating-Cell Nuclear Antigen 1 (PfPCNA1), which may be differentially phosphorylated by TNF treatment [35]. We have performed quantitative PCR to verify the expression of PfPCNA1. Since TNF increases cytosolic calcium concentration in P. falciparum and calcium is well known to cross talk with cAMP within several cells including malaria parasites [26], [27] we pre-treated P. falciparum not only with TNF but also with 6-Bnz cAMP. According to Fig. 4 , P. falciparum synchronized trophozoites treated with TNF (1 ng/ml) or 6-Bnz cAMP (20 μM) have a decrease in PfPCNA1 mRNA expression in comparison to control treatments (PBS). The data point to PfPCNA1 as a downstream target of the TNF�lcium�MP signaling pathway, perhaps for reducing cell cycle proliferation, but more work is needed to confirm this. Other candidates studied (e.g. PfRACK) showed no change upon TNF treatment (Supplemental data — Fig. S3).

Expression of mRNA in synchronized trophozoites treated with TNF and 6-Bnz cAMP in PfPCNA1. Real Time PCR for P. falciparum (3D7) control (PBS), TNF (1 ng/ml) or 6-Bnz-cAMP (20 mM), after incubation for 1 h at 37 ଌ. Bars represent mean ± S.E.M. in 3 independent experiments (P <਀.05*).

It is well established that inhibitors of TNF production reduced IE cytoadherence [9], [10]. As FK506 is a potent suppressor of inflammation and the endothelial cell receptor ICAM-1 expression is increased by pro-inflammatory cytokines [36], we have analyzed if FK506 inhibitor would affect the P. falciparum binding under flow and static conditions to HDMEC.

Fig. 5 (A and B) shows the results of experiments performed under flow conditions where parasites were first treated with FK506 (1.5 and 6 μM), calyculin (2 and 10 μM) or Nfkkb inhibitor (2 μM). Higher concentration of calyculin (10 μM) induced an increase in parasite binding indicating the involvement of phosphatases in cytoadherence. While at higher concentration FK506 (6 μM) and Nfkkb inhibitor (2 μM) showed a decrease in binding to HDMEC and highlight the potential importance of the host immune response on parasite signaling and sequential binding to endothelial cells. Under static protein assays ( Fig. 5 C and D) reduction in binding was not statistically different between FK506 at lower concentrations (0.375, 0.75 or 1.5 μM) compared to the controls, DMSO (0.04 %) or untreated parasites.

Effects of FK506 in Plasmodium-endothelial cell binding under flow and static conditions. (A) Adhesion of trophozoite-stage A4 IE after FK506 (1.5 or 6 μM), calyculin (2 or 10 μM), NFƘᦋ inhibitor (2 μM), DMSO (0.04%) or no treatment of parasites under flow assay showing bound IE per mm 2 or (B) percentage of control (DMSO treated) binding to human dermal microvascular endothelial cells (HDMEC). (C) Adhesion of trophozoite-stage ItG IE to protein ICAM-1 after FK506 (0.375, 0.75 or 1.5 μM), DMSO (0.04%) or no treatment under static conditions to ICAM1 showing bound IE per mm 2 or (D) percentage of control (DMSO treated) binding. Bars represent mean ± S.E.M. Note differences in A4 erythrocyte binding among 3𠄵 independent experiments (P <਀.05 *, P <਀.01 **, P <਀.001 ***) under flow condition and no difference observed in ItG erythrocyte binding among three independent treatments (P >਀.05) under static condition.

To determine whether the ability of parasites (treated as described in Fig. 5 ) to adhere was due to PfEMP-1 expression on the infected erythrocyte membrane we performed flow cytometry measurements using mAb BC6, specific for a surface epitope of the PfEMP-1 ITvar14 (A4var) variant [37]. The results clearly demonstrate that PfEMP-1 expression is not changed under the treatments ( Fig. 6 ).

Analysis of PfEMP-1 expression on the surface of live trophozoite-stage A4 IE using BC6 antibody (A) Quantitation of PfEMP-1 levels by flow cytometry analysis after BC6 primary antibody (Alexa 488 coupled secondary antibody) and counterstaining infected erythrocyte with ethidium bromide. (B) Density plots of uninfected erythrocyte population in the left and box IE (infected erythrocyte) representing BC6 positive erythrocytes (gated as ethidium bromide positive) (C) Representative graph showing quantitation of BC6 positive fluorescence intensity (X-axis APC-A) of the infected population without BC6 for negative control (red line) with BC6 and treated with DMSO 0.04% (black line) or (D) FK506 1.5 μM (black line) treated A4 trophozoites. Bars represent mean fluorescence intensity for PfEMP-1 positive population ± S.E.M. for 5 independent experiments. Note no difference in erythrocyte PfEMP-1 quantitation levels (P >਀.05).

3 Concluding remarks

Here, we have described several nutrient sensing pathways in P. falciparum that respond to variations in sugars, amino acids, and lipids. These nutrients result in different types of changes in gene expression, including transcriptional induction, epigenetic regulation, as well as translational repression. However, in almost all cases, the complete pathway from nutrient through metabolite, to sensors and gene expression changes are not fully delineated (Figure 1). For instance, although PbKIN and PfeIK1 appear to act as sensors for cellular energy and amino acid levels, respectively, the downstream effectors of these sensors and functional conservation of PfKIN in P. falciparum remain poorly defined. Moreover, primary versus secondary changes will need to be distinguished through fine temporal studies. Powerful postgenomic tools of metabolomics, transcriptomics, proteomics, and acetylomics, have now been developed to monitor downstream molecular and biochemical changes under different nutritional environments both in vitro and in vivo. Reverse genetics analysis of the nutrient sensor proteins can reveal their interaction with upstream effector metabolites using chemical genetic approaches, as well as their downstream transcriptional programs.

Several intriguing questions do arise relating to the in vivo relevance of nutrient sensing pathways. To what extent is nutritional variation sufficient to elicit gene expression and phenotypic changes in parasites in vivo? Does parasite infection of erythrocytes of different age have an impact on the metabolic state of parasites, which may be of particular relevance to the related P. vivax parasite? Do low and high transmission settings result in different epigenetic profiles (Rono et al., 2018 ), allowing parasites to vary their response to nutritional differences? How relevant is the nutritional status of the parasite to the efficacy of therapeutics? Moreover, several metabolic and epigenetic molecules have been proposed as the direct targets for antimalarials (reviewed in Carolino & Winzeler, 2020 Coetzee et al., 2020 Huthmacher et al., 2010 Kumar et al., 2018 Nawaz et al., 2020 Subbayya et al., 1997 ). It is important to know how these genes function in different metabolic environments, such as in severe malaria, or patients with metabolic disorders like diabetes, and children with malnourishment. So far, parasite metabolism and epigenetic processes have been largely studied independently. However, a picture is emerging of significant crosstalk between these two key processes, and further studies can elucidate their connection.

In conclusion, it will be critical to investigate the interdependency between metabolites, sensor proteins and gene expression changes as an interlocked molecular mechanism used by the parasite to survive under different environmental conditions. Interventions targeting any arm of these complex pathways could have profoundly variable impacts on parasite survival and transmission under diverse metabolic conditions.

Watch the video: Plasmodium vivax (June 2022).


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