# Transcript tinkering: RNA modifications in protozoan parasites

Tiziano Vignolini, Justine E.C. Couble, Gregory Doré, Sebastian Baumgarten

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Tiziano Vignolini, Justine E.C. Couble, Gregory Doré, Sebastian Baumgarten. Transcript tinkering: RNA modifications in protozoan parasites. Current Opinion in Microbiology, 2024, 79, pp.102477. 10.1016/j.mib.2024.102477. hal-05090451

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Review

# Transcript tinkering: RNA modifications in protozoan parasites

Tiziano Vignolini, Justine E.C. Double, Grégory R.G. Doré and Sebastian Baumgarten

<figure><img src="image_2.png" /></figure>

Apicomplexan and trypanosomatid parasites have evolved a wide range of post-transcriptional processes that allow them to replicate, differentiate, and transmit within and among multiple different tissue, host, and vector environments. In this review, we highlight the recent advances that point toward the regulatory potential of RNA modifications in mediating these processes on the coding and noncoding transcriptome throughout the life cycle of protozoan parasites. We discuss the recent technical advancements enabling the study of the 'epitranscriptome' and how parasites evolved RNA modification-mediated mechanisms adapted to their unique lifestyles.

**Address**

Institut Pasteur, Université Paris Cité, G5 Parasite RNA Biology, Department of Parasites and Insect Vectors, F-75015 Paris, France

Corresponding author: Baumgarten, Sebastian (sebastian.baumgarten@pasteur.fr)

**Current Opinion in Microbiology 2024, 79:102477**

This review comes from a themed issue on **Host-Microbe Interactions: Parasites**

Edited by **Mohamed-Ali Hakimi** and **Jonathan Weitzman**

For complete overview of the section, please refer to the article collection, "Host-Microbe Interactions: Parasites 2023"

Available online 24 April 2024

https://doi.org/10.1016/j.mib.2024.102477

1369-5274/© 2024 The Authors. Published by Elsevier Ltd. This is an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/).

## Introduction

The chemical modification of RNA nucleosides was observed for the first time over half a century ago [1]. Today, more than 170 distinct chemical modifications have been identified, including on mRNAs [2]. With the development of next-generation sequencing (NGS) and sensitive mass spectrometry techniques that allow for detailed localization and quantification of these RNA modifications, recent years have seen an ever-increasing interest in their regulatory potential. Although

significant advancements have been made in model eukaryotes, protozoan parasites have always been at the forefront of the field of extreme post-transcriptional regulation, ranging from RNA editing [3], RNA trans-splicing [4], rRNA fragmentation [5–10] to life stage—specific ribosome heterogeneity [11]. It is therefore not surprising that the first studies interrogating RNA modifications within this diverse group of organisms have already unveiled a plethora of highly divergent and intriguing roles for different RNA modifications. In this review, we highlight the recent advances and challenges in the study of the epitranscriptomes of apicomplexan and trypanosomatids and highlight the major knowledge gaps in these post-transcriptional regulatory mechanisms (Figure 1).

## Epitranscriptomes of protozoan parasites

Although the field of 'epitranscriptomics' has been dominated in recent years by the study of internal base modifications in protein-coding transcripts, especially N<sup>6</sup>-methyladenosine (m<sup>6</sup>A), the mechanisms used to regulate the processing, fate, or function of RNA transcripts are equally important to molecules other than mRNA. In fact, a mass spectrometry-based quantification of mRNA modifications exists only for *Plasmodium falciparum* [12,13], and m<sup>6</sup>A mRNA methylation and pseudouridine (Ψ) in *Toxoplasma gondii* and *Trypanosoma brucei* were identified by targeted approaches [14–18].

