Abstract
The pathology associated with malaria infection is largely due to the ability of infected human erythrocytes to adhere to a number of receptors on endothelial cells within tissues and organs. This phenomenon is driven by the export of parasite-encoded proteins to the host cell, the exact function of many of which is still unknown. Here we inactivate the function of one of these exported proteins, PFA66, a member of the J-domain protein family. Although parasites lacking this protein were still able to grow in cell culture, we observed severe defects in normal host cell modification, including aberrant morphology of surface knobs, disrupted presentation of the cytoadherence molecule PfEMP1, and a total lack of cytoadherence, despite the presence of the knob associated protein KAHRP. Complementation assays demonstrate that an intact J-domain is required for recovery to a wild-type phenotype and suggest that PFA66 functions in concert with a HSP70 to carry out host cell modification. Strikingly, this HSP70 is likely to be of host origin.
Taken together, our data reveal a role for PFA66 in host cell modification, implicate human HSP70 as also being essential in this process, and uncover a KAHRP-independent mechanism for correct knob biogenesis. Our observations open up exciting new avenues for the development of new anti-malarials.
Introduction
Plasmodium falciparum causes the most severe form of malaria in humans, malaria tropica, responsible for over 200 million clinical cases and 400,000 deaths per annum, mainly in children under the age of 5 and mostly in sub-Saharan Africa1. The pathology associated with malaria infection is largely due to the ability of infected human erythrocytes to adhere to a number of receptors on endothelial cells within tissues and organs1. This cytoadherence results in reduced blood flow in the affected areas, hypoxia and (in cerebral malaria) increased intracranial pressure1,2. The phenomenon of cytoadherence results from parasite-induced host cell modification in which parasite-encoded proteins are transported to and exposed at the surface of the infected host cell, where they mediate endothelial binding and antigenic variation3–5. In addition to these surface proteins, parasites also encode, express, and export a large number of other proteins to the infected erythrocyte6–8. Many of these proteins are specific to P. falciparum, and their function is still not well understood, partly due to limitations in reverse genetic systems8–10.
Within the predicted ‘exportome’ are 19 proteins belonging to the family of J-domain proteins (JDPs, also called Hsp40s), and this exported family appears to be expanded in the Laveranian clade, suggesting important functions in these parasite species8,11. In other systems, HSP40s act as co-chaperones for HSP70, a protein family that lies at the heart of proteostasis and other essential cellular processes12. Previous studies have localised several members of the exported PfHSP40 family to various structures within the infected erythrocyte, including red blood cell plasma membrane, Maurer’s clefts, knobs, and J-dots10,13,14. Knockout studies suggest that, although some of the exported HSP40s are essential, others can be deleted without any observable phenotype, and several can be deleted, resulting in aberrant cellular morphology, cytoadherence, and rigidity of infected erythrocytes, suggesting a potential role for this protein family in host cell modification and virulence characteristics10 (Supplementary Table 1). One exported HSP40, PFA66 (encoded by PF3D7_0113700, formerly PFA0660w), was previously localised to J-dots, novel structures within the infected erythrocyte containing further exported parasite-encoded HSP40s, an exported parasite-encoded HSP70 (PfHsp70-x), and a number of other exported proteins13,15,16. An earlier medium throughput knockout study failed to generate parasites deficient in PFA66, and therefore its function in the parasite’s lifecycle remains elusive9.
In this study, we utilise selection-linked integration-targeted gene disruption (SLI-TGD) to generate parasites expressing a non-functional PFA66 truncation mutant and characterise the resulting parasite lines. We find that inactivation of PFA66 function leads to dramatic aberrations in host cell modification, especially in knob morphology, capacity for cytoadherance and surface exposure of the virulence factor PfEMP1. Our data suggest an important role for exported HSP40s in parasite pathogenicity. Additionally, our data strongly implicate residual human HSP70 in parasite-induced host cell modification.
Results
Generation of PFA66 truncation and complementation cell lines
Genetic manipulation via single crossover was performed using a selection-linked integration targeted gene disruption strategy17 (Figure 1A). Integration of the plasmid would lead to the production of a GFP-tagged, truncated, non-functional PFA66 protein lacking the entire C-terminal substrate binding domain (SBD), referred to as dPFA. We transfected CS2 parasites that had previously been freshly selected for binding to chondroitin-sulphate-A as this binding phenotype would be essential for later characterisation18,19. Plasmid integration into the PFA0660w/PF3D7_0113700 locus was verified via PCR using primers designed to yield products upon integration of the plasmid into the genome via primers spanning the integration site (Figure 1A). Appearance of bands representing the 5’ and 3’ integration, as well as the disappearance of bands representing the endogenous PFA0660w locus, demonstrated specific integration of the plasmid into the genome (Figure 1B), yielding parasite line CS2ΔPFA (referred to as ΔPFA). Immunodetection using an α-GFP antibody revealed the presence of dPFA::GFP fusion in cell lysates derived from ΔPFA but not parental CS2 parasites (Figure 1C). T ogether, this data indicated successful integration of the vector into the PFA0660w locus, leading to the production of a truncated product dPFA lacking the J-domain required for function. To verify that any aberrant phenotypes observed were due to a lack of PFA66 and not second site events, we generated a complementation line that expressed a full-length, functional 3xHA (hemagglutinin)-tagged copy of PFA66 from an episome, under the control of PromPFA66, referred to as ΔPFA[PFA::HA]. Expression of this complementation construct was verified by immunoblotting using α-HA antibodies (Figure 1D).
