Metabolic crosstalk between the mitochondrion and the nucleus is essential for Toxoplasma gondii infection

Zhang, Hongxi; Ji, Nuo; Su, Shuxin; Zhao, Meng; Du, Huiyu; Sahoo, Lakesh Kumar; Wu, Yi; Feng, Yaoyu; Gupta, Nishith; Xiao, Lihua; Xia, Ningbo

doi:10.1038/s42003-025-07823-4

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Published: 07 March 2025

Metabolic crosstalk between the mitochondrion and the nucleus is essential for Toxoplasma gondii infection

Communications Biology volume 8, Article number: 384 (2025) Cite this article

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Abstract

Toxoplasma gondii, an intracellular pathogenic protist with a remarkable ability to infect a wide range of host cells, displays an equally exceptional design of its carbon metabolism. There are, however, critical gaps in our understanding of the metabolic network in T. gondii. We characterized the mito-nuclear metabolism and organelle coupling during its acute infection (lytic cycle). The major enzymes of the TCA cycle, i.e., citrate synthase (CS1), succinyl-CoA synthase alpha subunit (SCSα), succinate dehydrogenase (SDHA) and FAD malate dehydrogenase (MDH-FAD) located in the parasite mitochondrion support its asexual reproduction but are not needed for its survival. The SCSα and SDHA mutants are nearly avirulent in a mouse model, and they can protect the host against a lethal challenge infection. Genetic deletion of MDH-FAD dysregulated glucose-derived carbon flux, leading to a collapse of the mitochondrial membrane potential. The parasite also harbors a cytosolic isoform of MDH and a nuclear malic enzyme (ME) contributing to malate oxidation; however, only the latter is essential for the lytic cycle. Expression of ME in the nucleus is crucial for the parasite development. Besides, conditional knockdown of ME impairs the histone acetylation and disrupts the expression of several genes in tachyzoites. Our work discloses novel network design features of T. gondii and highlights the therapeutic and vaccination potential of the parasite metabolism.

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Introduction

Toxoplasma gondii, a prevalent zoonotic parasite, infects many warm-blood organisms and can reproduce in several nucleated host cells¹. It has a complex life cycle, with sexual reproduction occurring exclusively in felid hosts and asexual development in a wide range of intermediate hosts, such as pigs, cattle, sheep and humans¹. The parasite exists in two infectious stages in its hosts: fast-replicating tachyzoites and slow-growing bradyzoites. Acute toxoplasmosis in pregnant individuals or animals can cause abortion, stillbirth or fetal deformities¹. In contrast, the chronic disease usually persists for the entire life of the infected host and can result in recrudescence upon impairment of the immunity. These outcomes seriously threaten public health and socioeconomic development. Currently, pyrimethamine and sulfadiazine are the only common drugs for treating acute toxoplasmosis^2,3,4 and none are available for against chronic infection. Extended drug usage is also associated with adverse effects and drug resistance^2,3,4. There is only one commercial vaccine (Toxovax), which is weakly virulent and used exclusively in sheep, with a potential risk of reverting to virulence. Therefore, there is an urgent need to explore additional drug and vaccine targets to control the disease^5,6. Although the parasite metabolism remains central to drug development, its limited knowledge has hindered the discovery of new therapeutic vulnerabilities in T. gondii.

Toxoplasma’s acutely infectious tachyzoite stage utilizes glucose- and glutamine-derived carbon to fuel their carbon metabolism and, thereby, the lytic cycle^7,8,9,10,11. Glucose and glutamine are metabolized through glycolysis and the TCA cycle, respectively, and both nutrients cooperate to enable a rapid propagation of tachyzoites. Glucose enters the parasite through a high-affinity transporter located in the plasma membrane⁷ and then converted to pyruvate via glycolysis^8,10,12. Pyruvate is imported into the mitochondrion via the mitochondrial pyruvate carrier (MPC) complex, where it is converted to acetyl-coenzyme A (Ac-CoA) by a branched-chain α-ketoacid dehydrogenase E1 subunit (BCKDH-E1α), feeding into the TCA cycle^13,14. However, tachyzoite mutants lacking MPC or BCKDH-E1α are still viable and partially virulent^13,14. Glutamine is first converted to glutamate, entering the TCA cycle as α-ketoglutarate or as succinate via a γ-aminobutyric acid (GABA) shunt⁸. Tachyzoites can also survive the deletion of glutamate decarboxylase (GAD), the first enzyme of the GABA shunt, although their growth and virulence are impaired⁸. These studies reveal significant vulnerabilities and plasticity in the parasite metabolism. The TCA cycle is a vital eukaryotic pathway, but its significance for the asexual growth of T. gondii remained ambiguous. The tachyzoite stage harbors all enzymes expressed in the mitochondrion¹⁵, and stable isotopic labeling showed evidence of a canonical oxidative-type TCA cycle⁸. The first step is catalyzed by citrate synthase 1 (CS1), forming citrate from acetyl-CoA and oxaloacetate. Subsequently, aconitase (ACO) isomerizes citrate to isocitrate, which is then oxidized to α-ketoglutarate by isocitrate dehydrogenase I (IDH1). α-ketoglutarate serves as a substrate for α-ketoglutarate dehydrogenase (KDH), producing succinyl-CoA, which is converted to succinate by succinyl-CoA-synthetase (SCS). Depletion of the SCSα subunit only partially (~30% reduction) impairs parasite growth, suggesting an alternative pathway for succinate production¹⁵. A succinic semialdehyde dehydrogenase (SSADH) is also present in the parasite genome, which can potentially produce succinate from succinic semialdehyde¹⁶. The latter may originate from γ-aminobutyric acid (GABA) shunt in tachyzoites⁸. However, the physiological role of SSADH in T. gondii remains unknown.

Succinate is oxidized by succinate dehydrogenase (SDH) to form fumarate, which in turn is hydrated to produce malate by the catalytic action of fumarate hydratase (FH). Malate is oxidized to oxaloacetate by malate dehydrogenase (MDH). Oxaloacetate has multiple fates depending on the physiological status of the parasite. It can fuse with acetyl-CoA, completing the TCA cycle⁸, and is used for aspartate synthesis via aspartate aminotransferase/glutamic oxaloacetic transaminase (AST/GOT)¹⁰. Additionally, during gluconeogenesis in glucose-starved conditions, oxaloacetate drives the formation of phosphoenolpyruvate by a mitochondrial isoform of phosphoenolpyruvate kinase (PEPCK)¹¹. The latter two are cataplerotic reactions, which dispose of TCA cycle intermediates to other pathways. Glutamate dehydrogenase (GDH) catalyzing the formation of glutamate from α-ketoglutarate and the malic enzyme (ME) producing pyruvate from malate are other cataplerotic proteins encoded by the parasite, but their physiological relevance has not been studied. The parasite also harbors two variants of malate dehydrogenases: MDH and MDH-FAD¹⁵; however, neither has been examined in T. gondii.

