Heterochromatin dynamics during the initial stages of sexual development in Plasmodium falciparum

Nhim, Sandra; Tintó-Font, Elisabet; Casas-Vila, Núria; Michel-Todó, Lucas; Cortés, Alfred

doi:10.1038/s41598-024-73981-w

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Published: 05 October 2024

Heterochromatin dynamics during the initial stages of sexual development in Plasmodium falciparum

Sandra Nhim^1,2,
Elisabet Tintó-Font^1,2,
Núria Casas-Vila^1,2,
Lucas Michel-Todó^1,2^nAff4 &
…
Alfred Cortés^1,2,3

Scientific Reports volume 14, Article number: 23180 (2024) Cite this article

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Abstract

Asexual replication of Plasmodium falciparum in the human blood results in exponential parasite growth and causes all clinical symptoms of malaria. However, at each round of the replicative cycle, some parasites convert into sexual precursors called gametocytes, which develop through different stages until they become infective to mosquito vectors. The genome-wide distribution of heterochromatin, a type of chromatin generally refractory to gene expression, is identical at all asexual blood stages, but is altered in stage II/III and more mature gametocytes. However, it is not known if these changes occur concomitantly with sexual conversion or at a later time during gametocyte development. Using a transgenic line in which massive sexual conversion can be conditionally induced, we show that the genome-wide distribution of heterochromatin at the initial stages of sexual development (i.e., sexual rings and stage I gametocytes) is almost identical to asexual blood stages, and major changes do not occur until stage II/III. However, we found that at loci with heterochromatin alterations, transcriptional changes associated with sexual development typically precede, rather than follow, changes in heterochromatin occupancy.

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Introduction

In eukaryotic organisms, chromatin plays fundamental roles in the regulation of gene expression and other nuclear processes. A type of chromatin termed heterochromatin mediates transcriptional repression and also plays important roles in centromere and telomere function and in repressing transposable elements, thus securing chromosome stability^1,2,3,4. Conserved features of heterochromatin across eukaryotes include its ability to spread into adjacent regions and epigenetic inheritance of its distribution during cell division. Regions that are found as heterochromatin in some cells and as euchromatin in others, which often include protein-coding genes, are referred to as facultative heterochromatin, whereas regions that are always found as heterochromatin, such as subtelomeric and pericentromeric tandem repeats regions, are called constitutive heterochromatin. In the majority of eukaryotes, the main molecular determinants of heterochromatin are the histone modifications di- or tri-methylation of histone H3 lysine 9 (H3K9me2 or H3K9me3) or tri-methylation of histone H3 lysine 27 (H3K27me3). H3K27me3 is typically associated with dynamic facultative heterochromatin and regulates transient gene silencing during development and cell type specification in multicellular organisms. In contrast, H3K9me3 is the hallmark of constitutive heterochromatin, although it can also play important roles in the dynamic regulation of gene expression by forming facultative heterochromatin^1,2,3,4.

In Plasmodium falciparum, the parasite responsible for the most severe forms of human malaria, H3K9me3 is dynamic and frequently associated with facultative heterochromatin, whereas H3K27me3 has been identified only at specific stages of sexual development⁵ and its function still remains unclear. H3K9me3 has been extensively studied, as it plays fundamental roles in parasite biology. This mark, together with the conserved heterochromatin protein 1 (HP1) that binds to it, occupies subtelomeric repeats and clonally variant gene (CVG) loci, which are located in subtelomeric regions and a few chromosome internal islands^6,7,8,9,10. The distribution of H3K9me3 and HP1 is essentially identical^6,7,8,9. Presence of H3K9me3 and HP1 at the upstream regulatory regions of CVGs, where they form facultative heterochromatin, is associated with transcriptional silencing, whereas presence of acetylated H3K9 (H3K9ac) at these positions is associated with their active state⁹. The distribution of H3K9me3-based heterochromatin at CVGs is clonally transmitted from one generation of blood stage asexual parasites to the next, acting as a truly epigenetic mark¹¹. However, infrequent transitions between the heterochromatic and euchromatic states result in transcriptional switches^12,13,14.

The P. falciparum genome contains more than 500 CVGs, which participate in numerous host-parasite interactions, including antigenic variation, solute transport, erythrocyte invasion, erythrocyte remodelling and sexual conversion^12,13,14,15. Many of these genes belong to multigene families in which there is redundancy, as different genes of the same family often encode similar proteins that participate in the same process, albeit with antigenic or functional differences. Therefore, switches in the expression of CVGs ultimately result in antigenic or phenotypic variation. In general, epigenetic regulation of malarial CVGs plays an adaptive role: functional or antigenic diversity within parasite populations provides the grounds for dynamic natural selection when the conditions of the environment change. This is considered a bet-hedging adaptive strategy¹⁵.

The P. falciparum life cycle involves multiple well-differentiated stages in humans and in Anopheles spp. mosquito vectors. In the human blood, parasites undergo the asexual intraerythrocytic development cycle (IDC), which lasts ~ 48 h and involves the merozoite, ring, trophozoite and schizont stages. Repeated rounds of the IDC result in exponential parasite growth, which is associated with all clinical symptoms of malaria. However, at each round of the IDC, a small fraction of the parasites abandons asexual growth and converts into sexual precursors called gametocytes, in a process called sexual conversion¹⁶. Gametocytes are the only form of the parasite that, once mature, can infect mosquitoes. In the mosquito, after mating of male and female gametocytes and several additional stage transitions, parasites become salivary gland sporozoites, which can infect a new human host during a mosquito bite. Sporozoites invade hepatocytes, where they multiply until they are released to the blood stream to start a new blood infection, closing the cycle¹⁷.