dPFA is soluble within the host cell cytosol
Full-length PFA66 has previously been localised to the J-dots, and a follow up study suggested that the SBD of exported HSP40s is required for correct localisation13,20. As dPFA lacks the SBD, but still contains both an N-terminal signal peptide and a recessed PEXEL trafficking signal, we expect export of dPFA::GFP to the host cell6,7. Live cell imaging and immunofluorescence failed to detect dPFA, likely due to the low expression level of this protein. We therefore used equinatoxin (EQT) to selectively permeabilise the erythrocyte plasma membrane and allow sub-cellular fractionation of infected erythrocytes21,22. Immunodetection using antibodies against the compartment-specific markers SERP (parasitophorous vacuole), human Hsp70 (HsHSP70, red blood cell cytosol), aldolase (ALDO, parasite cytoplasm), and GFP reveals co-fractionation of dPFA with HsHSP70, verifying that dPFA::GFP is found in the host cell cytosol and furthermore that dPFA::GFP is found in the soluble phase and not in the membrane fraction as we have previously demonstrated for the full-length protein13 (Figure S1A). Taken together, these data suggests that deletion of the SBD of PFA66 leads to a non-functional protein.
Truncation of PFA66 affects novel permeability pathway (NPP) activity and confers a small growth advantage
Exported parasite proteins carry out a multitude of functions supporting the survival of P. falciparum parasites. One of these is the establishment of NPPs of the iRBC to support the uptake of essential nutrients23,24. To investigate a potential role of PFA66 in NPP activity, we used a sorbitol uptake / lysis assay, which revealed that erythrocytes infected with ΔPFA parasites show a significantly reduced sorbitol-induced lysis, implying a reduced NPP capacity (Figure S1B). To examine whether this reduction in NPP activity affects parasite viability, growth of both cell lines was compared over four growth cycles (~8 days) using flow cytometry. Surprisingly, a slight but significant growth advantage (calculated as below 1% advantage per cycle) of the truncation cell line compared to the parental cell line was observed (Figure S1C).
Truncation of PFA66 causes deformed knob morphology
Knobs are electron-dense protrusions of the iRBC surface that help correctly present the major virulence factor PfEMP1, thus facilitating iRBC cytoadhesion and concomitantly increasing clinical pathology3-5,25,26. As exported HSP40s have previously been implicated in knob formation9,14, we used scanning electron microscopy to visualize knobs on the surface of CS2- and ΔPFA-infected erythrocytes. RBCs infected with the parental parasite line CS2 showed normal knob morphology, with an even distribution of small knobs over the entire surface of the infected cell (Figure 2A, S2A). Small variations in knob number and size are observed and are likely due to slightly different developmental stages of the parasite (compare 2B, C to S6A, B, D, E). In stark contrast, RBCs containing ΔPFA displayed, in addition to a population of normal knobs, knobs with extremely aberrant morphology. These knobs varied in their aberration, and included vastly extended knobs (Figure 2A), wide tall knobs (Figure S2), wide flat knobs (Figure S2), and branched knobs (Figure 2A). Some of the elongated knobs reached lengths of ~0.7 μm. To distinguish these abnormal structures from classical knobs, we will refer to them as “mentula” (plural mentulae). For purposes of quantification, we counted and classified knobs/mentulae to one of three classes I) normal/ small knobs II) abnormal/enlarged mentulae III) deformed/elongated mentulae. This analysis revealed a significant reduction in the overall number of knobs/mentulae in ΔPFA-infected RBCs when compared to wild type CS2 (CS vs ΔPFA, Figure 2B). 22% of surface structures exhibited abnormal morphology in ΔPFA-infected RBCs (Figure 2C). Interestingly, we occasionally observed extended knoblike structures on the surface of RBCs infected with CS2 parasites (Figure 2C) at a level of 2%. As a control, we complemented ΔPFA function with full-length PFA66 expressed from an episome, and both density and morphology of mentulae/knobs returned to wild type levels (Figure 2B, C).
Transmission electron microscopy (TEM) on thin sections prepared from RBCs infected with either WT CS2 or ΔPFA parasites substantiated our observations. CS2 parasites produced clearly defined electron-dense knob structures of a restricted diameter and height above the erythrocyte plasma membrane, whereas ΔPFA displayed mentulae extending from the red blood cell surface into the external medium (Figure 2D, S3). The lumen of these mentulae was often extremely electron dense, hinting at their molecular relation to knobs. Occasionally we observed membrane-bound structures extending from RBCs infected with CS2 parasites, but the lumen of these structures was not electron dense; therefore, these structures cannot be classed as mentulae. Although not highly abundant in either sample, the morphology of Maurer’s clefts appeared comparable in both samples (data not shown).
Electron-dense material is a scaffold for mentula structure
Intrigued by the apparent electron-dense core of mentulae in TEM, we conducted electron tomography analysis on thick sections prepared from ΔPFA-infected erythrocytes. Subsequent 3D reconstruction and surface rendering of the distinct densities in tomographic volumes allows a high-fidelity glimpse into the fine structure of mentulae. This analysis verified the presence of electron-dense material within and at the base of mentulae (Figure 2E). This material fills out the entire mentula rather than only lining the structure, and the distribution of this material closely matched the structure of the mentula. In some cases (3 out of 10), the electron-dense material at the base of the mentula was connected via a thin bridge to the material inside of the mentula (Figure 2F, Supplementary Video 1).