The most heavily modified RNA transcripts in many eukaryotes on the other hand are tRNAs, with 22 and 28 different modifications quantified in *P. falciparum* [19] and *T. gondii* [20,21], respectively, by mass spectrometry. rRNAs also commonly feature a diverse set of base and ribose modifications, most notably 2'-O-methylation (Nm) and Ψ. These rRNA modifications are mediated by the C/D and H/ACA class of small nucleolar RNA (snoRNA), respectively, which guide universal 2'-O-methyltransferases (2'-O-MTase) and pseudouridylases (PUS) to their rRNA target sites. Ψ and Nm on rRNA have been identified in *T. brucei* [22–25], and modification sites have also been predicted from snoRNA annotations in *P. falciparum* [26]. For *Leishmania* spp, individual rRNA modifications have been characterized [27,28] on top of a complete rRNA modification annotation generated by mass spectrometry [29]. In addition,
<figure><img src="image_3.png" /><figcaption>Figure 1</figcaption></figure>

Identified RNA modifications on tRNA, rRNA, mRNA, and small noncoding RNA in *P. falciparum* (top left), *T. gondii* (top right). *T. brucei* (bottom right) and *Leishmania spp.* (bottom left). Where known, the modification ‘writers’ are noted on the right of the modification.

the majority of rRNA modifications was also visualized in the 3D structure of the *Leishmania donovani* and *T. brucei* ribosomes generated by cryo-electron microscopy (cryo-EM) [5,30]. Importantly, for both parasites, the number of Nm is substantially higher than that in yeast, which has a similar genome size, suggesting a particular importance of these modifications for ribosome function in these parasites [27,31].

In contrast to the discovery of RNA modifications present, the identification of the diverse writer enzymes that facilitate these modifications is a more challenging task. For example, the *P. falciparum* genome encodes ~29 RNA methyltransferases alone [32], of which only four could be assigned to their corresponding RNA modification thus far [12,13,33,34]. Among the better characterized ‘writers’ in trypanosomes and *T. gondii* are PUS, including enzymes of the RNA-independent PUS family [18,21,35] and those associating with H/ACA snoRNAs [36,37]. However, as with many proteins in protozoan parasites, the high divergence in primary protein sequence and domain architecture often makes a homology-based annotation using known enzymes from model organisms unfeasible, and the individual disruption of each possible candidate remains difficult [12,18,38]. A recent example is the m⁶A machinery in *T. brucei*, where a putative regulatory binding motif necessary for m⁶A methylation was identified in the RNA sequence, but the identities of the responsible ‘writer’ proteins remain elusive [14,15]. Conversely, the *P. falciparum* tRNA m⁵C (5-methylcytosine)

methyltransferase DNMT2 was identified using orthology-based annotations and was shown to be responsible for the methylation of C38 in tRNAAsp (GTC) [33,34], where it is crucial for tRNA stability and the stress response of malaria parasites [33]. Interestingly, this protein has also been proposed to be responsible for m⁵C DNA modifications [39,40], highlighting how some protozoan enzymes might have evolved additional modes of action beyond their predicted function. Of note, while enzymes that can remove base modifications, especially m⁶A, have been identified in mammalian cells, no ‘erasers’ have so far been identified or characterized in a parasite genome (Table 1).

## From mapping to mechanism

The ability to interrogate the regulatory potential of RNA modifications has only been made possible with the concurrent development of increasingly sensitive techniques to localize, quantify, and, in some cases, even visualize RNA modifications.

Among protozoan parasites, mass spectrometry-based, global maps of RNA modifications have been conducted in *P. falciparum* [12,13,19], *T. gondii* [20], and *T. brucei* [29]. While providing absolute quantitation, mass spectrometry cannot provide specific information as to where a modified RNA nucleotide is located within a molecule. One approach to circumvent this issue is the identification of modified RNA nucleoside positional isomers from RNase T1- or A-digested RNA samples using a
<table><caption>**Table 1**