The consolidated model for sexual conversion postulates that PfAP2-G, an ApiAP2 transcription factor, is the master regulator of the process^18,19,20,21. During the IDC, the pfap2-g locus is in a heterochromatic state that maintains the gene silenced. Expression of the GDV1 protein mediates heterochromatin depletion at the pfap2-g locus²², which results in expression of PfAP2-G and sexual conversion. The heterochromatin-based regulation of PfAP2-G^18,23 illustrates the cross-talk between transcription factor networks and epigenetic regulation of gene expression. Heterochromatin may also play a role in the regulation of gdv1, as heterochromatin occupancy at this locus correlates with basal sexual conversion rates⁹. PfAP2-G regulates the expression of several early gametocyte genes, which drive sexual differentiation²¹. Depending on the time of activation of PfAP2-G expression during the IDC, parasites can convert directly into gametocytes via the same cycle conversion (SCC) pathway, or go through one additional cycle of multiplication as sexually-committed forms before converting via the next cycle conversion (NCC) pathway²⁴. Regardless of the conversion pathway used, a cascade of transcription factors activation ensues^25,26 and drives development through the sexual ring stage and then stage I to V gametocytes, in a process that lasts ~ 10 days until male or female gametocytes are mature and ready to infect a mosquito.

Heterochromatin distribution has been compared between clonal parasite lines that differ in the expression of specific CVGs, which revealed alternative heterochromatin patterns associated with the active or silenced states of these genes^{9,12,27,28,29}. However, heterochromatin distribution differences associated with life cycle progression have also been reported. While several studies found that the distribution of heterochromatin at specific P. falciparum CVG loci^27,28,29 or at a genome-wide level⁷ is almost identical between different stages of the IDC, the genome-wide distribution of heterochromatin is different in gametocytes^7,30 or mosquito stages (oocysts and sporozoites)^31,32. The most prominent heterochromatin alterations observed during these transmission stages were expansion of heterochromatic domains at some subtelomeric regions and opening of heterochromatin at a small number of specific loci. Major changes in heterochromatin distribution in transmission stages, involving subtelomeric heterochromatin expansions, were also observed in the simian malaria parasite P. cynomolgi³³, but not in the murine malaria parasite P. berghei³⁴. Another recent study reported heterochromatin differences at some specific loci between male and female P. falciparum gametocytes³⁵. However, none of these studies included early sexual stages such as sexual rings or stage I gametocytes. Therefore, it is currently not known if changes in heterochromatin distribution during sexual development occur concomitantly with sexual commitment, at the initial stages of sexual development or later on during sexual development.

Here we took advantage of a recently developed inducible parasite line that enables controlled massive sexual conversion²⁰ to investigate heterochromatin distribution in early sexual stages. To assess the impact of heterochromatin redistribution during sexual development on gene expression, we characterised the relative temporal dynamics of changes in heterochromatin and in gene expression. We also used existing transcriptomic and epigenomic datasets to develop an integrated model of heterochromatin dynamics in transmission stages.

Results

Heterochromatin distribution is almost identical between asexual blood stages, sexual rings and stage I gametocytes

To characterise the distribution of heterochromatin at the initial stages of P. falciparum gametocyte development, we used the transgenic line E5ind, in which synchronous sexual conversion of the vast majority of parasites can be conditionally induced²⁰. In brief, addition of rapamycin to E5ind cultures at the trophozoite stage results in activation of the expression of PfAP2-G and sexual conversion via the NCC pathway²⁴ in ~ 90% of the parasites, such that they develop into sexually committed schizonts and, after reinvasion, into sexual rings and subsequent stages of sexual development (stage I to V gametocytes)²⁰.

Using the E5ind line, we performed H3K9me3 chromatin immunoprecipitation followed by sequencing (ChIP-seq) for sexual rings, stage I and stage II/III gametocytes, and also for asexual rings prepared in parallel without adding rapamycin (Fig. 1a and Supplementary Fig. S1). Since heterochromatin distribution remains stable throughout the full IDC⁷, the asexual ring stage is representative of all other IDC stages. Visual inspection of the general distribution of heterochromatin in two independent biological replicates did not reveal any major differences between asexual rings, sexual rings and stage I gametocytes, whereas in stage II/III gametocytes expansion of heterochromatin was apparent at several subtelomeric regions (Fig. 1b). Pearson correlation analysis confirmed a very high level of heterochromatin distribution similarity in pairwise comparisons between different stages (r > 0.93 in all comparisons), with the two replicates of stage II/III gametocytes forming a separate cluster (Supplementary Fig. S2a). Consistently, the proportion of the genome covered by a H3K9me3 peak (according to MACS2 peak calling) was slightly higher in stage II/III gametocytes than at the other stages (Supplementary Fig. S2b). These results indicate that heterochromatin distribution is almost identical between asexual blood stages and sexual rings or stage I gametocytes, and that the previously reported changes in heterochromatin distribution during sexual development^7,30 do not occur until at least the stage II of gametocyte development.

Subtelomeric heterochromatin expansions occurring from gametocyte stage II/III are similar among parasite lines of different genetic backgrounds

Since the most prominent heterochromatin distribution change during sexual development was expansion of heterochromatin at several subtelomeric regions, we developed an in-house pipeline to measure the length of the region covered by heterochromatin at each of the 28 subtelomeric regions and also at seven internal heterochromatin islands. The method was based on MACS2 peak calling and joining peaks within the same subtelomeric regions or internal island (Supplementary Fig. S3a). This analysis confirmed that the size of all heterochromatic regions was almost identical between asexual parasites, sexual rings and stage I gametocytes, whereas marked heterochromatin expansions (relative to asexual parasites) occurred in several subtelomeric regions in stage II/III gametocytes (Fig. 2a and Supplementary Fig. S3b). Different chromosome ends showed highly variable degrees of heterochromatin expansion, ranging from 0 to > 60 Kb. Heterochromatin islands remained invariant across all stages (Supplementary Fig. S4).