KAHRP distribution is changed in ΔPFA-infected erythrocytes
KAHRP has long been held to be a crucial knob-associated protein as parasites lacking this protein no longer form knobs25,26, and KAHRP truncations show varying aberrant knob phenotypes27. For this reason we investigated the localisation of KAHRP in RBCs infected with our ΔPFA cell line. Indirect immunofluorescence (IFA) on fixed cells demonstrated a punctate distribution of KAHRP in cells infected with both wild type and mutant cell lines (Figure 3A). Automated analysis with a self-generated ImageJ plugin revealed that KAHRP +ve structures were equally numerous but statistically larger in diameter in RBCs infected with ΔPFA than with CS2 (Figure 3B, C). Localisation of other exported parasite proteins via IFA showed no significant difference between the cell lines (Figure S4). To exclude that the KAHRP result was due to non-specific binding of antibodies, for example, we episomally expressed a KAHRP::mCherry fusion in both wild-type and ΔPFA parasites. Surprisingly, considering the relatively low resolution of live cell epifluorescence microscopy, (but preserving membrane integrity and cell morphology), KAHRP +ve structures could be seen to emerge from the surface of RBCs infected with ΔPFA, possibly representing mentulae (Figure 3D, Supplementary Videos 2A-D). Based on the previous result, we wanted to understand whether KAHRP is directly associated with mentulae and used immunogold labelling of thin sections derived from CS2 and ΔPFA iRBCs to localise KAHRP. Although we encountered high background staining of the RBC cytosol in both cases, analysis revealed considerably more label was associated with knobs/mentulae than with the cytosolic background (Figure 3E, F). Analysis of the location of the KAHRP label in relation to the length of the knob/mentula or the closeness to the RBC membrane (RBCM) showed no difference between WT and mutant, or between knob/mentula morphologies (Figure S5A, B). To gain more insight into the nature of the KAHRP +ve structures, we paired membrane shearing with immunolabelling and STED (stimulated emission depletion28) microscopy to study the nature of the KAHRP +ve structures observed above. RBCs infected with CS2 or ΔPFA were allowed to bind to a cover slip, hypotonically lysed to obtain access to the internal leaflet of the RBC plasma membrane, fixed, immunodecorated using an α-KAHRP antibody, and imaged via STED microscopy. This technique allows super-resolution visualisation of KAHRP +ve structures from the luminal side of the erythrocyte plasma membrane and can be used to monitor the assembly of KAHRP into knobs (and in this case, mentulae). This analysis revealed a punctuate distribution of KAHRP beneath the membrane of RBCs infected with both wild type and mutant ΔPFA cells. Although the data initially suggests that there are more KAHRP +ve structures in membranes from ΔPFA, automated counting and measurement shows that this impression is likely to be imparted due to a higher mean size of KAHRP +ve objects rather than an actual increase in the absolute number of objects (Figure 3G, H, I). KAHRP +ve structures from ΔPFA, as well as being larger, also appear by eye to have a lower circularity, although we were not able to substantiate this with image analysis. This might be in part due to the differences in morphology between knobs and mentulae.
Mentulae contain ring/tubelike KAHRP structures but only small amounts of actin
To further investigate the relation of KAHRP with mentula we used rSTED (rescue-stimulated emission depletion29) to image both RBCs infected with CS2 or ΔPFA parasites episomally expressing KAHRP::mCherry as above (Figure S5C). Infected cells were treated with RFP booster to amplify the mCherry signal, fluorescently labelled wheat germ agglutinin (WGA) to label the RBC glycocalyx (delineating the RBC membrane, RBCM), and phalloidin to stain host cell actin (Figure 4).
Line scan analysis reveals that mentulae are bounded by the RBCM. In most cases, KAHRP is found between the RBCM and the actin cytoskeleton, and the KAHRP signal also extends into the central cavity of the mentulae. Distribution of host actin closely follows that of WGA, apart from at the base of mentulae where it follows a path below the KAHRP staining. Fortuitous sectioning of several membrane-bounded mentulae extending from the RBC reveals a ring of KAHRP staining lining the luminal face of the mentula. Phalloidin staining was absent in this case, although it was associated with a shorter mentula (Figure 4, see Figure S5D for CS2 cell line).
Chelation of membrane cholesterol but not actin depolymerisation or glycocalyx degradation causes reversion of the mutant phenotype in ΔPFA
A number of lipid- or protein-dependent mechanisms can induce the initiation of curvature of biological membranes and stabilisation of the resulting structures30. The mentulae we observe here extend up to 0.7 μm from the surface of the red blood cell and appear stable enough that we were able to observe them in live cell imaging. To understand how such structures can be generated, we treated erythrocytes infected with ΔPFA parasites with cytochalasin-D (cyto-D) to depolymerise actin, methyl-β-cyclodextrin (MBCD) to chelate membrane cholesterol, or the glucosidases hyaluronidase (HA) and neuraminidase (NA) to degrade RBC glycocalyx. Following treatment, cells were fixed and prepared for SEM, followed by image acquisition and analysis. Neither cyto-D nor HA/NA treatment caused a statistically significant reduction in the number of mentulae in cells infected with ΔPFA (Figure S6A, B, C, D, E). Treatment with MBCD, while not causing a total reversion of mentulae to knobs, did cause a statistically significant alteration in the observed type of mentulae, with a decrease in the number of deformed, elongated mentulae, and an increase in abnormal, enlarged mentulae.