Observed phenotypes of different RNA modifications in protozoan parasites.</caption><thead><tr><th>Organism</th><th>Modification</th><th>RNA type</th><th>Observed phenotype</th><th>Reference</th></tr></thead><tbody><tr><td rowspan="10">T. brucei</td><td rowspan="10">Ψ</td><td>snoRNA</td><td>Increases efficiency of rRNA Nm</td><td>[37]</td></tr><tr><td>snRNAs</td><td>Strengthen RNA–RNA and RNA–protein interaction</td><td>[36]</td></tr><tr><td>mRNA</td><td>Growth advantage at increased temperatures</td><td>[18]</td></tr><tr><td>rRNA</td><td>Repels RBSR1, (de-)stabilizes mRNA</td><td>[23]</td></tr><tr><td>rRNA</td><td>Alters ribosome structure</td><td>[23]</td></tr><tr><td>rRNA</td><td>Growth advantage at increased temperatures</td><td>[31]</td></tr><tr><td>rRNA</td><td>Ribosomal protein recruitment</td><td>[24]</td></tr><tr><td>tRNA</td><td>Differential methylation across life cycle</td><td>[74]</td></tr><tr><td>m<sup>6</sup>A</td><td>Stabilizes VSG transcripts (in poly-A tail)</td><td>[14]</td></tr><tr><td>Wyosine</td><td>Translation fidelity (putative)</td><td>[65]</td></tr><tr><td>L. donovani</td><td>Ψ</td><td>rRNA</td><td>In vitro fitness gain</td><td>[28]</td></tr><tr><td rowspan="5">T. gondii</td><td rowspan="5">Ψ</td><td>tRNA/mRNA</td><td>mRNA destabilization</td><td>[21]</td></tr><tr><td>m<sup>6</sup>A</td><td>Inhibits tachyzoite-to-bradyzoite differentiation</td><td>[16,17]</td></tr><tr><td>m<sup>6</sup>A</td><td>mRNA stabilization</td><td>[13,66]</td></tr><tr><td>m<sup>5</sup>C</td><td>Translational repression (Reader: YTH2)</td><td>[33]</td></tr><tr><td>m<sup>5</sup>C</td><td>tRNA cleavage, decreases stress tolerance</td><td>[33]</td></tr><tr><td>P. falciparum</td><td>m<sup>5</sup>C</td><td>mRNA</td><td>mRNA stabilization, mediates gametocyte development (Reader: ALYREF (?)</td><td>[12]</td></tr></tbody></table>

VSG, variant surface glycoprotein.

two-step mass spectrometry approach. In the first step, this technique generates sequence-specific RNA fragments that have a unique mass and liquid chromatography elution profile, which can be mapped to the original sequence. Simultaneously, the RNA fragments dissociate predictably along the phosphodiesterase bond and generate a spectral map of the modified oligonucleotide within each fragment. With the sequence information that surrounds a modified base, this approach allows for the localization of the modification within an RNA molecule of interest, and it already has been used to map rRNA modifications in *Leishmania donovani* [29] and tRNA modifications in *P. falciparum* [19]. In principle, this method could therefore also be used to quantify RNA modification dynamics of a specific species in samples containing both host (or vector) and parasite material. Recently, the use of cryo-EM has advanced the field even further, especially regarding localization of rRNA modifications in ribosomes in the 3D space. This technique has provided deeper insight into the functional importance of rRNA modifications, as seen, for example, in *L. donovani* [5] and *T. brucei* [31].

The most widely used approaches to identify, quantify, and localize individual RNA modifications are NGS-based methods, which led to the surge in mRNA epi-transcriptomic studies over 10 years ago [41–43]. These methods rely either on immunoprecipitations (IP) using antibodies specific for the modification of interest (e.g. m<sup>6</sup>A) or chemical alteration of the RNA sample. IP experiments are largely analogous to chromatin IPs and can provide single-nucleotide resolution when combined with an antibody–RNA cross-linking step, as well as quantitation on a modification site, gene, and sample

level [44,45]. Chemical alteration approaches, on the other hand, rely on the pretreatment of an RNA sample. This can either lead to a conversion of a modified base to another (or protection against it), differential RNA cleavage, or the alteration of a modified base in such a way that the reverse transcriptase (RT) during cDNA generation of the sequencing library introduces a base substitution, deletion, or truncation at the modified site. These conversions and RT signatures are readily detected *in silico* and have already been applied to provide single-base resolution mapping of a range of different RNA modifications, including m<sup>5</sup>C [46], Nm [47,48], Ψ [49,50], m<sup>1</sup>A [51], ac<sup>4</sup>C [52], queuosine [53], and m<sup>6</sup>A [54–56]. Importantly, while antibody-based approaches can provide quantitative information when input, flow-through, or spike-in controls are sequenced in parallel [45], chemical alteration-based methods can give even absolute quantitation, for example, in the case of m<sup>5</sup>C, Nm, Ψ, ac<sup>4</sup>C, and m<sup>6</sup>A [46,48,49,52,54] when near-complete conversion/protection of a modified base is achieved. Thus, for parasite research comparing RNA modification dynamics between life cycle stages, the choice of mapping technique has therefore to be carefully considered.