Next, we compared the size of subtelomeric or internal heterochromatin domains between our samples (E5ind line, of 3D7 genetic background) and published heterochromatin (H3K9me3 or HP1) ChIP-seq data from stage II/III or IV/V gametocytes of the Pf2004 genetic background⁷ (a culture-adapted Ghanian isolate), and sporozoites from a field isolate from Burkina Faso³¹. Overall, similar heterochromatin expansions (relative to asexual parasites), affecting the same subtelomeric regions and with similar limits, were observed between our E5ind stage II/III gametocytes and Pf2004 gametocytes⁷ (Fig. 2a-b and Supplementary Fig. S4-7). The few different expansions between E5ind and Pf2004 could be explained by differences in heterochromatin distribution between the two lines that were already present in asexual parasites, as a consequence of regular epigenetic variation^9,15 (i.e., some genes are silenced and heterochromatic in asexual parasites in one parasite line and active and euchromatic in the other). However, in many of the regions where the expansions occurred, H3K9me3 coverage was low compared with other heterochromatic regions, mainly in our E5ind stage II/III gametocytes (Fig. 1b, 2b and Supplementary Fig. S3a). This is suggestive of population heterogeneity, such that heterochromatin had already expanded in older individual gametocytes but not yet in the younger ones. Of note, heterochromatin distribution in sporozoites³¹ was generally similar to stage II/III or IV/V gametocytes (Fig. 2a–b and Supplementary Fig. S5b), although some additional expansions were observed. Since ChIP-seq data from asexual parasites of the same genetic background as the sporozoites sample was not available, changes in heterochromatin distribution in sporozoites had to be determined relative to E5ind asexual parasites. Therefore, some of the differences in heterochromatin expansions between gametocytes and sporozoites may be attributable to the different genetic backgrounds.

In addition to subtelomeric heterochromatin expansions, an apparent retraction of heterochromatin was observed at the distal end of chromosome 12 in E5ind stage II/III gametocytes and at the distal end of chromosome 14 in sporozoites (Fig. 2a). This could be explained by small changes in the very low level heterochromatin coverage at a specific locus (the clonally variant geco gene³⁶, which was fully heterochromatic at all stages in the Pf2004 line) or by loss of heterochromatin in two genes encoding the proteins CRMP4 and Ccp1 that play important roles in mosquito stages^37,38, respectively (Supplementary Fig. S6). In addition to the changes in the limits of specific subtelomeric heterochromatin domains, there were changes in heterochromatin distribution within some of the domains. In some subtelomeric regions, there was an expansion of heterochromatin that did not affect the estimated limit of the domain, involving loci that were euchromatic in asexual parasites (Fig. 2c).

Together, these results show that during sexual development there is expansion of heterochromatin at specific subtelomeric regions, starting at the gametocyte stage II or III. The heterochromatin expansions observed were similar between parasites of different genetic background, indicating that this is a conserved, controlled process intrinsic to P. falciparum sexual development.

Distribution of changes in heterochromatin coverage during sexual development

To analyse in more detail the changes in heterochromatin distribution, regardless of whether or not they affect the size and limits of large heterochromatin domains, we modified a previously described custom differential peak calling method⁹ to identify regions with differential H3K9me3 coverage between different stages. Comparison of E5ind asexual stages and stage II/III gametocytes revealed 139 differential coverage regions, of which 33.8% were localised in subtelomeric repeats regions devoid of protein coding genes and containing mainly TARE repeats³⁹ (Fig. 3a and Supplementary Table S1). Heterochromatin coverage in these subtelomeric repeats was generally higher in sexual compared to asexual parasites. Of note, 21 of the previously reported 22 subtelomeric lncRNA-TARE transcripts⁴⁰ intersected with differential coverage regions (Supplementary Fig. S8 and Supplementary Table S1).

A large proportion (49.6%) of the 139 differential peaks overlapped with the -1,000 to + 500 bp (from the ATG) region of a gene, and 14.4% of the peaks overlapped only with other parts of a gene (including the 3’UTR, when annotated). Lastly, three of the differential peaks (2.2%) occurred in intergenic regions that did not overlap with any of the annotated genetic elements mentioned above (“Others”) (Fig. 3a).

In contrast to the 139 differential H3K9me3 coverage regions identified when comparing asexual rings with stage II/III gametocytes, only 21 and two differential coverage regions were identified between asexual rings and sexual rings or between asexual rings and stage I gametocytes, respectively (Supplementary Table S1). In the case of sexual rings, the majority (76%) of the differential coverage regions overlapped with the -1,000 to + 500 bp region of a gene (Supplementary Fig S9a), whereas the two regions detected in stage I gametocytes were intergenic.

Genes with differential heterochromatin coverage at putative regulatory regions between asexual and sexual parasites

Given that heterochromatin changes at proximal upstream regions and the beginning of the coding sequence show the strongest association with gene expression⁹, we analysed in detail the differential heterochromatin coverage regions overlapping with the -1,000 to + 500 bp (from the ATG) region of a gene. In the comparison between asexual rings and stage II/III gametocytes, 107 genes were identified in which this region overlapped with a differential H3K9me3 peak and had a coverage difference higher than twofold (Fig. 3b and Supplementary Table S2). These will be referred to as ‘differential coverage genes’. For these genes, H3K9me3 coverage in the -1,000 to + 500 bp region was almost identical between E5ind asexual rings, sexual rings and stage I gametocytes, and markedly different only in stage II/III gametocytes. Heterochromatin changes at these genes in stage II/III gametocytes were similar to changes in stage II/III or IV/V gametocytes of the Pf2004 line⁷ (Fig. 3b). Indeed, there was a large overlap between the lists of differential heterochromatin coverage genes in E5ind stage II/III and Pf2004 gametocytes (Supplementary Table S3).

101 of the E5ind stage II/III differential coverage genes showed an increase in heterochromatin occupancy in stage II/III gametocytes compared to asexual parasites, reflecting mainly the previously described subtelomeric heterochromatin expansions. The majority of these 101 genes were not gametocyte markers and about half were previously described as CVGs⁴¹, with a predominance of genes encoding exported proteins involved in erythrocyte remodelling and PfEMP1 export, consistent with previous reports⁷ (Fig. 3b and Supplementary Table S2). However, in the remaining six differential coverage genes there was a reduction of heterochromatin coverage. For these genes, reduced occupancy was also observed only in stage II/III gametocytes. These genes included a cluster of early gametocyte markers located in the distal subtelomeric region of chromosome 14, including pfg14-744 and pfg14-748⁴², and two neighbour genes in chromosome 6 (Fig. 3b and Supplementary Table S2). A previous study reported that heterochromatin depletion in maturing gametocytes at the genes in the subtelomeric region of chromosome 14 occurs only in female gametocytes³⁵.