PFA0660w truncation results in negligible cytoadhesion and aberrant PfEMP1 presentation
As previously mentioned, mature stage infected P. falciparum iRBCs develop additional adhesive capabilities to cells of the host, and this process underlies P. falciparum pathogenicity1. IRBC cytoadherence to purified ligands can be assessed with a binding assay18. Prior to genetic manipulation we selected our parasite population for expression of the var2CSA variant of PfEMP1 by repeatedly binding them to chondroitin sulphate-A (CSA)19. We chose to use the CS2 strain of P. falciparum as this strain stably expresses var2CSA18. A static cytoadherence assay against immobilised CSA demonstrated that, in direct comparison to the parental CS2 strain, ΔPFA exhibits massively reduced levels of binding (Figure 5A). Although all experiments were carried out on parasites that had not been maintained in culture for extended time periods (to avoid switching to another var/PfEMP1 variant), we wanted to verify that ΔPFA still expressed PfEMP1var2csa. We therefore used flow cytometry on intact iRBCs using antisera specific against VAR2CSA (a kind gift of Benoît Gamain) to verify VAR2CSA expression and surface exposure. On erythrocytes infected with ΔPFA, surface expression of VAR2CSA was reduced 60% compared to wild type parasites (Figure 5B, Figure S6F for histograms). Immunofluorescence (IFA) using the same antisera on fixed cells demonstrated that both CS2 and ΔPFA express VAR2CSA to similar levels, with punctuate staining distributed across the host cell. A control IFA on cells infected with 3D7 strain parasites that had not been selected for VAR2CSA expression showed only background fluorescence, verifying the specificity of the result (Figure S6G).
Complementation of PFA66 function requires an intact J-domain
HSP40s such as PFA66 are known interactors and regulators of HSP70s11,12,31, and HSP70-independent functions of HSP40s are rare. The J-domain of HSP40s is crucial both for recruitment of the partner HSP70 and stimulating the ATPase activity of the HSP70 partner31. Having shown above that we can achieve functional complementation via episomal expression of a full-length copy of PFA66, we were interested to know whether a functional J-domain (and hence a functional interaction with a HSP70) is required for phenotypic complementation. To this end, we expressed in the ΔPFA parasite line a full-length copy of PFA66 with a H111Q amino acid replacement converting the HPD motif of the J-domain into the non-functional QPD sequence (Figure S6H) and assayed knob/mentula morphology and density as a surrogate for all other phenotypic assays. While expression of full-length PFA66 was able to revert the mutant knob phenotype (Figure 2B, C), similar expression of the non-functional full-length protein failed to complement the wild-type phenotype (Figure 5C, D) in either density or morphology of knobs/mentulae.
Discussion
Although it has long been recognised that malaria parasites export a substantial number of proteins to their host cell, the mature human erythrocyte, the function of many of these proteins remains unknown6,7,10. P. falciparum exports a larger number of proteins to the host cell than related species, and one family that is highly represented amongst this expansion is that of J-domain proteins (Hsp40s)8. A previous medium-throughput study identified the exported Type II Hsp40 PFA66 as likely to be essential and resistant to inactivation via double-crossover integration9. In this study, we have used selection-linked integration (SLI)17 to generate parasites expressing a severely truncated form of PFA66. This strategy deletes the entire substrate-binding domain of the expressed protein, resulting in a non-functional truncation mutant that is incapable of carrying out its biological function, and may well exhibit a dominant negative effect. The resulting parasites are highly viable but exhibit severe abnormalities in host cell modification.
Parasite growth following truncation of PFA66 was slightly higher than that of wild type CS2 parasites, this effect only becoming evident after four growth cycles. A similar effect was previously observed in a ΔGEXP07 parasite line; however, the significance of this result remains unclear as the reduction in metabolic burden by the loss of only one gene/protein is likely to be negligible32. Lysis of infected erythrocytes in a sorbitol uptake assay was also slightly reduced, implying either a reduction in novel permeation pathway activity, or potentially an increase in host cell stability via mechanical means. As a reduced NPP activity would be expected to lead to slower, not faster, parasite growth, we interpret this to be the result of increased robustness of the erythrocyte plasma membrane through an unknown mechanism.