The cataplerotic contribution of the TCA cycle to biosynthetic pathways necessitates anaplerotic replenishment of its intermediates to ensure continued operation. Pyruvate carboxylase (PyC), which converts pyruvate to oxaloacetate in the mitochondrial matrix, is the typical anaplerotic enzyme and is also present in T. gondii. However, it is dispensable in both glycolysis-competent and glycolysis-deficient tachyzoites¹¹. Hence, its metabolic role remains unknown. This study presents a systematic dissection of the central carbon metabolism in T. gondii by genetic mutagenesis of CS1, SSADH, SCSα, SDHA, ME, MDH-FAD and MDH. In-depth phenotyping of the mutants coupled with metabolomics discloses novel design features and therapeutic targets in the core metabolic network of T. gondii.

Results

The CS1 is needed for the growth but not for the survival of tachyzoites

The genome of T. gondii harbors three isoforms of citrate synthase (www.ToxoDB.org); however, only CS1 is located in the tachyzoite mitochondrion¹⁵. We started our work confirming the subcellular location of CS1 in the mitochondrion by 3’-genomic tagging (Fig. 1a). The CS1-specific CRISPR/Cas9 construct and the homology donor amplicon encoding YFP-DHFR* were transfected into tachyzoites to generate a Δcs1 mutant (YFP⁺) (Supplementary Fig. 1a). Surprisingly, we could isolate a CS1 deletion strain despite its predicted essentiality in tachyzoites^17,18. Genomic PCR screening (Supplementary Fig. 1b), genome sequencing (Supplementary Fig. 4a), and indirect immunofluorescence assays (IFA) confirmed the deletion of the CS1 locus in the mutant (Supplementary Fig. 1c). The Δcs1 mutant grew slower in routine culture and produced smaller plaques due to poor replication than the parental strain (Fig. 1b–d). The Δcs1 mutant was severely attenuated in mice as judged by their higher survival rate at doses of 10² parasites/mouse (84%, Fig. 1e). Even at higher doses (up to 10⁴ parasites/mouse), 84% of animals survived the infection (Fig. 1e). The parasite load in mutant-infected animals was almost 10⁴ times lower than the parental (DiCre) strain (Fig. 1f). These results reveal that CS1 is required for optimal growth and virulence but is not essential for tachyzoite survival.

**Fig. 1: CS1 is required for parasite growth.**

Deletion of SCSα compromises parasite growth, whereas SSADH is dispensable

To gain additional insight into the importance of the TCA cycle, we investigated SCSα. The HA-tagged SCSα is also expressed in the mitochondrion (Fig. 2a). Earlier work has shown that ATc-mediated depletion of SCSα transcript inflicts only minor growth defects in tachyzoites¹⁵. Such an unexpected (poor) phenotype upon conditional knockdown can be attributed to a lack of stringent regulation by ATc or a nonessential role of SCSα. We, therefore, engineered a SCSα deletion mutant by CRISPR/Cas9-assisted homologous gene replacement (Supplementary Fig. 1f). The Δscsα (YFP⁺) mutant was isolated after pyrimethamine selection, diagnostic PCR screening and immunostaining (Supplementary Fig. 1g, h). The Δscsα strain was notably viable in prolonged culture despite severely impaired plaque formation and replication defect (Fig. 2b–d). The data indicate a partial TCA cycle operation is sufficient for parasite survival.

Fig. 2: SCSα is vital for the lytic cycle of *T. gondii.*

We also investigated SSADH, which can produce succinate from succinic semialdehyde¹⁶. The 3’HA-tagged SSADH was expressed in the mitochondrion, co-localizing with HSP60 – a known organelle marker (Fig. 3a). We next engineered a Δssadh strain using CRISPR/Cas9-mediated homologous recombination (Supplementary Fig. 1d), pyrimethamine selection and PCR screening (Supplementary Fig. 1e). The gene deletion had only a minor impact on plaque formation (Fig. 3b, c), replication (Fig. 3d) and virulence (Fig. 3e), ruling out SSADH as a primary source for succinate entry into the TCA cycle. Functional redundancy of SSADH and SCSα was examined by deleting SSADH in the Δscsα strain, resulting in a double mutant (ΔscsαΔssadh) (Supplementary Fig. 2a–c). Phenotyping revealed a further reduction in the replication of ΔscsαΔssadh (Fig. 3f); however, the mutant could be maintained in cultures (Fig. 3g). This finding consolidates our other data on the metabolic flexibility in the TCA cycle.

**Fig. 3: *SSADH* deletion only mildly affects parasite development.**

The Δscsα mutant is avirulent, and it can protect mice from challenge infection

Our further work evaluated the in vivo relevance of the Δscsα strain by intraperitoneal injection into the ICR mice. As expected, the parental (DiCre) strain killed almost all animals within 2 weeks (Fig. 2e). In contrast, mice infected with Δscsα survived and exhibited no clinical signs. Even at much higher doses of the mutant (~10⁶ tachyzoites/mouse), no apparent virulence was observed (Fig. 2e). The Δscsα and DiCre strains (10⁴ tachyzoites/animal) were also quantified in the peritoneal fluid by qPCR five days post-infection. The parasite burden in mice parasitized by the mutant was at or below the detection threshold of qPCR (Fig. 2f). We tested whether the host immune response underpins the attenuated growth of the Δscsα strain in mice. Immunodeficient Balb/c-nu mice were infected (10⁴ parasites/animal), and the parasite load was quantified. The outcome resonated with the immune-competent ICR mice, as the Δscsα tachyzoites were barely detectable in the Balb/c-nu mice (Fig. 2f).

In the next step, we explored the potential of genetically attenuated Δscsα strain as a whole-cell vaccine because it could be cultured in vitro but failed to proliferate in vivo. Indeed, the Δscsα-immunized mice rapidly cleared the challenge infection by the RH-Luc tachyzoites, as evident by bioimaging of luminescence (Fig. 2g, h). Furthermore, all Δscsα-vaccinated animals survived infection of RH-Luc and ME49 tachyzoites (Fig. 2i, j). In conclusion, our data highlight the therapeutic and vaccination potential of SCSα protein and Δscsα mutant, respectively. The inability of the Δscsα strain to propagate itself in mice also suggests a critical in vivo role of SCSα in the mitochondrial metabolism.