In contrast to the many differential coverage genes in stage II/III gametocytes (relative to asexual rings), an analogous analysis for sexual rings and stage I gametocytes revealed only seven and zero differential coverage genes, respectively (Supplementary Fig. S9b-c and Supplementary Table S4). In all seven differential coverage genes in sexual rings, there was a decrease in coverage of very low magnitude. Furthermore, the decrease was no longer observed in stage I or II/III gametocytes, suggesting that differential coverage in these few genes was likely explained by technical reasons.

To investigate the transcriptional changes during sexual development associated with the changes in heterochromatin occupancy, we used published gene expression data for the E5ind line²⁰. This analysis included the sexually-committed schizont, sexual ring and stage I gametocyte stages and their asexual counterparts (asexual schizont, asexual ring and trophozoite). In general, there were lower transcript levels in sexual stages (compared with the asexual counterparts) for the genes that gained heterochromatin during sexual development, and higher transcript levels for genes that lost heterochromatin. However, for many of the genes, transcript level changes already occurred in stage I gametocytes or even earlier sexual stages, in contrast to heterochromatin changes that were not observed until stage II/III gametocytes (Fig. 3b).

For the same 107 genes, we analysed changes in expression in a previously published transcriptomic dataset⁴³ of full gametocyte development that used parasites with a wild type pfap2-g locus. This analysis confirmed that the majority of genes with more heterochromatin in gametocytes were downregulated during gametocyte development and genes with less heterochromatin were upregulated (Supplementary Fig. S10 and Supplementary Table S2).

Transcriptional changes during sexual development may precede heterochromatin changes

To explore in more detail the relative timing of transcriptional and heterochromatin changes, we compared the temporal dynamics of heterochromatin coverage and transcript levels during sexual development for selected representative genes (using the same data as in Fig. 3b). For these genes, changes in heterochromatin did not occur until stage II/III gametocytes, whereas changes in transcript levels²⁰ occurred at an earlier stage (Fig. 3c-d). In the genes kahrp, clag3.1, pfd80 or pf332, increased H3K9me3 coverage was not observed until stage II/III gametocytes, whereas reduced transcript levels (compared with their asexual counterparts) were already observed at the sexually-committed schizont (clag3.1), sexual ring (kahrp) or stage I gametocyte (pfd80 and pf332) stages (Fig. 3c). Likewise, reduced H3K9me3 coverage was observed in pfg14-744 and a neighbour phist gene (PF3D7_1477400) only in stage II/III gametocytes, but increased transcript levels were already observed in stage I gametocytes (Fig. 3d). All six genes with reduced heterochromatin occupancy during sexual development are direct targets of PfAP2-G²¹. Comparison with published PfAP2-G binding data (ChIP-seq)²¹ revealed that this transcription factor already binds to the putative regulatory region of these genes in stage I gametocytes, at positions that overlap with the regions where heterochromatin occupancy is reduced in stage II/III gametocytes (Fig. 3e). Therefore, the temporal dynamics of PfAP2-G binding appears to correlate with changes in gene expression and precede changes in heterochromatin coverage.

To confirm these results, we performed RT-qPCR and H3K9me3 ChIP-qPCR analysis from time-course experiments that included five time points from the sexual ring to stage III gametocyte stages, with a non-induced asexual rings culture as a control (Fig. 4a). In addition to genes in Fig. 3c–d, in this analysis we also included pfg14-748, PF3D7_0601600 (putative tetratricopeptide repeat protein), for which data from the previous microarray-based transcriptomic analysis²⁰ was not available, and geco, which did not pass the threshold for differential heterochromatin coverage but showed a peculiar pattern of early heterochromatin alterations (Supplementary Fig. S6a). For all genes analysed, the same trend (increase or decrease) in heterochromatin occupancy during sexual development as in the ChIP-seq analysis was observed. These experiments confirmed that changes in H3K9me3 coverage generally occurred from stage II or II/III onwards, whereas transcript levels started to increase or decrease one to two days earlier (Fig. 4b–c). In general, in genes with increased H3K9me3 coverage the change was observed from day 5 post-induction (stage II/III gametocytes), whereas in genes with reduced coverage it was observed from day 3 (stage II gametocytes). Changes in transcript levels during sexual development (silencing of genes with increased heterochromatin or activation of genes with reduced heterochromatin) were typically observed from day 2 (stage I gametocytes) or even earlier, confirming that these transcriptional changes precede, rather than follow, heterochromatin changes. For pf14-744 and pf14-748, high transcript levels occurred transiently only until stage II gametocytes, as previously reported^42,43.

Changes in heterochromatin distribution at specific CVG families

Many of the 107 stage II/III differential coverage genes belong to multigene CVG families. Different families of CVGs occupy a relatively conserved position within subtelomeric regions, such that large hypervariable families involved in antigenic variation such as var, rif, pfmc-2tm and stevor tend to be located closer to the telomeres, whereas other CVG families that confer functional plasticity to the parasites tend to be located in more telomere-distal positions within subtelomeric regions^44,45. Heterochromatin expansion during sexual development mainly affected the latter type of CVG families, in addition to genes that did not belong to gene families previously annotated as containing CVGs. The CVG family with the largest absolute number of genes showing altered heterochromatin occupancy during sexual development was phist (23 of 78 genes), involved in erythrocyte remodelling^46,47,48, followed by hyp (11 of 41) that comprises several families involved in the same process⁴⁶ and fikk (8 of 20), which encodes exported kinases also involved in erythrocyte remodelling^46,49 (Fig. 5, Supplementary Fig. S11 and S12a and Supplementary Table S5). Other CVG families with a high proportion (≥ 20%) of differential coverage genes were acyl-coA synthetases (acs), acyl-coA binding proteins (acbp), glycophorin binding proteins (gbp), clag, eba, surfin and hrp (Fig. 5, Supplementary Fig. S11 and S12a and Supplementary Table S5), involved in various processes that include solute transport, erythrocyte invasion and lipid metabolism, among others. As a control, a gene family encoding non-clonally variant essential genes, aminoacyl tRNA synthetases (ars), did not show changes in H3K9me3 coverage. Four large families of CVGs located in telomere-proximal positions within subtelomeres (var, rif, pfmc2-tm and stevor) showed very few or no heterochromatin changes (Supplementary Fig. S11 and S12b).