NGS-based methods have recently been joined by third-generation sequencing methods, such as Oxford Nanopore direct RNA sequencing (DRS). With DRS, native RNA is pulled through a nanopore embedded in a membrane across which a flow of ions is established. The time that each stretch of bases resides within a pore and the change in the ion flow across the membrane can first be used to identify each base. Sites of RNA modifications are then derived based either on base-calling
errors or from the raw signal space (i.e. changes in
membrane current). However, the identity of an RNA
modification depends on (1) a training set generated *a priori* [57–59], (2) comparison of a native RNA to either a
sample without (i.e. an *in vitro* transcribed RNA) or with
reduced levels of an RNA modification (i.e. a ‘writer’
knock-down/knock-out sample) [60,61], or (3) a combi-
nation of both *a priori* and comparative approaches [62].
The reliance on a writer knock-down makes DRS chal-
lenging when the writer is unknown or genetic mod-
ification is difficult, as is the case in many protozoan
parasites. However, once RNA modification sites of in-
terest have been identified with an orthogonal NGS-
based method, DRS library preparations are often sub-
stantially easier than their NGS counterparts and, in
combination with long sequencing reads, are even able
to detect interdependencies of RNA modifications on
the same molecule [63].

NGS-based methods have been applied broadly in pro-
tozoan parasites for the detection of m⁶A [13], m⁵C [12],
and Ψ [18,37], and DRS has been used to identify m⁶A
sites in *T. gondii* [17] and Ψ in *T. brucei* [31]. Importantly,
the latter study compared the performance of multiple
orthogonal NGS approaches and DRS and showed that
technical limitations unique to each method can lead to
small discrepancies when quantifying the dynamics of Ψ
[31]. Similarly, a recent comparison of DRS and NGS
methods to identify m⁶A sites in human cells also
showed that both among NGS methods and between
NGS and DRS methods, substantially different numbers
of m⁶A sites are identified with overall little overlap [59].
This further highlights the importance of using multiple
orthogonal approaches to accurately identify the location
and dynamics of RNA modifications.

The only limiting factor in some NGS (i.e. especially
when antibody based) and DRS methods for protozoan
parasites is the required input quantity, which can pose a
particular challenge when working with certain low
abundance life cycle stages or even patient samples.
However, the latest NGS methods can detect mod-
ifications with input material in the range of picograms of
RNA [64], and in many assays, the multiplexing of
samples and replicates can further decrease the initial
input requirement and technical variability [45].

Special lifestyles with special mechanisms

One of the most comprehensive functional character-
izations of a single RNA modification and its effects on a
parasite transcriptome was performed in *T. brucei* [31].
Here, a single Ψ-guiding H/ACA snoRNA was deleted
that usually deposits Ψ530 in helix h69 of the ribosome
large subunit. Absence of Ψ530 changed the composition
of the ribosome, namely, the dislodging of the ribosomal
protein eS12, which, in turn, led to the differential

translation of stage-specific proteins. Although the au-
thors found that Ψ530 is not differentially regulated
across different life cycle stages of *T. brucei*, this study
convincingly demonstrated how ribosome heterogeneity
and specialization can emerge and be regulated by a
single rRNA modification. In addition, given the dy-
namics of many RNA modifications across the life cycle
of *T. brucei* and other parasites, this study provided a
proof of concept for the functionality of these dynamics
and their post-transcriptional and translational regulatory
potential.

Among tRNA modifications, a notable example is the
presence of wyosine. While this modification can also be
found in cytoplasmic tRNAs of eukaryotes and archaea,
its presence in mitochondrial tRNA is unique among
eukaryotes. One possible explanation for this could be
that wyosine serves the maintenance of translation fi-
delity in the U-rich mitochondrial transcriptome that
results from extensive RNA editing [65].