The most striking result of our study was the observation that, in the ΔPFA66 cell line, normal knob biogenesis was significantly inhibited with regard to both knob density and morphology. Although earlier knockout studies have observed a reduction in knob formation, or slight alteration in knob morphology, to our knowledge ours is the first study to demonstrate such a dramatic alteration in knob structure upon inactivation of a single gene9,32. Indeed, so different are the structures we observe to classic knobs that we suggest calling them mentula to distinguish them from the ‘normal’ surface extensions. Mentulae differ from knobs both in their size/length, reaching up to 0.7μm from the erythrocyte surface. Additionally, mentulae that split into separate branches can be observed. KAHRP, a protein known to be required for correct knob formation, can be localised to mentulae, although its distribution (based on live cell imaging, immunofluorescence, and membrane shearing paired with STED microscopy) seems to be different from that observed in cells infected with wild type parasites. Several studies suggest that KAHRP is integrated into higher order assemblies during parasite development, eventually resulting in a ring structure underlying the knob33,34. Deletions in specific KAHRP domains lead to less incorporation into such structures and also appear to influence the generation of sub-knob spiral structures of unknown molecular composition34. The possibility exists that KAHRP, while being necessary for correct knob formation, is itself not the major structure-giving component but merely serves as a scaffold for assembly of further higher order molecular structures, which themselves generate the necessary vector force to allow membrane curvature and push the knobs above the surface of the erythrocyte. KAHRP has been proposed to be especially vulnerable to misfolding due to its unusual amino acid composition, which would make it a likely client for chaperones / cochaperone systems14. Following this logic, if PFA66 is involved in the assembly of KAHRP into higher order assemblies, interruption of this process could cause knock-on misassembly of the spiral and therefore aberrant knob structures. Although we were not able to visualise spiral structures at the base of mentulae, our observed phenotype could be explained by runaway lengthening of such a spiral structure (Figure 6). Indeed, the electron density we observed on tomograms is consistent with a high molecular weight structure as a form-giving scaffold for the formation of mentulae. Within the resolution limit of our study, KAHRP itself appears to line the inner leaflet of the membrane-bounded mentulae and is thus unlikely to itself be a major component of the electron dense core of the mentulae. The action of host actin is known to be required for the generation of knobs34; however, although we could successfully visualise actin below the erythrocyte plasma membrane, it appears to be largely excluded from mentulae and thus cannot be responsible for maintaining the form of these structures, a view supported by the lack of action of cyto-D on mentula structure. Similarly, enzymatic removal of the erythrocyte glycocalyx had no effect on mentulae, excluding a role for this in membrane shaping. Chelation of membrane cholesterol via treatment with MBCD did cause change in the morphology of mentulae, but not a complete reversion to normal knob structures. Removal of cholesterol from biological membranes has been observed to cause an increase in membrane stiffness35,36, and this may explain the morphological reversion upon MBCD treatment, with a stiffer membrane being more resistant to the pushing force within the mentulae. Alternatively, removal of cholesterol and subsequent breakdown of so-called lipid rafts may interfere with the higher order organisation of membrane-bound factors involved either directly in membrane curvature or required for coordination of proteins internal to the mentulae that generate force. Taken together, our data strongly implies that both knobs and mentulae contain a so far unknown component, likely proteinaceous, which is required for force generation, subsequent membrane curvature, and resulting morphology. Interruption of correct assembly of this factor leads to aberrations in knob formation, eventually leading to the appearance of mentulae. Our, and others’, data implicates KAHRP as having a role in this process, but it is unlikely to be the only structural protein involved.
In other systems, HSP40s, through their role as a regulator of HSP70 chaperone activity, have been shown to have a role in both assembly and disassembly of protein complexes37, and it is tempting to suggest that the phenotype we observe here is due to incorrect complex assembly. Alternatively, PFA66 may be required for the correct transport of accessory proteins required for complex formation such as those proposed in a recent study34, and thus play an indirect role in correct assembly of high-molecular weight complexes. In support of this hypothesis, PFA66 is known to associate with J-dots, highly mobile structures within the infected erythrocyte that are also known to contain a number of HSP70s13,15,16,20. It is also feasible that PFA66 is required for the disassembly of incorrectly folded or assembled knob protein complexes and that our knockout reveals so far unknown quality control mechanisms.
Knobs are required for correct presentation of the major virulence factor PfEMP1, and high affinity binding of such to endothelial receptors38. Although previous studies suggested that KAHRP and PfEMP1 formed a ‘precytoadherance complex’ at the Maurer’s clefts39, later evidence suggest strongly that PfEMP1 is incorporated only into knobs once they have been at least partly formed34,40. Erythrocytes infected with ΔPFA66 parasites showed a 63% reduction in knob/mentulae density and a 10-fold increase in the frequency of abnormal knob phenotype, and we observed an almost total lack of cytoadherence in iRBC infected with ΔPFA66 parasites. This data strongly supports the view that, even in the normal knobs present, less PfEMP1 was correctly presented and could take part in cytoadherence. Flow cytometry determined an almost 60% drop in cell surface recognition of the VAR2CSA variant of PfEMP1, although immunofluorescence suggests that both cell lines express similar amounts of this protein. The total loss of cytoadherence may be the consequence of several distinct factors: a) less total surface PfEMP1, b) fewer knobs with correctly loaded PfEMP1, and c) a significant number of aberrant knobs/mentulae. Moreover, we cannot exclude that PfEMP1, which needs to be correctly folded to bind specific receptors, does not assume the correct tertiary structure due to the lack of the necessary chaperone/co-chaperone system.