Some gametocyte markers are considered CVGs, as they show variant expression and/or carry heterochromatin marks in asexual parasites. By overlapping previously published lists of CVGs and gametocyte markers^9,41, we identified 27 genes that fall in this category. It is not known if, once parasites convert into sexual forms, the chromatin state of these genes in asexual parasites affects their expression in gametocytes. In four genes, including pfg14-744 and pfg14-748, H3K9me3 occupancy decreased in putative regulatory regions in stage II/III gametocytes (Supplementary Fig. S13 and Supplementary Table S6). This suggests that chromatin remodelling during sexual development secures that these genes are in an euchromatic state in sexually-developing parasites, regardless of their state in asexual parasites. In contrast, surfin 13.1 and surfin 4.1 became heterochromatic as a consequence of subtelomeric heterochromatin expansion in stage II/III gametocytes. Of note, surfin 13.1 has been identified as one of the earliest sexual conversion markers, with high expression already at the sexually-committed schizont stage^20,50 and reduced expression later during gametocyte development⁴³. This raises the intriguing possibility that heterochromatin expansion in stage II/III gametocytes may contribute to developmental silencing of this sexual marker when its product is no longer needed. In the other clonally variant gametocyte genes, including well-known early gametocyte markers (e.g., gexp5 and pfg27/25), H3K9me3 coverage remained invariant at all the stages analysed. Therefore, it remains possible that presence or absence of heterochromatin at these genes in asexual parasites may influence their expression during sexual development.

Exploratory analysis to characterise heterochromatin changes at the pfap2-g locus after sexual conversion

Our ChIP-seq analysis using the E5ind line does not inform about heterochromatin changes during sexual conversion at the locus encoding the master regulator PfAP2-G, because it has been edited in this transgenic line²⁰. The pfap2-g locus, which is heterochromatic in asexual parasites^18,23, is activated by GDV1-mediated heterochromatin displacement in parasites that commit to sexual development²², but the distribution of heterochromatin at the pfap2-g locus soon after sexual conversion has not been characterised.

We used the NF54-gexp02-Tom line, in which sexual parasites express a fluorescent marker from the sexual ring stage onwards⁵¹, to sort asexual and sexual rings using flow cytometry and compare their heterochromatin distribution (Supplementary Fig. S14a-c). ChIP-seq analysis of the sorted populations revealed reduced heterochromatin occupancy in sexual rings at the region upstream of pfap2-g, including the putative promoter, but not at the 5’UTR or coding sequence (Supplementary Fig. S14d). This result is consistent with previous observations of reduced heterochromatin at the pfap2-g locus in later gametocytes occurring specifically at the upstream region³⁵. Our analysis also revealed absence of heterochromatin coverage differences between sexual and asexual rings in genes with differential coverage in sexual rings or stage II/III gametocytes in the E5ind line (Supplementary Fig. S14e). This result supports the idea that general heterochromatin changes do not occur until stage II/III gametocytes and the small differences in sexual rings in E5ind were likely attributable to technical reasons. Of note, the analysis of the NF54-gexp02-Tom line was complicated by the very small amount of chromatin obtained from sorted parasites. The results of this exploratory analysis will require confirmation using chromatin analysis techniques more suitable for experiments with low input material, such as CUT&TAG or CUT&RUN^52,53.

Discussion

Our results show that, in parasites using the canonical NCC pathway, global heterochromatin changes do not occur at the initial stages of sexual development (i.e., sexual rings and stage I gametocytes) and are only observed from gametocyte stage II/III onwards. Therefore, global heterochromatin redistribution likely plays a role in intermediate or late steps of gametocyte development but not in sexual conversion. The regulatory role of heterochromatin during gametocyte development is additional to its well-established adaptive role in the dynamic regulation of CVGs expression in asexual blood stages^12,13,14.

Life cycle progression in malaria parasites is mainly regulated by a cascade of ApiAP2 transcriptional regulators, such that expression of different ApiAP2 factors at different stages of the life cycle results in different transcriptomes⁵⁴. For the development of gametocytes, the cascade involves PfAP2-G, PfAP2-G2, PfAP2-G5, AP2-FG and several others^18,25,26,55. As previously noted⁷, the majority of genes that are activated or silenced during gametocyte development do not acquire heterochromatin at any sexual or asexual stage. This indicates that transcription factor networks are the main mechanism governing changes in the transcriptome during gametocyte development, whereas heterochromatin alterations are a complementary regulatory mechanism. Other changes in chromatin modifications during gametocyte development^5,56, besides changes in heterochromatin distribution, may also contribute to the regulation of gene expression in gametocytes.

Heterochromatin changes in maturing gametocytes affected several genes previously classified as CVGs^15,41 and positive for heterochromatin marks in asexual blood stages^6,7,8,9, but also a similar number of non-CVGs: several genes that are never heterochromatic during the IDC can be regulated by heterochromatin in gametocytes. Many of these genes belong to multigene families that include other genes that are variantly expressed during the IDC. The majority of the genes with differential heterochromatin coverage in gametocytes encode exported proteins, as previously reported⁷, consistent with the prominent role of erythrocyte remodelling during gametocyte development and differences with remodelling during the IDC^57,58. Many exported proteins involved in PfEMP1 traffic in asexual blood stages are no longer needed in gametocytes. Some specific gene families such as phist, fikk or hyp, involved in erythrocyte remodelling, are overrepresented among the genes with differential heterochromatin coverage in gametocytes. The expansion of heterochromatin at some specific subtelomeric regions and not others may reflect the distribution of these gene families in the genome. It is tempting to speculate that the conserved telomere-distal position of some gene families within subtelomeric regions⁴⁴ may have evolved to facilitate their silencing by heterochromatin spreading during sexual development.