With m⁶A being the most abundant internal base mod-
ification of mRNAs in model systems, it has also taken
the center stage in protozoan mRNA epitranscriptome
studies. Indeed, in all three species investigated, m⁶A
has immediately shown its versatility by mediating reg-
ulatory processes not seen in any model organism in-
vestigated before. *T. brucei* once again served as a model
to identify a novel post-transcriptional mechanism, re-
vealing the potential for extensive m⁶A methylation in
poly-A tails of mRNA transcripts [14]. Interestingly, al-
though m⁶A occurs throughout the protein-coding tran-
scriptome of this parasite [15], most m⁶A was found in
the poly-A tail of transcripts encoding variant surface
antigens, where it serves as an mRNA-stabilizing factor
and protects the transcript from degradation [14].

In most cases, the effect of mRNA base modifications
has been found to be mediated by specific ‘reader’
proteins that can translate the modification into different
functional outcomes. Both the *P. falciparum* and *T. gondii*
genomes encode for two proteins of the YT521-B
homology protein (YTH) family, which represent con-
served m⁶A ‘readers’ among eukaryotes [13,16,17,66,67].
YTH1 in *T. gondii* is a homolog of the 30 kDA subunit of
the cleavage and polyadenylation specificity factor
(CPSF4) [68], and its m⁶A binding activity was con-
firmed recently [16,17]. Knock-down of this protein led
to aberrant 3' end processing and subsequent read-
through transcription, providing the first link between
m⁶A methylation and 3' processing. Intriguingly, the
YTH1 domain architecture of *T. gondii* and *P. falciparum*
is shared with that of green plants, which show a similar
defective 3' end processing phenotype when the corre-
sponding protein is knocked down [69]. This suggests a
potential functional convergence of these distant phyla.
The YTH2 protein of *P. falciparum*, on the other hand,
shows no resemblance to any other known YTH proteins, and a functional characterization of this protein suggested that it acts as a translational repressor [13,66]. As this phenotype has yet to be observed in other eu-karyotes, it remains unknown how exactly YTH2 can effect this regulation [66]. Beyond m⁶A, m5C mRNA modifications have been identified in *P. falciparum*. Although a putative m⁵C 'reader' protein is still unknown, disruption of the NSUN2 'writer' revealed that m⁵C is important for RNA stability and is involved in regulating the development of the host-to-vector transmission stage [12].

Importantly, characterizing individual 'reader' proteins of mRNA modifications removes the potential bias that arises from disrupting a 'writer' enzyme whose modification can be recognized by multiple 'readers' and generating overlapping effects that are difficult to disentangle afterward. In addition, some specific 'readers' might also be able to recruit secondary effectors, such as mammalian YTHDC1 and proteins of the splicing factor family serine/arginine-rich splicing factor (SRSF) [70]. On the other hand, for some modifications such as m⁶A, it has also been shown that they can affect RNA-binding proteins that are not specific modification 'readers'. Here, by unfolding of the RNA secondary structure, m⁶A can (un-)mask binding sites of these proteins ('m⁶A-switch') [71]. As of today, an untargeted mass spectrometry-based approach to identify specific modification 'readers' has only been performed in *P. falciparum*. Of the 10 proteins that preferentially bind m⁶A, all of them still require orthogonal and functional validation [66].

## Dynamics of RNA modifications across parasite life cycles

The dynamic nature of RNA modifications makes their regulatory potential especially intriguing within the context of the vast changes that take place across the life cycle of a parasite. While no RNA modification erasers have been identified in any protozoan parasite yet, differential expression of the m⁶A 'writer' enzymes, as in *P. falciparum*, gives rise to highly dynamic m⁶A levels during parasite development [13]. Beyond the m⁶A sites in the poly-A mentioned above, largely differing numbers of m⁶A-methylated transcripts were also identified when comparing mRNA transcriptomes across the *T. brucei* life cycle. Here, most transcripts were found to be methylated in the host-to-vector transmission stage, opening the possibility for a role of m⁶A in mediating the parasite's transmission [72].