As previously mentioned, HSP40s generally act in concert with members of the HSP70 family12. The erythrocyte is known to contain significant amounts of residual human HSP70s41 and additionally a parasite-encoded HSP70, PfHSP70-X 15. We have previously demonstrated that a knockout of PfHSP70-X leads to a reduction in virulence characteristics, including cytoadherence42. Significantly, however, iRBCs infected with Δ70-x parasites were covered with normal knob structures at a density comparable to that of the wild type42. Hence, although both knockouts showed defects in virulence functions, the mutant phenotype of iRBCs infected with ΔPFA66 is distinct and significantly more dramatic than that in iRBCs infected with Δ70-x parasites. PFA66 has been reported to undergo a functional interaction with PfHSP70-X, HsHSP70, and HsHSC7043,44. Considering the difference in phenotype between Δ70-x and ΔPFA66, here we must conclude that, if a HSP70 is involved, it is more likely to be of human rather than parasite origin. We cannot formally exclude that PFA66 functionally interacts with both human and parasite HSP70 homologues and that the phenotype we observe is a combination of the effect on both chaperones, but the balance of probabilities suggests that morphological abnormalities observed in this current study are largely due to an interruption of functional PFA66-HsHSP70/HSC70 interactions.
The J-domain of HSP40s contains a characteristic HPD motif that is required both for binding and ATPase activation of partner HSP70s37. As the unlikely possibility existed that our observed phenotype was due to a HSP70-independent function of PFA66, we carried out complementation analysis with a full-length copy of PFA66 expressed from an episome, or a copy containing a H111Q mutation that renders the J-domain inactive. As only the wild type protein was able to complement the mutant knob phenotype, we conclude that the effects we observed upon PFA66 truncation are due to either a dominant-negative effect of the truncated protein on proper functioning of the HsHSP70 chaperone system within the host erythrocyte, or an effect due to lack of co-chaperone activity via deletion of an essential functional domain (SBD) of the HSP40. Either way the results support HsHSP70 involvement. A potential role for residual human HSP70 in host cell modification and parasite virulence has been suggested for almost 20 years45, but to our knowledge our current study is the first to provide strong experimental evidence implicating human HSP70s in these processes.
To conclude, in this study we show data suggesting that correct biogenesis of knobs in malaria-infected erythrocytes is a complex process necessitating a number of proteins, the molecular identity of some of which remains enigmatic. Our data suggests that KAHRP, while obviously required for knob generation, may not directly provide a scaffold for knob structure. More importantly, our data also reveals that residual human HSP70 within the infected erythrocyte is involved in parasite-driven host cell modification processes. To our knowledge, this is the first time a host cell protein has been implicated in parasite virulence, and this observation opens up exciting new avenues for the development of new anti-malarials.
Materials & Methods
Vector construction
The ~1kbp PFA66 targeting region was amplified using the primers PFA_NotI_F and PFA_MluI_R and cloned into pSLITGD using NotI-HF and MluI (NEB)17 (kind gift of Tobias Spielmann). The complementation plasmid PFA::HA was generated by excising the PFA0660w coding sequence and promoter from the plasmid pAD-A660-GFP13 with the restriction enzymes NotI-HF and BssHII and cloning them into pARL-PFF1415c-3xHA (BSD, a kind gift of Sarah Charnaud). The resulting plasmid was subsequently used to generate the QPD::HA plasmid via QuickChange PCR using PFA_Quick_QPD_F and PFA_Quick_QPD_R primers. Upon verification of the QPD mutation, the insert consisting of the PFA0660w promoter and coding sequence was re-cloned into the same vector to avoid mutation due to the PCR step. The KAHRP::mCherry plasmid was generated by amplifying mCherry with the primers mCherry_AvrII_F and mCherry_XmaI_R and cloning them into a pre-existing plasmid pARL2_KAHRP containing the native KAHRP promoter and KAHRP coding sequence (kind gift of Cecilia Sanchez). All primers are listed in Supplementary Table 2A.
Cell culture methods
P. falciparum parasites were cultured at 37°C with 90% N2, 5% CO2, and 5% O2 according to established methods46. Parasites were maintained at a haematocrit of ~5% in A+ or O+ human blood obtained from the blood banks in Marburg and Heidelberg, respectively, and maintained with RPMI1640 (Gibco) containing 200 μM hypoxanthine, 160 μM neomycin (Sigma Aldrich) and 10% human plasma. Parasitaemias were evaluated from smears prepared from the blood cultures, which were fixed in 100% MeOH and stained with ddH2O/10% Giemsa solution (Merck). Parasites were transfected with 150 μg of plasmid and treated with 2.5nM WR99210 (HS lines) or 12μg/ml blasticidin (Invivo Gen). Transfectants were propagated in fresh O- blood using RPMI1640 (Gibco) with 5% human plasma, 5% albumax II (Invitrogen) and other additives as above until parasites re-appeared47. Selection-linked integration was performed according to Birnbaum et al.17. Briefly, following reappearance after the initial transfection, parasites were treated with 400 μg/ml G418 (Thermo Fisher Scientific) until resistant parasites were observed. Parasites were synchronized before experiments using sorbitol-induced lysis48. For this mixed-culture parasites were incubated in 5% sorbitol for 10 min, washed with parasite culture medium and re-cultivated. Routine selection for CSA-binding parasites was performed according to standard protocols19. Late-stage parasites were enriched via gelatine flotation for 1 hr / 37 °C 49. Subsequently parasites were resuspended in cytoadhesion media (pH 7.2, prepared from RPMI1640 powder (Life technologies)) and incubated in cell culture flasks pre-treated with CSA (PBS pH 7.2/1 mg/ml CSA overnight / 16 °C and blocked with PBS pH 7.2 / 1% BSA) for 1 hr / 37 °C. After careful washing with cytoadhesion media, the remaining bound parasites were washed off and re-seeded.