Comparison of our results with previously published data describing the distribution of heterochromatin in sporozoites³¹ revealed that the boundaries of subtelomeric heterochromatin domains in sporozoites are more similar to maturing gametocytes than to asexual blood stages. This suggests that the expansion of subtelomeric heterochromatin observed in gametocytes is maintained during development in the mosquito. Previous studies analysing P. falciparum blood stages from controlled human malaria infections (CHMI) showed that var gene expression patterns are reset during transmission stages⁵⁹, and the epigenetic memory for the expression of clag3 genes⁶⁰ and several other (if not all) CVG families is also erased⁴¹. Furthermore, the observation of heterogeneous CVG expression patterns in blood stage parasites of a volunteer infected by a single sporozoite suggested that the reestablishment of variant gene expression patterns likely occurs during liver stages⁴¹. Based on these observations and the data presented here, we propose a model for heterochromatin dynamics across the full life cycle (Fig. 6) in which the global heterochromatin distribution in asexual blood stages is first modified in stage II/III gametocytes, involving mainly subtelomeric heterochromatin expansions. After this, heterochromatin distribution is maintained through the rest of gametocyte development and mosquito stages with only small additional changes. A second round of global heterochromatin redistribution occurs during liver stages, before parasites are released again to the bloodstream. This second round of global remodelling reverts subtelomeric heterochromatin expansions and likely involves removal and de novo formation of heterochromatin at most loci, resulting in a reset of the expression patterns of specific CVGs. Experiments following changes in heterochromatin distribution in the same parasite line across the full life cycle, which now can be reproduced in vitro⁶¹, will be necessary to confirm this model. It is not known if a similar heterochromatin dynamics across the life cycle occurs in other human malaria parasite species, such as P. vivax. The lack of a suitable in vitro culture system for P. vivax precludes a detailed characterisation of heterochromatin distribution at different life cycle stages in this parasite species. Of note, the majority of Plasmodium species, including P. vivax and P. cynomolgi, show much faster gametocyte development (~ 2 days) than P. falciparum (~ 10 days)⁶². In P. cynomolgi, heterochromatin expansions were observed in sporozoites, but it was not determined if they occurred in gametocytes or later during mosquito stages³³.

The mechanism underlying the expansion of subtelomeric heterochromatin during gametocyte development is not known. Research in model organisms has established that heterochromatin has the ability to spread into adjacent regions in a discontinuous and sequence-independent manner unless limited by boundary elements^1,2,3, which are not well-described in most eukaryotes, including Plasmodium spp. Our data shows that the boundaries for several subtelomeric heterochromatin domains are different between P. falciparum asexual blood stages and maturing gametocytes, but conserved between parasite lines of different genetic backgrounds. A possible mechanism that could explain the changes between asexual blood stages and gametocytes involves a different dose of specific heterochromatin regulators between asexual blood stages and gametocytes. The dose of some chromatin modifiers has been shown to determine the extension of heterochromatin domains in Drosophila position effect variegation (PEV) and in other systems^1,2,3,63. In this regard, the expression levels of the gene encoding the putative P. falciparum H3K9 methyltransferase, PfSet3 (PF3D7_0827800)^8,64, show a clear increase from roughly stage II/III gametocytes onwards⁴³, pointing to this protein as a candidate for underlying heterochromatin expansion during gametocyte development.

An unexpected observation from our results was that, in genes with differential heterochromatin coverage between asexual blood stages and gametocytes, transcriptional changes during sexual development preceded changes in heterochromatin occupancy. For genes silenced during sexual development, such as kahrp or pf332, this may simply reflect that the transcription factors necessary for their expression are already absent before stage II/III gametocytes, as a consequence of the new transcriptional program in gametocytes. This indicates that heterochromatin expansion is not the primary event for the silencing of these genes, but rather a secondary mechanism that locks the silenced state. For genes activated during sexual development, such as pfg14-744 and pfg14-748, increased transcript levels and also binding of the transcription factor PfAP2-G were first observed in stage I gametocytes, whereas heterochromatin depletion was not observed until stage II/III. This suggests that these genes may be activated within a heterochromatin environment, which would raise the intriguing possibility that PfAP2-G is a pioneer factor able to bind to and activate its target loci in spite of the presence of heterochromatin. Pioneer factors are a type of transcription factors that can bind DNA wrapped in nucleosomes and in a closed (heterochromatic) chromatin environment, and facilitate transcription by opening chromatin⁶⁵. However, we cannot completely rule out the possibility that population heterogeneity, for instance between male and female gametocytes³⁵, accounts for these results. Experiments testing if heterochromatin is present at these genes in the same individual stage I gametocytes in which PfAP2-G binding and active transcription occurs will be needed to directly establish if PfAP2-G is a pioneer factor.

Materials & methods

Parasite cultures and preparation of sexual and asexual parasite samples

The E5ind and NF54-gexp02-Tom lines have been previously described^20,51. Cultures were regularly maintained under standard P. falciparum culture medium containing Albumax II and no human serum. The E5ind line was cultured under 10 nM WR99210 (Jacobus Pharmaceuticals Co., USA) pressure to maintain the cam promoter integrated at the pfap2-g locus in an active state, but drug pressure was removed at the cycle of induction²⁰. The NF54-gexp02-Tom line was regularly maintained in medium supplemented with 2 mM choline (Sigma-Aldrich, C7527) to repress sexual conversion⁵¹.