SDHA and FH are vital, but MDH and OMT are dispensable in tachyzoites

In the next step, succinate is oxidized to fumarate by SDH, also known as complex II, because it delivers TCA cycle-derived electrons to the mitochondrial electron transport chain (mETC)¹⁹. Toxoplasma SDH is a large complex of over 500 kDa, broadly divided into a membrane-anchoring domain and an enzymatic core. The latter, localizing in the matrix, comprises SDHB and SDHA. The SDHA contains the succinate-binding site and the FAD cofactor¹⁹. We generated an SDHA mutant confirmed by PCR screening and immunostaining (Supplementary Fig. 1i–k). The Δsdha was severely compromised compared to the parental strain (Fig. 4a–c). Accordingly, the mutant could not propagate in vivo (Fig. 4d), resulting in the survival of all infected mice (Fig. 4e). Even at much higher doses of the Δsdha mutant (~10⁵ tachyzoites/mouse), no significant virulence was observed (Fig. 4f). Similar to the Δscsα mutant, immunization of mice with Δsdha induced effective host immune protection against challenge with wild-type RH-Luc and ME49 (Fig. 4g–j). Next, we tried to ablate the fumarate hydratase (FH) protein, converting SDHA-derived fumarate to malate. However, our efforts to delete the FH gene were unsuccessful, suggesting its necessity for parasite survival.

**Fig. 4: Knockout of *SDHA* is detrimental for the parasite.**

Our following work focused on MDH, oxidizing malate into oxaloacetate. It was earlier shown to localize on the parasite mitochondrion¹⁵. However, 3’-genomic tagging of MDH with an HA tag revealed its expression in the cytoplasm, localizing with ALD (Fig. 5a). To assess its physiological importance, we engineered a knockout strain using the CRISPR/Cas9 system (Supplementary Fig. 3a). PCR screening (Supplementary Fig. 3b) and genome sequencing (Supplementary Fig. 4b) verified the deletion of MDH. The plaque (Fig. 5b, c), replication (Fig. 5d) and virulence (Fig. 5e) assays displayed that loss of MDH had no discernible effect on the parasite. We were, therefore, promoted to consider alternative pathways that could supply malate to the cytoplasm. Tachyzoites encode a 2-oxoglutarate/malate translocase (OMT, TGGT1_274060), which may transport malate from the mitochondrion to the cytoplasm. Indeed, the HA-tagged OMT was expressed in the mitochondrion. Subsequently, the Δomt mutant was isolated by CRISPR/Cas9, pyrimethamine selection, screening PCR and immunostaining (Supplementary Fig. 5a, b). Deleting the OMT gene only slightly affected the tachyzoite growth, as evaluated by plaque, replication, virulence and parasite load assays (Supplementary Fig. 5c–f). The data imply a functional redundancy between OMT and MDH for supplying malate to the parasite cytosol.

**Fig. 5: MDH is not required for the lytic cycle.**

Toxoplasma encodes a functional MDH-FAD located in mitochondria

Our data above describing cytoplasmic expression and the nonessential nature of MDH indicated the presence of an isoform for malate oxidation in the parasite mitochondrion. The database search identified a FAD-dependent malate-dehydrogenase (MDH-FAD). 10×HA fusion of the native MDH-FAD by 3′-genomic tagging disclosed a mitochondrial expression, co-localizing with HSP60 (Fig. 6a). TgMDH-FAD was further examined by functional complementation of a Saccharomyces cerevisiae mutant (ScΔmdh1), which cannot grow on a non-fermentable carbon source²⁰. As shown (Fig. 6b), the parental (BY4741) and ScΔmdh1 strains transformed with the empty plasmid (pRS416) or construct (pRS416-MDH-FAD) were all able to grow on the fermentable carbon source. As expected, the mutant growth was restored by expressing TgMDH-FAD, and pRS416 failed to rescue its growth on glycerol (YPG, non-fermentable carbon source) (Fig. 6c). We constructed an ScΔmdh-TgMDH-FAD strain expressing the parasite enzyme from chromosome XI to validate our results. Consistently, TgMDH-FAD rescued the ScΔmdh1 mutant on YPG plates (Fig. 6c), showing its functionality in yeast.

**Fig. 6: *Toxoplasma* expresses MDH-FAD in the mitochondrion.**

The MDH-FAD is required for the asexual reproduction of tachyzoites

In the next step, we investigated the metabolic relevance of MDH-FAD in tachyzoites. A pool of transgenic parasites expressing rapamycin-inducible dimerizable Cre (DiCre) and harboring loxP-flanked MDH-FAD (TgMDH-FAD-cKD) was generated. Subsequently, a Δmdh-fad strain was produced by treating the TgMDH-FAD-cKD strain with rapamycin (Supplementary Fig. 3c). The mutant was validated by PCR (Supplementary Fig. 3d), genome sequencing (Supplementary Fig. 4c) and immunofluorescence (Supplementary Fig. 3e) assay. Although the Δmdh-fad strain formed notably smaller plaques (Fig. 7a, b) and exhibited much smaller parasitophorous vacuoles (Fig. 7c), the Δmdh-fad mutant could be maintained in routine culture for a long time. In extended work, we infected mice with the Δmdh-fad strain and quantified the parasite load in the peritoneal fluid (Fig. 7d). Indeed, the deletion of MDH-FAD significantly reduced the parasite proliferation. Besides, mice parasitized by the Δmdh-fad mutant survived longer than those infected with the parental strain (Fig. 7e), indicating a reduced virulence upon deletion of MDH-FAD. These results emphasize the crucial function of MDH-FAD in promoting tachyzoite development in vitro and in vivo.