Chemical or enzymatic treatment of iRBCs
For these experiments, magnetically purified iRBCs (~1 x 107 per condition) were used. Cytochalasin-D treatment was carried out with RPMI1640 / 10μm cyto-D for 10 min at RT, incubation with RPMI1640/10 mM MBCD was performed for 20 min at 37 °C, RPMI1640/ 30mU neuraminidase or RPMI1640/30U hyaluronidase treatments were performed for 1 hr at 37 °C. Following treatment, samples were processed for SEM.
MACS purification
For some protocols, late-stage parasites were magnetically purified using a VARIOMACS with a CS-column50. Briefly ~1 ml packed erythrocytes (~10% parasitaemia) were applied to a CS column, washed with PBS / 3% BSA and finally eluted into PBS.
Microscopy methods
Live cell imaging was performed on DAPI-stained (1 ng/ml) parasites using a Zeiss Axio-Observer microscope and AxioVision software. For immunofluorescence assays, parasites were fixed on microscopy slides using 90% acetone / 10% MeOH for 5 min / −20 °C. Cells were then blocked using PBS / 3% BSA for 1 hr / RT and incubated in a humid chamber overnight with the primary antibody diluted in blocking buffer (for antibodies see Supplementary Table 2B). On the next day, PBS-washed slides were treated with the secondary antibody, diluted 1:2,000 in blocking buffer for 2 hr / RT, subsequently washed again, DAPI stained (0.1 ng/ml in PBS), and imaged using a Zeiss Axio-Observer microscope and AxioVision software. RSTED imaging was carried out as recently reported in great detail51. Wheat germ agglutinin Alexa Fluor® 488 conjugate (Thermo Fisher) was used according to the supplier’s specification. Phallodin-Atto 647N (Thermo Fisher) was diluted 1:500 in PBS and incubated for 30 min / RT to stain the actin cytoskeleton. RFP booster ATTO594 was used 1:200/2hr/RT to enhance RFP fluorescence. IMSpector imaging software (Abberior Instruments GmbH) was used for image capture and deconvolution of STED images, and AxioVision software was used for other acquisitions. Images were processed using ImageJ. Brightness and contrast was adjusted to reduce background and enhance visibility. No gamma adjustments were applied to any images, and all data is presented in accordance with the recommendations of Rossner and Yamada52.
Protein-based methods
Protein extracts were prepared from 1 x 108 MACS-purified iRBCs. These were resuspended in PBS and boiled in Laemmli loading buffer for 10 min at 99 °C. Soluble fractions were separated via centrifugation (4 °C, 35,000 g) and an equivalent of 1 x 107 parasites loaded onto each well of 12% acrylamide gels. Equinatoxin (EQT) treatment and fractionation of MACS-purified iRBCs was carried out as described by Külzer et al.22 but using 4 haemolytic units of EQT at RT for 6 min. Western blot / immunodetection was carried out via semi-dry blotting, blocking in 5% milk powder (1 hr / RT), incubation with primary (overnight / 4 °C), washing three times with PBS, incubation with the secondary (2 hr / RT) antibody in blocking buffer, washing three times with PBS, and visualization via x-ray films. Antibody sources and dilutions can be found in Supplementary Table 2B.
Membrane shearing
For investigation of the internal structure of the RBC cytoskeleton membrane shearing was employed according to established protocols34,53. Briefly, a (3-aminopropyl)triethoxysilane-treated ibidi dish was incubated with 150 μl PBS / 1 mM with Bis(sulfosuccimidyl)suberate for 30 min / RT, washed with PBS, and incubated with 150 μl ddH2O / 0.1 mg/ml erythroagglutinating phytohaemagglutinin for 2 hr / RT. Dishes were rinsed three times with PBS and quenched using PBS / 0.1 M glycine for 15 min / RT. Approximately 1 x 107 MACS-purified iRBCs were added and incubated for 3-4 hr, washed, and sheared using 5P8-10 buffer (5 mM Na2HPO4 / NaH2PO4, 10 mM NaCl, PH 8), while angling the dish at 20°. Samples were then blocked using 150 μl of PBS / 1% BSA, treated with the primary α-KAHRP antibody overnight / 4 °C in blocking buffer, washed three times, incubated with the secondary α-mouseATTO549 for 1 hr / RT in blocking buffer, and finally washed three times before imaging via rSTED.
Sorbitol lysis
Assessment of NPP activity was carried out according to Baumeister et al.23. For each measurement, 40 μl of 2% trophozoite culture was resuspended in 150 μl lysis buffer (290 mM sorbitol, 5 mM HEPES, pH 7.4) and incubated for 30 min / 37 °C. Remaining RBCs were then pelleted at 1,600 g / 2 min, and the absorbance of the resulting supernatant was measured at OD570nm. Samples were then compared to a total lysis control, which was generated in parallel using ddH2O instead of lysis buffer.
Flow cytometry
Infected erythrocytes were fixed for 24 hr at 4 °C using PBS/4% paraformaldehyde/0.0075% glutaraldehyde and stained with DAPI (1 ng/ml) prior to analysis with a BD Canto. In the growth experiments, both cell lines were diluted after every growth cycle with the same factor in order to support parasite growth. Both parasite cell lines were seeded with the same parasitaemia and diluted after every cycle to avoid ‘crashing’ the culture. Parasitaemias were measured before and after every dilution by staining of iRBCs with DAPI and flow cytometry. For staining VAR2CSA on the RBC surface, live parasites were incubated with VAR2CSA antiserum (11P, rabbit, a kind gift of Benoit Gamain) and α-rabbit-Cy3 for 30 min each and then processed for flow analysis as detailed above.