Induction of sexual conversion in the E5ind line was performed as previously described by adding 10 nM rapamycin (Sigma-Aldrich, R0395) for 1 h to sorbitol-synchronised cultures at the trophozoite stage (20 h post-synchronisation)²⁰. Control cultures were treated, in parallel, with DMSO solvent. From one cycle before induction, cultures were maintained in erythrocytes filtered with Plasmodipur columns (R-Biopharm, 8011FILTER10U) to remove residual leukocytes and avoid human DNA contamination. After reinvasion (about 24 h after induction), 2 mM choline and 50 mM N-acetylglucosamine (Sigma-Aldrich, A3286) were added to the cultures, to improve gametocyte maturation and to prevent asexual parasite growth, respectively. Recombination at the edited pfap2-g locus after induction was confirmed by PCR analysis of genomic DNA extracted ~ 24 h after induction, as described²⁰.

To obtain TdTomato positive (TdTom +) and negative (TdTom-) rings, NF54-gexp02-Tom cultures were tightly synchronised to a 5 h age window using Percoll purification of mature forms followed by sorbitol lysis to eliminate mature forms 5 h later⁶⁶. Immediately after sorbitol lysis, when parasites were 0–5 h post-invasion (hpi), the choline supplement was removed to stimulate sexual conversion^22,67,68. The sexual conversion rate, determined by flow cytometry⁵¹, was ~ 50% in this experiment. After reinvasion, at ~ 11–16 hpi of the next cycle, TdTom + and TdTom- parasites were sorted in a BD FACS Aria SORP cell sorter. Erythrocytes were gated on SSC-A/FSC-A and then single cells were gated on FSC-H/FSC-A to avoid cell doublets or aggregates. Next, TdTom + and TdTom- cells were separated using the combination of 488 nm (detects autofluorescence) and 561 nm (detects TdTomato) lasers (Supplementary Fig. S14b). Sorting parameters included: sheath pressure set at 20 psi, use of a 100 µm nozzle tip and temperature maintained at 37ºC. Sorted cells were collected into tubes containing culture medium supplemented with 2 mM choline. We collected a total of 3.87 × 10⁶ TdTom + cells (sexual rings and possibly some stage I/II gametocytes formed at the previous cycle and diluted during culture growth) and 7.8 × 10⁷ TdTom- cells (containing 3.74 × 10⁶ asexual rings and uninfected erythrocytes). Sorting took approximately 7 h. The purity of the resultant TdTom + and TdTom- preparations was > 90%, according to flow cytometry analysis (Supplementary Fig. S14c). Uninfected erythrocytes were added to the TdTom + sorted cells to adjust the parasitaemia to that of the TdTom- sorted cells (~ 5%).

ChIP-seq experiments

ChIP-seq experiments were performed essentially as described^20,41. Parasites were isolated from erythrocytes using saponin lysis. After formaldehyde cross-linking, chromatin was extracted with the MAGnify kit (Life Technologies). Sonication to obtain 150—200 bp DNA fragments was performed with a Covaris M220 apparatus. Immunoprecipitation was done with 2 µg of chromatin and 4 µg of antibody against H3K9me3 (Diagenode, C15410193, lot A2217P). Sequencing libraries were prepared from 5 ng of immunoprecipitated DNA as previously described^20,41, using conditions adapted to the extremely AT-rich P. falciparum genome⁶⁹. NEBNext Multiplex Oligos for Illumina (New England BioLabs) were used for multiplexing, and 8 to 10 PCR cycles with the KAPA HiFi PCR kit (Kapa Biosystems) were used for amplification. For all purification steps, AMPure XP beads (Beckman Coulter, A63882) were used. Sequencing was performed using an Illumina NextSeq 550 system, obtaining 8 to 15 million 150 bp paired-end reads per sample.

For the ChIP-seq analysis of NF54-gexp02-Tom FACS-sorted parasites, we used a similar protocol but library preparation was performed using the NEBNext Ultra DNA Library Prep Kit (New England BioLabs, ref. E7370) for Illumina following manufacturer’s specifications except for end repair, which was performed for 1 h at 45ºC (instead of 30 min at 65ºC). DNA was amplified for 15–19 cycles.

RT-qPCR and ChIP-qPCR assays

ChIP-qPCR analysis of uninduced E5ind cultures at the asexual ring stage and induced cultures at different sexual stages was performed approximately as described⁷⁰. In brief, chromatin was extracted as for the ChIP-seq experiments, but DNA was sonicated using a Bioruptor 300 (Diagenode) to obtain ~ 500 bp fragments. After immunoprecipitation (as for ChIP-seq experiments), input and immunoprecipitated samples were analysed by qPCR in triplicate wells using the standard curve method (with a standard curve included in each plate for each primer pair) and the primers described in Supplementary Table S8. In each experiment, we included ama1 and uce as heterochromatin negative controls and the var gene PF3D7_1240300 and clag3.2 as heterochromatin positive controls.

RNA for RT-qPCR analysis was prepared using the TRIzol method, treated with DNAse I and purified as previously described^66,70. Reverse transcription with a mixture of random and oligo (dT) primers and qPCR analysis of the cDNAs using the PowerSYBR Green master mix (Applied Biosystems) and the standard curve method were also performed as previously described^66,70. All primers used are listed in Supplementary Table S8. Transcript levels were normalised against the serine–tRNA ligase gene (serrs, PF3D7_0717700), which is expressed at relatively stable levels in all blood stages except for very young rings, which were not included in this study.

ChIP-seq data analysis

New ChIP-seq data generated in this study as well as previously published data^7,31 were analysed approximately as previously described⁹. After quality control and trimming, reads were aligned against the P. falciparum 3D7 genome (PlasmoDB v. 55) using bowtie2 (v. 2.3.0). Duplicates reads were removed and peak calling for each individual sample was performed using MACS2 software (v. 2.2.7) with parameters: -f BAMPE -B -g 2.41e7 –keep-dup all –fe-cutoff 1.5 -nomodel –extsize 150. IP tracks were normalised against their inputs using the Deeptools suite, with the bamCompare option. Coverage at specific genome regions was calculated from normalised mean log₂ (IP/input) bedgraph tracks, using the bedtools -map option.