**Fig. 7: MDH-FAD is required for *T. gondii* growth.**

The Δmdh-fad strain displays impaired metabolism and membrane potential

To investigate whether deletion of MDH-FAD impacted the intermediates of the central carbon metabolism, we labeled extracellular tachyzoites of the Δmdh-fad and parental strains with [¹³C]-glutamine for 4 h. We determined the inclusion of glutamine-derived carbon (¹³C) into glycolytic and TCA metabolites using LC-MS. A change in ¹³C labeling of α-ketoglutarate and succinate was not apparent (Fig. 7f). However, the tracer incorporation into malate was significantly increased (Fig. 7f). We also noted that the inclusion of glucose-derived ¹³C into fumarate and malate was increased upon deleting MDH-FAD (Fig. 7g). As expected, the abundance of malate was considerably increased in the mutant (Fig. 7h). We observed a difference in malate labeling of the Δmdh-fad strain fed with [¹³C]-glutamine and [¹³C]-glucose (Fig. 7h, i). The glucose-derived ¹³C into the TCA cycle requires oxaloacetate, whereas the influx of [¹³C]-glutamine-derived carbon into malate is not dependent on oxaloacetate. A deletion of MDH-FAD may reduce the availability of oxaloacetate in the mitochondrion, resulting in a lower glucose-derived ¹³C inclusion into malate. Our attempts to detect oxaloacetate in the parasites were unsuccessful, likely due to its rapid conversion to aspartate by aspartate aminotransferase (AST). We identified a mitochondrion-localized AST in T. gondii, as shown by the 10×HA gene fusion of the native AST (Supplementary Fig. 6a). The deletion of AST in the DiCre strain resulted in a notable reduction in plaque formation, replication, virulence, and parasite load (Supplementary Fig. 6b–g).

Interconversion of malate and oxaloacetate by MDH-FAD depends on NAD⁺/NADH. We, therefore, measured the total NAD⁺ and NADH levels in the mutant (Fig. 7j), which declined by about 80% upon loss of MDH-FAD expression. Because NADH is an electron donor in the inner mitochondrial membrane, we examined the membrane potential (Δψm) using mitotracker staining (Fig. 7k). Intracellular tachyzoites of the parental strain had well-stained mitochondrion, classified as Δψm⁺(Fig. 7l). In contrast, mitochondrial staining was not detectable in over 80% of the Δmdh-fad parasites (Fig. 7l), signifying a collapse of Δψm in the organelle upon deletion of MDH-FAD.

The ME, localized in the nucleus, is essential for parasite development

Our follow-up work focused on ME, which catalyzes malate to pyruvate while producing NADPH. We first determined its subcellular localization using antisera and by ectopic expression of epitope-tagged ME (Fig. 8a, b). Immunofluorescent staining showed that ME co-localized with Hoechst (DNA dye, Fig. 8a, b). In the next step, we made a conditional mutant of ME using the auxin-inducible degron (AID) system (Supplementary Fig. 3f) because its knockout via CRISPR/Cas9-mediated homologous replacement was not feasible. PCR screening of the drug-resistant parasite clones identified a miniAID-ME mutant (Supplementary Fig. 3g), in which the protein could be rapidly degraded by indole-3-acetic acid (IAA, Fig. 8c). A knockdown of ME resulted in reduced plaque size (Fig. 8d) and smaller vacuole size distribution (Fig. 8e). Besides, the ICR mice infected by the miniAID-ME tachyzoites (10⁴, i.p.), followed by treatment with IAA in drinking water, demonstrated a highly-reduced parasite load compared to the untreated control (-IAA) and the parental strain (Fig. 8f). These results underline the critical role of ME in tachyzoite propagation.

**Fig. 8: ME is critical for the lytic cycle of tachyzoites.**

The ME-depleted strain exhibits a selective perturbation of transcriptome

ME produces pyruvate from malate, a crucial precursor for several metabolic pathways in the cytoplasm, apicoplast, and mitochondrion¹². While not known in T. gondii, pyruvate metabolism in the nucleus is critical for histone acetylation and chromatin remodeling in mammalian cells^21,22. To determine whether the ME depletion impacts histone acetylation in tachyzoites, we conducted an immunofluorescence analysis using the specificity of the H3K9ac-directed antibodies (Fig. 8g). Our findings revealed that ME depletion markedly decreased the H3K9ac signal intensity in the parasite nucleus (Fig. 8h), indicating that ME-derived pyruvate plays a pivotal role in acetyl-CoA synthesis for epigenetic regulation in tachyzoites.

The miniAID-ME strain was grown with or without IAA for 12 h, and then samples were collected for RNA sequencing to assess the global consequences of ME knockdown. The ME-depleted strain exhibited a lower expression of 2342 transcripts, while 1731 were more abundant (Supplementary Fig. 7a). Several histone deacetylases and histone acetyltransferases, including HDAC1, HDAC2, HDAC5, SiR2, MYST, GCN5, and GNAT family protein, displayed dysregulated expression (Supplementary Fig. 7b). The qPCR verified the lower expression of SIR2 and more abundance of GCN5-B, HDAC5 and MYST-B transcripts in the ME-depleted mutants (Supplementary Fig. 7c). Furthermore, KEGG enrichment analysis revealed that DNA replication, mismatch repair and amino sugar and nucleotide sugar metabolism were the most significantly affected in instances where ME expression was lacking (Supplementary Fig. 7d).

Nuclear localization of ME is imperative for tachyzoite growth

ME comprises the N-terminal, Tudor, and C-terminal domains (Fig. 9a). Notably, the C-terminal contains two nuclear localization signals (Fig. 9a). To determine the functional relevance of these domains, we complemented the miniAID-ME strain by the N-terminal domain, both the N- and C-terminal domains, or the complete open reading frame of ME at the UPRT locus (Fig. 9b). A diagnostic PCR of each strain confirmed the integration of the respective insert (Supplementary Fig. 8a–c, comp-ME, comp-ME-N/C, comp-ME-N). Immunostaining revealed a nuclear localization in the comp-ME and comp-ME-N/C strains (Fig. 9c). In contrast, the comp-ME-N strain showed a cytosolic expression (Fig. 9c), indicating a role of the C-terminal in the nuclear transport of ME. As anticipated, the comp-ME strain displayed a fully restored lytic cycle (plaque formation, Fig. 9d) and replication (Fig. 9e) in the IAA-treated/ME-depleted mutant. A recovered phenotype of the comp-ME-N/C strain (+IAA) suggested a nonessential nature of the Tudor domain. Unlike the nuclear-localized constructs, the IAA-exposed comp-ME-N strain could not grow (Fig. 9d, e), highlighting the importance of ME expression in the parasite nucleus.

**Fig. 9: Functional analysis of the ME domain.**

Discussion

Herein, we demonstrate the network design of carbon metabolism in the mitochondrion and nucleus of a globally prevalent and clinically relevant human and animal pathogen, Toxoplasma gondii. Our integrative approach discloses previously unknown plasticity, inter-organelle cooperativity, vulnerability and therapeutic potential of major metabolic enzymes in the parasite mitochondrion, cytosol and nucleus (Fig. 10).