Cytoadherence
IRBC cytoadhesion to immobilized CSA was investigated using MACS-purified late-stage parasites18. Parasites were applied in cytoadhesion media (pH 7.2, made from RPMI1640 powder (Life technologies) to pre-treated spots (PBS pH 7.2/1 mg/ml CSA overnight at 16 °C, blocked with PBS pH 7.2/1% BSA for 1 hr at RT), and then washed with PBS on a Petri dish. After incubation for 1 hr at RT, non-bound parasites were washed away using cytoadhesion medium. Parasites were then fixed using PBS/2% glutaraldehyde for 2 hr at RT and stained with PBS/10% Giemsa for 10 min at RT prior to imaging using a Zeiss Axio Observer microscope and counting with Ilastik54 and ImageJ software.
Electron microscopy
For scanning electron microscopy, purified parasites were fixed using PBS /1% glutaraldehyde for at least 30 min at RT. After washing, parasites were bound to coverslips (pre-treated with 0.1% polylysine for 15 min at RT), washed again, and dehydrated in acetone gradients (ddH2O, 25% Ac, 50% Ac, 75% Ac, 100% Ac, 10 min each) followed by critical point dehydration and coating with 5 nm Pd-gold. Cells were imaged using a Zeiss Leo 1530 electron microscope (SE2 detector, ~12,000 x magnification)55. For transmission electron microscopy, parasites were fixed in 100 mM Ca-cacodylate/4% paraformaldehyde/ 2% glutaraldehyde, embedded in Spurr and cut into ~70 nm sections. Some samples were fixed using 100 mM Ca-cacodylate/4% paraformaldehyde/ 0.1% glutaraldehyde and treated according to the Tokuyasu protocol for immunogold labelling of KAHRP using an α-KAHRP (rabbit) antibody and a secondary goat α-rabbit-gold conjugated antibody56. Some of the EM sections were used without post-contrasting, while some were post-contrasted using 3% uranyl acetate and ddH2O/0.15 M Na-citrate/0.08 M Pb(NO3)2 / 0.16 M NaOH for 2 min. Imaging was performed using a Jeol 1400 microscope operating at 80kV. For electron tomography ~350 nm thick sections were used and examined in a TECNAI F30, 300kV FEG, FEI electron microscope (EMBL Heidelberg). The resulting tomograms were processed using IMOD, ETOMO image/ volume processing software package and the Amira, volume visualisation software.
Statistics
Statistics were calculated in prism or Excel using unpaired, two-tailed t-tests. p > 0.05 = non-significant (ns); *p < 0.05; **p < 0.01; ***p < 0.00. Figures show mean and standard deviation.
ImageJ Macro
The ImageJ/Fiji57 macro computes the local maxima of each object on the smooth probability map (PM) images generated by ilastik pixel classification. The ilastik 54 pixel classification workflow is used to reduce the background in the images and enhance the foreground pixels. To segment each object in the probability map images the local maxima is used as a seed for the 3D watershed plugin. The approach allows to separate close objects and creates masks that are used to measure size and intensity on the raw images. The macro and the instructions on how to use it can be found at: https://github.com/cberri/2D_AutomatedObjectsDetection_ImageJ-Fiji
Supplementary Video 1. 3D reconstruction and surface render of mentulae depicted in Figures 2E, F. Red, erythrocyte plasma membrane; blue, electron dense material.
Supplementary Video 2A. Z-stack of erythrocytes infected with CS2[KAHRP::mCherry] in mCherry channel.
Supplementary Video 2B. Z-stack of erythrocytes infected with CS2[KAHRP::mCherry] in mCherry channel.
Supplementary Video 2C. Z-stack of erythrocytes infected with ΔPFA[KAHRP::mCherry] in mCherry channel.
Supplementary Video 2D. Z-stack of erythrocytes infected with ΔPFA[KAHRP::mCherry] in mCherry channel.
Acknowledgements
We wish to thank the blood banks of the University Hospitals in Giessen and Marburg for providing blood. Further we would like to thank the EMCF at the University of Heidelberg (in particular Stefan Hillmer), the EMCF of EMBL Heidelberg (Yannick Schwab and Martin Schorb), the FACS core facility at the ZMBH (Monika Langlotz), as well as the Infectious Diseases Imaging Platform (Vibor Laketa) and Marie Freudenberg. We thank Tim Bostick for proofreading. This work was supported by DFG grant PR1099/8-1 to JMP.
- Abbreviations
- SLI
- selection-linked integration
- (i)RBC
- (infected) red blood cell
- PV(M)
- parasitophorous vacuole (membrane)
- RBCM
- red blood cell membrane
- PPM
- parasite plasma membrane
- BSD
- blasticidin
- NEO
- neomycin
- KAHRP
- knob-associated histidine-rich protein
- SEM
- scanning electron microscopy
- TEM
- transmission electron microscopy
- (r)STED
- (rescue) stimulated emission depletion
- NPP
- new permeability pathways
- IFA
- immunofluorescence assay
- EQT
- equinatoxin
- CSA
- chondroitin-sulphate-A
- MC
- Maurer’s clefts