An additional layer of regulation is timing. The presence or absence of modification 'readers' can define how exactly a modification marks mRNA transcripts for different

fates [13,16,17]. For example, the *P. falciparum* m⁶A-reader YTH2 is expressed at its highest levels in the two transmission stages, suggesting that it serves a specific function in these developmental stages [13]. Dynamics of specific tRNA modifications during blood stage development of *P. falciparum* have also led to the hypothesis of a 'code of codons'. Here, an increase of wobble modifications in tRNA coincides with increased translation of proteins featuring a bias for codons read by these modified tRNA isoacceptors. Lowly abundant proteins, on the other hand, are enriched in synonymous codons translated by nonwobble-modified tRNAs [19]. Similar developmental dynamics of RNA modifications have also been observed in *T. brucei*, where the total number of Ψ and Nm in mRNAs and noncoding RNAs remains largely stable, but the expression of guiding snoRNAs and the level of modifications is highly dynamic, leading to hyper-or hypo-modifications depending on the life cycle stage or growth condition [23,24,36,37].

On top of these developmental regulations, the dynamics of RNA modification can also be influenced by the environment of a cell. For protozoan parasites, this is particularly important, given how they can survive and differentiate within a considerable range of different environments dictated by their hosts and vectors. An extremely interesting example is the nucleoside queuosine (Q), a wobble modification found on position 34 of a subset of tRNAs. Importantly, the nucleobase queuine must be foraged from the environment since only prokaryotes can synthesize it de novo. Therefore, *T. brucei* parasites alter the levels of Q on some tRNA depending on the nutritional status of their host [73]. More curious still is the fact that Q-modified tRNAs are preferentially imported into the mitochondria of *T. brucei* [74]. Once imported, they are important for the efficient translation of the U-rich RNAs that result from RNA editing [74]. These findings reveal a single tRNA modification to be at the center of a putative nutrient-sensing mechanism and the metabolic response of the parasite.

Increasing evidence also points toward a more direct effect of the environment on the RNA modification landscape. In the hyperthermophilic archaea *Thermococcus kodakarensis*, ac⁴C levels on rRNA increase proportionally with temperature, helping to stabilize the ribosome at high temperatures [52]. First indications for a similar mechanism have already been observed in *T. brucei*, where increased snoRNA expression and rRNA pseudouridylation can potentially alter the ribosome structure, leading to increased parasite growth at elevated temperatures [23]. For malaria parasites, such a dynamic rRNA epitranscriptome could give rise to an even more complex regulation of protein synthesis, considering how these parasites already transcribe highly divergent sets of rRNAs depending on whether they are in the host or vector [11].
## Future directions

The first detailed studies of phenotypes and mechanisms mediated by RNA modifications already highlight their regulatory potential throughout the life cycles of many different parasites. Research on these processes would benefit greatly from the advent of more high-throughput, targeted screening methods of gene function [75-79], which could help to identify which enzyme (or snoRNA) is responsible for each individual RNA modification. The identified 'epitranscriptomes' of parasites are extraordinary with regard to how the parasites utilize these modifications to mediate different post-transcriptional processes specific to their unique lifestyles. Characterizing the underlying mechanisms could ultimately help to identify novel Achilles' heels to combat the spread of the diseases these parasites cause. The study of how parasite 'epitranscriptomes' respond to and/or are shaped by their environment will provide exciting new avenues for developing interventions. Importantly, the ability of apicomplexan and trypanosomid parasites to cycle among many different cells, tissues, hosts, and vectors gives them remarkable potential as model systems for elucidating how RNA modifications translate environmental signals into a functional molecular outcome.

## CRediT authorship contribution statement

TV: Writing – original draft; JECC: Writing – original draft; GRGD: Writing – original draft; SB: Funding acquisition, Conceptualization, Writing – original draft.

## Data Availability

No data were used for the research described in the article.

## Declaration of Competing Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Acknowledgements

The authors would like to thank Dr. Jessica M. Bryant and Dr. Ameya Sinha for the discussions during manuscript preparation. The work in the Parasite RNA Biology Group is supported by the European Commission (ERC Staring Grant 'PlasmoEpiRNA', #947819) and Institut Pasteur baseline funding. Graphics created with BioRender.com.

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