Fig. 10: Mito-nuclear carbon metabolism in *Toxoplasma.*

We show a significant role of the mitochondrial isoform of citrate synthase (CS1) for in vitro growth of tachyzoites and in vivo virulence. A similar outcome was reported recently²³. Our study revealed a more pronounced reduction in virulence of the Δcs1 mutants, which may be attributed to the disparate genetic backgrounds of the parental strains. The depletion of CS1 partly phenocopies the mutants of MPC1 and MPC2 transporting glycolysis-derived pyruvate into the mitochondrion and BCKDH-E1α subunit facilitating acetyl-CoA synthesis from MPC-imported pyruvate. These knockout strains are not lethal to the parasite survival in vitro, but exhibit varying degrees of attenuation in their virulence in mice^8,14. It has been reported that the mitochondria-localized 2-methylcitrate synthase (PrpC) has citrate synthase activity, although the catalytic efficiency of CS1 is notably higher than that of PrpC²³. Consequently, when CS1 is knocked out, PrpC may facilitate citrate formation from acetyl-CoA, thereby maintaining the functionality of the TCA cycle (Fig. 10). T. gondii also expresses a CS2 in the cytoplasm, producing citrate²³, which is hypothesized to be transported into the apicoplast and subsequently serve as a substrate for ACO to generate isocitric acid. The latter is metabolized by IDH2 to produce α-ketoglutarate and NADPH, contributing to FASII pathways in the organelle (Fig. 10). This premise is supported by the evidence that the parasite encodes an aconitase (ACO) and two isocitrate dehydrogenases (IDH1 and IDH2). The ACO is dual-localized in the mitochondrion and the apicoplast²⁴, whereas The IDH1 and IDH2 are present in the mitochondrion and apicoplast, respectively¹⁵. Whether CS2-derived citrate can traffic to the apicoplast and mitochondrion to support biosynthetic pathways in both organelles requires additional research.

SCSα and SSADH may synergize to supply succinate to the cycle (Fig. 10). A notably mild phenotype of the Δssadh mutant suggests that the GABA-derived succinic semialdehyde is not a significant contributor of succinate. Similarly, loss of GABA-AT, which converts GABA to succinic semialdehyde, does not affect parasite growth¹⁶. By contrast, the Δscsα strain exhibited a more severe phenotype which indicates its primary function in supplying succinate. However, the Δscsα mutant could be maintained in culture, suggesting that SSADH may compensate for succinate, albeit not sufficiently to ensure normal growth without SCSα. A double deletion of SSADH and SCSα further detrimented parasite growth but the mutant was viable in culture. We postulate that propionate-derived succinate via the 2-methylcitrate cycle²⁵ may partly fuel the TCA cycle in tachyzoites (Fig. 10).

We also found that SDHA is vital for parasite growth, and the Δsdha mutant is avirulent in mice. Our attempts to delete the FH protein, converting SDHA-derived fumarate to malate, were unsuccessful, indicating its crucial role for the growth of T. gondii. Interestingly, FH-deficient Plasmodium berghei is viable²⁶. A conditional mutant of FH in tachyzoites of T. gondii is required to define its role better. Besides PEPCK_mt¹¹, MDH-FAD likely enables the parasite to reprogram its oxaloacetate supply in varying milieus (Fig. 10). It can complement a Δmdh1 yeast mutant impaired in growth on glycerol (non-fermentable carbon source), and the Δmdh-fad tachyzoite shows a mild phenotype. The mutant accumulates malate, its NADH pool is severely compromised and the mitochondrial membrane potential collapses. Despite such metabolic anomalies, the mere survival and virulence of the Δmdh-fad mutant is unprecedented. We believe PEPCK_mt may offset the deficiency of oxaloacetate¹¹, and ATP can be supplied through glycolysis and salvage pathways. Indeed, tachyzoites express an ATP carrier (AAC1) in the mitochondrion^27,28 and can acquire host-derived ATP²⁹. However, there appears to be no substitute mechanism for maintaining the NADH and membrane potential in the Δmdh-fad mutant. The adaptive reprogramming in the Δmdh-fad strain warrants further investigation.

A second isoform of MDH is expressed in parasite cytosol, indicating the presence of a malate-aspartate shuttle for the transfer of NADH/NAD⁺ to the mitochondrion (Fig. 10). Previous work showed the myc-tagged MDH localizing to the mitochondrion of T. gondii¹⁵. However, our findings indicate that MDH-HA co-localizes with the cytoplasmic protein ALD. Consistent with our finding, the location of MDH was predicted to be in the cytoplasm by HyperLOPIT³⁰. This discrepancy in the localization of MDH can be attributed to the use of different transgenic approaches. The Δmdh mutant shows no growth phenotype, and therefore seems dispensable for malate synthesis. Tachyzoites encode a 2-oxoglutarate/malate translocase (OMT, TGGT1_274060) located in the mitochondrion-membranes³⁰, which could potentially compensate for the loss of cytosolic MDH. On the other hand, the malic enzyme, producing NADPH and pyruvate in the nucleus, is essential. NADPH is required for cellular processes, including nucleotide biosynthesis and redox balance. Likewise, acetyl-CoA synthesis from pyruvate in the nucleus is vital for histone acetylation and chromatin remodeling^21,22. Our transcriptomics data displayed that conditional knockdown of ME is associated with the modulation of replication, mismatch repair, amino sugar and nucleotide sugar metabolism. Loss of ME did not impact the total NADPH, likely due to other NADPH-generating pathways, including oxidative pentose phosphate pathway (oxPPP) and folate metabolism^31,32.

Previous research indicated that Tudor domain proteins are molecular adaptors involved in RNA metabolism, DNA damage response, and chromatin modification^33,34,35. However, it is noteworthy that we could rescue the ME-depleted strain by complementing the C-terminus and N-terminus, excluding the Tudor domain. Our data also show that the ME expression in the nucleus is regulated via the C-terminal, and its nuclear expression is essential for the lytic cycle. In mammalian cells, increasing evidence shows nuclear-mitochondrial crosstalk involving the translocation of metabolic enzymes³⁶. There is even relocation of pyruvate dehydrogenase complex to the nucleus and the occurrence of a nuclear TCA cycle regulating the metabolic-epigenetic circuitry^21,22,37. It is, therefore, tempting to surmise a role of ME-derived pyruvate in acetyl-CoA synthesis for epigenetic regulation in tachyzoites – a premise supported by our initial studies of histone acetylation. Future work shall focus on yet-uncharacterized isoforms of metabolic enzymes to study the nuclear metabolism and mito-nuclear crosstalk in T. gondii. Lastly, ME, SCSα and SDHA emerge as potential therapeutic targets against acute toxoplasmosis.