To calculate the proportion of the genome of each parasite line covered by a MACS2 H3K9me3 ChIP-seq speak, each .narrowPeak file containing all peak intervals were converted into a fasta file using Bedtools (-getfasta option) and, after counting the number of base pairs covered by the mark, it was divided by the size of the full genome.

To identify differential peaks, which here refers to regions with differential coverage between asexual rings and sexual stages, we used a previously described in-house method⁹, with some modifications. Each sexual stage was compared with asexual rings from the same experiment. In brief, we first calculated the input-normalised coverage for all 100 bp bins overlapping a MACS2 peak in one or more of the samples and adjusted the distribution of coverage intensities to a gaussian mixture distribution with two components (the rationale being that one component accounts for the non-enriched regions and the other one for the enriched regions). Next, we calculated for each bin in each sample a cumulative density function (CDF) value for each of the two components. Bins with a CDF value difference of > 0.2 (for any of the two components) were classified as having differential coverage. We merged bins with differential coverage separated by < 1,000 bp, and filtered out differential peaks of < 150 bp after merging. Finally, we only retained as differential peaks the intervals that were common between the two replicates and removed regions of ≤ 500 bp or with an absolute value of the log₂ of the coverage fold-change between the sexual stage and asexual rings of < 1. The final differential peaks were then intersected with protein coding genes (either the −1,000 to + 500 bp from the ATG region or the rest of the annotated transcript products, including 5’UTR and 3’UTR when available in PlasmoDB) and other genetic elements. Genes for which the -1,000 to + 500 bp region overlapped with a differential coverage peak were retained in the differential coverage gene lists if the log₂ of the H3K9me3 coverage fold-change between asexual rings and the sexual stage tested was ≥ 1 in this −1,000 to + 500 bp region.

Determination of the extension of heterochromatin domains at subtelomeric regions or internal islands

From the initial .narrowPeak files generated by individual MACS2 peak calling, we merged peaks separated by ≤ 10 Kb using bedtools (parameters merge -i .narrowPeak -d 10000) and eliminated the peaks with a length < 500 bp. Next, to define the full extent of each heterochromatin domain we merged regions separated by ≤ 60 Kb and, for each subtelomeric region, extended the regions to the extreme of the chromosome regardless of whether or not it was covered by a MACS2 peak (Supplementary Fig. S3a). Finally, we used visual inspection of IGV tracks to confirm consistency between replicates and edited manually the limits of the heterochromatin region in one of the replicates when there was a large discrepancy with the other replicate and an error of the algorithm was apparent (one subtelomeric domain was manually edited).

Data availability

The new ChIP-seq data presented in this article was deposited in the GEO database with accession n. GSE252334. ChIP-seq data can also be visualised at the UCSC genome browser (http://genome.ucsc.edu/cgi-bin/hgTracks?db=hub_4042823_pfa2&lastVirtModeType=default&lastVirtModeExtraState=&virtModeType=default&virtMode=0&nonVirtPosition=&position=chr1%3A1%2D266268&hgsid=2358478401_dIujXaA0tJH9ttmYZznyqY3gb48N ).

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Acknowledgements

We are grateful to Alba Pérez-Cantero for help with training and advice with parasite cultures and ChIP experiments, and for critical reading of the manuscript. This work was supported by grants PID2019-107232RB-I00 and PID2022-137863OB-I00 to A.C. from the Spanish Ministry of Science and Innovation (MCIN)/ Agencia Estatal de Investigación (AEI, https://doi.org/10.13039/501100011033), co-funded by the European Regional Development Fund (ERDF/FEDER, European Union), and “la Caixa” Banking Foundation (HR18-00267 to A.C.). L.M.-T. was supported by a fellowship from the Spanish Ministry of Economy and Competitiveness (BES-2017-081079), co-funded by the European Social Fund (ESF). This research is part of the ISGlobal’s Program on the Molecular Mechanisms of Malaria, which is partially supported by the Fundación Ramón Areces. We acknowledge support from the grant CEX2023-0001290-S funded by MCIN/AEI/ https://doi.org/10.13039/501100011033, and support from the Generalitat de Catalunya through the CERCA Program.

Funding

Ministerio de Ciencia e Innovación (MCIN), BES-2017-081079, PID2019-107232RB-I00, PID2022-137863OB-I00, ‘la Caixa’ Foundation, HR18-00267.

Author information

Lucas Michel-Todó
Present address: Vall d’Hebron Institute of Oncology (VHIO), Barcelona, Spain

Authors and Affiliations

ISGlobal, Barcelona, Spain
Sandra Nhim, Elisabet Tintó-Font, Núria Casas-Vila, Lucas Michel-Todó & Alfred Cortés
Facultat de Medicina i Ciències de la Salut, Universitat de Barcelona (UB), Barcelona, Spain
Sandra Nhim, Elisabet Tintó-Font, Núria Casas-Vila, Lucas Michel-Todó & Alfred Cortés
ICREA, Barcelona, Spain
Alfred Cortés

Authors

Sandra Nhim
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Elisabet Tintó-Font
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Núria Casas-Vila
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Lucas Michel-Todó
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Alfred Cortés
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Contributions

S.N. performed the majority of experiments, analysed the data and prepared the figures (except for Fig. 6, prepared by E.T.-F.). E.T.-F. and N.C.-V. performed the experiments with FACS-sorted parasites and provided support for other experiments. L.M.-T. contributed to data analysis. A.C. and S.N. designed the experiments, interpreted the results and wrote the manuscript. A.C. conceived the project, obtained funding and provided overall supervision. All authors reviewed the manuscript.

Corresponding author

Correspondence to Alfred Cortés.

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Nhim, S., Tintó-Font, E., Casas-Vila, N. et al. Heterochromatin dynamics during the initial stages of sexual development in Plasmodium falciparum. Sci Rep 14, 23180 (2024). https://doi.org/10.1038/s41598-024-73981-w

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Received: 12 June 2024
Accepted: 23 September 2024
Published: 05 October 2024
DOI: https://doi.org/10.1038/s41598-024-73981-w