Materials and methods

Biological resources and ethics statement

The RHΔku80³⁸, RHΔku80-TiR1, RH DiCre_T2A Δku80Δhxgprt (DiCre)^39,40,41, RH_Luc and ME49 strains were provided by Bang Shen (Huazhong Agricultural University, Wuhan), respectively. Indole-3-acetic acid (IAA) was purchased from Sigma-Aldrich. Anti-Ty (mouse), anti-HSP60 (mouse), and anti-ALD (rabbit) antibodies were provided by Bang Shen (Huazhong Agricultural University, Wuhan). Anti-HA (mouse, Cat#M180-3) was purchased from MBL (Medical & Biological Laboratories Co., Japan). Rabbit anti-H3K9ac antibodies (Cat#9649) were purchased from CST (Cell Signaling Technology, USA). Alexa Fluor 488 (Cat#4408) and Alexa Fluor 594 (Cat#8889) antibodies were purchased from CST (Cell Signaling Technology, USA). Hoechst dye (Cat#94403) were purchased from Sigma-Aldrich. The female ICR mice (six-week-old) were obtained from the Guangdong Medical Experimental Animal Center (Guangdong, China). The Balb/c-nu mice were acquired from Guangzhou Ruige Biotechnology Co., LTD. We have complied with all relevant ethical regulations for animal use. Animals were raised under standard conditions approved by the Ethics Committee of South China Agricultural University (Permit 2023f266).

Cell culture

Genetically modified strains were generated using the RHΔku80³⁸, RHΔku80-TiR1 and DiCre as the parental strains^39,40,41. Parasites were propagated in human foreskin fibroblasts (HFF) (ATCC, USA) using Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 2% fetal bovine serum, 4500 mg/L glucose, 2 mM glutamine, 10 U/mL penicillin and 100 μg/mL streptomycin.

Making of transgenic parasites

Supplementary Tables S1 (Supplementary Data 1) and S2 (Supplementary Data 2) describe primers and plasmids deployed in this study. The CRISPR/Cas9 plasmid was constructed using pEASY-Basic Seamless Cloning and Assembly Kit (TransGen Biotech, China). Other plasmids were constructed using the ClonExpress II one-step cloning kit (Vazyme Biotechnology, China). For 3’-genomic tagging, we amplified the 10×HA sequence flanked by homologous arms of the target gene and co-transfected it with the matching CRISPR/Cas9 constructs into the RHΔku80 or DiCre strain. Transgenic strains were selected with 1 μM pyrimethamine (Sigma Aldrich, USA) and identified through diagnostic PCR and indirect immunofluorescence (IFA) assays. The CS1 (TGGT1_268890), SSADH (TGGT1_257480), SCSα (TGGT1_290600), SDHA (TGGT1_215590), MDH (TGGT1_318430), OMT (TGGT1_274060), AST (TGGT1_248600) and MDH-FAD (TGGT1_288500) knockout strains (Supplementary Data 3) were generated by co-transfecting YFP-DHFR* or DHFR* cassettes flanked by homologous arms of the corresponding target genes and CRISPR/Cas9 constructs into the DiCre strain. The strains were selected with 1 μM pyrimethamine, cloned by limiting dilution and clonal strains were screened by diagnostic PCRs and the YFP⁺ signal. The ΔscsαΔssadh strain was generated by replacing SSADH with Tub-CAT-mCherry via the CRISPR/Cas9 method and chloramphenicol selection (30 μM, Sigma-Aldrich, USA). The double mutant was isolated by diagnostic PCR screening and the mCherry⁺ signal.

The conditional knockdown strain for ME (TGGT1_286440) was generated by co-transfecting the locus-specific CRISPR/Cas9 construct and homology donor template into the RHΔku80-TiR1 strain (refer Supplementary Data 2), followed by selection with 25 μg/mL mycophenolic acid and 50 μg/mL xanthine. Degradation of ME in the eventual mutant (miniAID-ME) was induced by 500 μM IAA. The miniAID-ME strain was used further to generate the comp-ME, comp-ME-N/C and comp-ME-N strains by inserting the specific expression cassette into the UPRT locus (selection with 10 μM 5-fluorodeoxyuracil). Clonal strains were identified by diagnostic PCRs and IFA.

TgMDH-FAD complementation in S. cerevisiae

The empty vector (pRS416) and the indicated TgMDH-FAD construct (pRS416-MDH-FAD) were transformed into the ScΔmdh1 mutant. Briefly, the ScΔmdh1 strain was grown in 2% yeast extract, 1% tryptone, and 2% glucose. Following transformation with the plasmids described above, all yeast strains were cultured in synthetic dropout (uracil-free) minimal medium (0.67% yeast nitrogen base) supplemented with appropriate amino acids and 2% glucose. The transfectants were cloned on selective plates and tested for growth complementation on a medium with glucose or glycerin as a sole carbon source by serial dilution assay (30 °C for 3 days). The TgMDH-FAD expression cassette was introduced into chromosome XI of the ScΔmdh1. Clonal strains were selected on plates and tested for growth complementation on a medium with glucose or glycerin as a sole carbon source by serial dilution assay (30 °C for 3 days).

Production of polyclonal antibodies

The open reading frames of ME were amplified from the RHΔku80 strain and cloned into the pCold vector containing a 6×His tag. The pCold-ME construct was transformed into the BL21(DE3) strain of E. coli, and protein expression was induced by IPTG. The recombinant proteins were purified via Ni-NTA affinity resin and used to immunize female mice (six-week-old). The purified protein preparation and positive sera were aliquoted and stored at −80 °C.

Immunofluorescence assays

Based on a protocol described previously⁴², HFF cells were infected by the parental and transgenic strains and then fixed with 4% paraformaldehyde (15 min), permeabilized with 0.1% Triton X-100 (20 min) and blocked with 10% FBS (2 hours). Samples were treated with mouse anti-HA (1:1000), rabbit anti-ALD (1:1000) and rabbit anti-H3K9ac antibodies (30 min, 1:400) and stained with Alexa Fluor488 (goat anti-mouse, 1:1000), Alexa Fluor594 (goat anti-rabbit, 1:1000) IgG and Hoechst dye (30 min, 1:1000). Parasitized cells were imaged using a BX53 fluorescence microscope (Olympus, Japan) or ZEISS LSM 980 with Airyscan 2 (Carl Zeiss, Germany) and refined using ZEN software (Carl Zeiss, Germany). Image J software was used to process the fluorescent images.

Parasite phenotyping

Plaque assay were set up in 6-well plates seeded with HFF cells (100 tachyzoites/well, 3 wells per strain), essentially as reported earlier⁴³. Cultures were incubated unperturbed at 37 °C and 5% CO₂ for 6–10 days. Samples were fixed with 4% paraformaldehyde (20 min), washed by PBS, stained with 0.1% crystal violet (20 min) and scanned (Microtek Scan Marker i600, Microtek, China) to analyze the plaque size. To assess the replication efficiency, HFFs were infected with parental (RHΔku80-TiR1) or miniAID-ME strains preincubated for 2 h with or without 500 μM IAA. Cells were fixed using 4% paraformaldehyde and then incubated with mouse anti-Tg IgG (30 min). Samples were permeabilized by 0.1% Triton X-100 (20 min) and blocked with 10% FBS (2 h). Subsequently, they were incubated with rabbit anti-ALD (30 min, 1:1000), followed by Alexa Fluor 488 (goat anti-mouse, 1:1000) and Alexa Fluor 594 (goat anti-rabbit, 1:1000), and Hoechst (1:1000) was used for 30 min. The DiCre and Δmdh parasites were immunostained by mouse anti-Tg IgG (before permeabilization, 1:1000) and rabbit anti-ALD (after permeabilization, 1:1000) to visualize non-invaded and total parasites, respectively. The Δcs1, Δssadh, Δscsα, ΔscsαΔssadh, Δsdha and Δmdh-fad parasites were visualized by YFP⁺ signal and mouse anti-Tg IgG (1:1000), followed by Alexa Fluor 594 (goat anti-mouse, 1:1000). At least 100 vacuoles per sample were scored for the number of parasites developing within them.

NAD⁺/NADH measurement

Freshly isolated tachyzoites of the DiCre and Δmdh-fad strains were lysed in 200 μL of NAD⁺/NADH extraction buffer on ice (30 min) and centrifuged at 12,000 g (10 min). The total NAD⁺ and NADH levels in the supernatant were detected using an NAD⁺/NADH assay kit with WST-8 (Beyotime, Shanghai, China)⁴⁴. Briefly, a 20 μL sample or NADH standards (0–20 μM) was added to a 96-well plate. Subsequently, 90 μL of alcohol dehydrogenase was added and incubated at 37 °C (10 min), followed by the addition of 10 μL chromogenic solution (37 °C, 45 min). The absorbance values were determined at 450 nm using a plate reader (Synergy, BioTek Instruments, USA).

Parasite virulence and burden

The female ICR mice were infected with extracellular parasites of the specified strains by intraperitoneal injection (100 tachyzoites/mouse, 10 mice/strain). Infection by the Δscsα mutant was performed at doses of 10³, 10⁴ parasites (8 mice/group) and 10⁵, 10⁶ parasites (5 mice/group). The clinical signs and survival were monitored daily, and blood samples were collected from the surviving mice. Seronegative animals were not included in the data analysis^45,46. To determine the parasite proliferation, we infected female ICR or Balb/c-nu mice by intraperitoneal injection (10⁴ parasites, 5–6 mice/strain). Animals infected with the Δcs1, Δssadh, Δscsα, Δsdha, Δmdh-fad, and DiCre strains were provided with normal purified water, while those harboring the RHΔku80-TiR1 and miniAID-ME strains were given water with or without 500 μM IAA. Peritoneal fluids were collected 5 days post-infection, and genomic DNA was extracted using the TIANamp Blood DNA Kit from Tiangen Biotechnology Co., Ltd. (China). Parasites in the peritoneal fluids were detected by amplifying the non-coding fragment (529 bp, primers in Supplementary Data 1) using Power SYBR Green PCR Master Mix from Toyobo Co., Ltd. (Japan)^45,46.

Bioluminescence imaging

ICR mice were immunized with the Δscsα or Δsdha mutant (10²) via intraperitoneal injection. 34–39 days post-immunization, mice were infected with the luciferase-expressing RH-Luc strain (10⁴ tachyzoites/animal, 3 or 4 mice/group, i.p.). Naïve mice were inoculated with the same dose of the indicated strains as control groups (3 or 4 mice/group, i.p.). Animals were anesthetized (1.25% tribromoethanol, MeilunBio Co., Ltd, China) five days after infection and 300 μL of 15 mg/mL D-luciferin was injected intraperitoneally (Yeasen Biotechnology Co., Ltd, Shanghai, China). The IVIS spectrum imaging (PerkinElmer, USA) was performed to detect the parasite load⁴⁷.

Genomics and transcriptomics

The Δcs1, Δmdh and Δmdh-fad tachyzoites were purified, and genomic DNA was extracted using the TIANamp Blood DNA Kit (Tiangen Biotechnology Co., Ltd., China). The genome was then sequenced as reported previously⁴⁸. The curated datasets were compared to the reference genome of the GT1 strain (ToxoDB), and the results were visualized using the integrated genome viewer. For transcriptomics, the miniAID-ME strain was pre-treated with −/+500 μM IAA for 12 h, and parasites were purified. Total RNA was extracted using Transzol UP reagent (TransGen Biotech Co., Ltd., China). RNA sequencing was performed as described before⁴⁹. The transcriptome datasets were analyzed using the Majorbio Cloud Platform (www.majorbio.com).

Metabolomics

Metabolic labeling of extracellular tachyzoites with stable isotopes of glucose or glutamine was carried out following a modified method described previously^10,11. Tachyzoites (5 × 10⁷) of the DiCre and Δmdh-fad strains were cultured for 4 hours in a glucose-free medium containing 8 mM ¹³C₆-glucose or ¹³C₅-glutamine. Subsequently, parasites were washed with PBS and treated with 1 mL of ice-cold methanol (80%)⁵⁰. Samples were sonicated (5 cycles, 1 min each with intermittent ice cooling) and stored at −20 °C for 30 min.

Metabolites were extracted by sample centrifugation, and the supernatant was evaporated to dryness. The residue was reconstituted in 100 μL of water and then mixed with 50 μL of 175 mM 3-NPH (Sigma Aldrich, USA) in 75% methanol, 50 μL of 105 mM EDC (Sigma Aldrich, USA) in methanol and 50 μL of 2.5% pyridine (Sigma Aldrich, USA) in methanol⁴⁶. After reaction at 4 °C for 30 min, the mixtures were evaporated to dryness and re-dissolved in 50 µL of 50% methanol for the UHPLC-HRMS assay⁵¹. The chromatographic separation was performed on a Thermo Fisher Ultimate 3000 UHPLC system using a Waters HSS T3 column (2.1 mm × 100 mm, 1.7 μm) (Waters, USA). The mobile phases comprised water with 0.1% formate (phase A) and acetonitrile with 0.1% formate (phase B). Metabolites were separated by linear gradient mode (0–0.5 min, 5% B; 2.5 min, 30% B; 5.5 min, 60% B; 9–13 min, 100% B; 13.1–15 min, 5% B) and mass spectrometry data were collected using Thermo Fisher Q Exactive Hybrid Quadrupole-Orbitrap Mass Spectrometry (QE) in Heated Electrospray Ionization Negative (HESI-) mode. The spray voltage was set to 2800 V. The capillary and probe heater temperatures were adjusted to 320 °C and 350 °C, respectively. The sheath gas flow rate was 50 Arb (arbitrary unit), the auxiliary gas flow rate was tuned to 15 Arb, and the S-Lens RF level was 50 Arb. The full scan was operated at a high-resolution 7 × 10⁴ FWHM (m/z, 200) at 100–1000 m/z with an AGC target setting of 1 × 10⁶. Data were analyzed using Xcalibur software, and IsoCor v2 was used to correct the original mass spectra⁵¹.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Numerical data were plotted using GraphPad Prism 8 (GraphPad Software Inc., USA). Statistical analysis was performed using the Student’s t test, two-way ANOVA and log-rank Mantel–Cox test. The genome sequencing data are deposited and released with the accession numbers PRJNA1108415, PRJNA1109294 and PRJNA1108442. Transcriptomics sequencing data are available in the short read archive (SRA) of the National Center for Biotechnology Information database (accession number PRJNA943448). Reference genome of the Toxoplasma GT1 strain: https://toxodb.org/toxo/app/record/dataset/NCBITAXON_507601. All data are presented in the article or the supplementary information. The source data for the graphs and images can be found in the Supplementary Data 4. All resources described herein are available upon reasonable request.

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Acknowledgements

We thank Na Li, Yaqiong Guo, Dongjuan Yuan and Rui Xu (College of Veterinary Medicine, South China Agricultural University, Guangzhou) for the discussion. We thank Xianfu Gao (Shanghai Profleader Biotech Co., Ltd.) and Wenchao Wang (Phenions Biotech Co., Ltd.) for LC-MS/MS. The authors also thank the Institute of Hematology, Jinan University, for providing flow cytometric sorting. This research was supported by the National Key Research and Development Program of China (2023YFD1801000); Guangdong Major Project of Basic and Applied Basic Research (2020B0301030007); National Key Research and Development Program of China (2022YFD1800200; 2022YFD1801700); Natural Science Foundation of Guangdong Province (2022A1515011104; 2024A1515011346); 111 Project (D20008); Double first-class discipline promotion project (2023B10564003). Supplementary funding was provided via a Core Research Grant to Nishith Gupta (CRG/2021/000919) by the Department of Science and Technology – Science and Engineering Research Board (DST-SERB), India. Nishith Gupta also acknowledges the extended support of the Senior fellowship by the DBT–Wellcome Trust (India Alliance, IA/S/19/1/504263) and the Scholar Mobility Program from the Sino-German Science Center (M0074). The funders had no role in the design, data collection, analysis, preparation or decision to publish this work.

Author information

These authors contributed equally: Hongxi Zhang, Nuo Ji, Shuxin Su.

Authors and Affiliations

State Key Laboratory for Animal Disease Control and Prevention, South China Agricultural University, Guangzhou, China
Hongxi Zhang, Nuo Ji, Meng Zhao, Huiyu Du, Yaoyu Feng, Lihua Xiao & Ningbo Xia
Frontiers Science Center for Synthetic Biology and Key Laboratory of Systems Bioengineering (Ministry of Education), School of Chemical Engineering and Technology, Tianjin University, Tianjin, China
Shuxin Su & Yi Wu
Intracellular Parasite Education And Research Labs (iPEARL), Department of Biological Sciences, Birla Institute of Technology and Science, Pilani (BITS Pilani), Hyderabad, India
Lakesh Kumar Sahoo & Nishith Gupta
Guangdong Laboratory for Lingnan Modern Agriculture, Center for Emerging and Zoonotic Diseases, College of Veterinary Medicine, South China Agricultural University, Guangzhou, China
Yaoyu Feng, Lihua Xiao & Ningbo Xia
Department of Molecular Parasitology, Institute of Biology, Faculty of Life Sciences, Humboldt University, Berlin, Germany
Nishith Gupta

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Contributions

Hongxi Zhang: Resources; Data curation; Validation; Investigation; Methodology; Writing–original draft. Nuo Ji: Resources; Data curation; Validation; Investigation; Methodology; Writing–original draft. Shuxin Su: Resources; Data curation; Validation; Investigation; Methodology; Writing–original draft. Meng Zhao: Resources; Data curation; Validation; Investigation; Methodology. Huiyu Du: Resources; Data curation; Validation; Investigation; Methodology. Lakesh Kumar Sahoo: Resources; Validation; Investigation; Methodology. Yi Wu: Resources; Data curation; Validation; Investigation; Methodology; Supervision; Writing–original draft. Yaoyu Feng: Resources; Data curation; Formal analysis; Supervision; Funding acquisition; Project administration; Nishith Gupta: Resources; Data curation; Supervision; Funding acquisition; Project administration; Writing—review and editing. Lihua Xiao: Resources; Data curation; Formal analysis; Supervision; Funding acquisition; Project administration; Ningbo Xia: Conceptualization; Resources; Data curation; Software; Formal analysis; Validation; Investigation; Visualization; Methodology; Supervision; Funding acquisition; Project administration; Writing—original draft; Writing—review and editing.

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Correspondence to Nishith Gupta, Lihua Xiao or Ningbo Xia.

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Zhang, H., Ji, N., Su, S. et al. Metabolic crosstalk between the mitochondrion and the nucleus is essential for Toxoplasma gondii infection. Commun Biol 8, 384 (2025). https://doi.org/10.1038/s42003-025-07823-4

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Received: 06 August 2024
Accepted: 26 February 2025
Published: 07 March 2025
DOI: https://doi.org/10.1038/s42003-025-07823-4