
==== Front
J Exp Bot
J Exp Bot
exbotj
Journal of Experimental Botany
0022-0957
1460-2431
Oxford University Press UK

38592734
10.1093/jxb/erae151
erae151
Review Papers
AcademicSubjects/SCI01210
Experimental approaches to studying translation in plant semi-autonomous organelles
https://orcid.org/0000-0002-1966-9455
Kwasniak-Owczarek Malgorzata Department of Cellular Molecular Biology, Faculty of Biotechnology, University of Wroclaw, F. Joliot-Curie 14A, Wroclaw, 50-383, Poland

https://orcid.org/0000-0002-6542-343X
Janska Hanna Department of Cellular Molecular Biology, Faculty of Biotechnology, University of Wroclaw, F. Joliot-Curie 14A, Wroclaw, 50-383, Poland

Janda Martin University of South Bohemia in České Budějovice, Czech Republic
Editor
Correspondence: malgorzata.kwasniak-owczarek@uwr.edu.pl
11 9 2024
09 4 2024
09 4 2024
75 17 51755187
18 1 2024
04 4 2024
08 4 2024
27 4 2024
© The Author(s) 2024. Published by Oxford University Press on behalf of the Society for Experimental Biology.
2024
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Abstract

Plant mitochondria and chloroplasts are semi-autonomous organelles originated from free-living bacteria that have retained reduced genomes during evolution. As a consequence, relatively few of the mitochondrial and chloroplast proteins are encoded in the organellar genomes and synthesized by the organellar ribosomes. Since both organellar genomes encode mainly components of the energy transduction systems, oxidative phosphorylation in mitochondria and photosynthetic apparatus in chloroplasts, understanding organellar translation is critical for a thorough comprehension of key aspects of mitochondrial and chloroplast activity affecting plant growth and development. Recent studies have clearly shown that translation is a key regulatory node in the expression of plant organellar genes, underscoring the need for an adequate methodology to study this unique stage of gene expression. The organellar translatome can be analysed by studying newly synthesized proteins or the mRNA pool recruited to the organellar ribosomes. In this review, we present experimental approaches used for studying translation in plant bioenergetic organelles. Their benefits and limitations, as well as the critical steps, are discussed. Additionally, we briefly mention several recently developed strategies to study organellar translation that have not yet been applied to plants.

A review outlining the distinctive characteristics of the methods currently used for examining translation in plant organelles along with plausible future methodological perspectives in plant organellar translation studies.

Arabidopsis
chloroplast
mitochondria
plant
polysome profiling
protein synthesis
radioisotope labeling
ribosome
ribosome profiling
translation assay
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pmcIntroduction

The plant cell contains two types of bioenergetic organelles of bacterial origin, mitochondria and chloroplasts, each with its own DNA and a gene expression system. Compared with bacterial genomes, organellar genomes are significantly smaller and encode just a tiny fraction of the organellar proteins. Only about 50 of the approximately 2000–3000 proteins found in mitochondria are encoded by the mitochondrial genome, while the chloroplast genome encodes ~80 of the 3000 chloroplast proteins (van Wijk, 2015; Zoschke and Bock, 2018; Møller et al., 2021). The remaining proteins are encoded by nuclear genes, synthesized in the cytosol, and imported to the organelles. Thus, the biogenesis of both mitochondria and chloroplasts requires a coordinated expression of the organellar and nuclear genes. The mitochondrial genome encodes mainly components of oxidative phosphorylation complexes (OXPHOS) and ribosomal proteins, whereas the chloroplast genome encodes mostly proteins involved in photosynthesis and gene expression. Due to the key role of those proteins in the organellar energy transducing systems, their efficient expression is critical for plant growth and development. It has been shown that the expression of the plant organellar genomes is regulated mainly post-transcriptionally (Hammani and Giegé, 2014; Sun and Zerges, 2015), the last stage of the expression, translation, being particularly important in this respect.

The protein synthesis within mitochondria and chloroplasts is carried out by mitoribosomes and chlororibosomes. Similarly to their cytosolic and bacterial counterparts, they are composed of a small subunit (SSU) that binds mRNA and decodes the genetic information and a large subunit (LSU) that catalyses the synthesis of polypeptide chains. Due to their evolutionary origins, the organellar ribosomes were long believed to be closely related to prokaryotic ones. While this is true for the chlororibosomes (Bieri et al., 2017; Perez Boerema et al., 2018), which strongly resemble bacterial ribosomes, the mitoribosomes have in fact diverged markedly from the bacterial ancestor. This divergence resulted from the acquisition of a large number of proteins and the expansion of large rRNA segments during evolution (Tomal et al., 2019). While the Escherichia coli ribosome contains 54 proteins (Chen and Williamson, 2013), and spinach chlororibosomes a total of 57 proteins (Bieri et al., 2017), as many as 94 proteins have been found in the Arabidopsis mitoribosome (Waltz et al., 2019). Also the chloroplast rRNAs are similar in length to their E. coli counterparts (Bieri et al., 2017), while the mitochondrial rRNAs are longer (Waltz et al., 2019). Notably, the mitochondrial 18S rRNA, the scaffolding for the SSU mitoribosomal proteins, carries a large insertion in the 3ʹ domain that binds plant-specific mitoribosomal proteins such as ribosomal pentatricopeptide repeat proteins (rPPRs) and/or translation factors (Tomal et al., 2019). Both the acquisition of the plant-specific proteins and the expanded rRNAs are reflected in the unexpected large size of the Arabidopsis mtSSU, larger than the mtLSU (Waltz et al., 2019). Another feature distinguishing mitoribosomes from the bacterial and chloroplastic ribosomes is the absence of the anti-Shine–Dalgarno (anti-SD) sequence at the 3ʹ end of the 18S rRNA, which facilitates the ribosome binding to mRNA (Hammani and Giegé, 2014). This absence indicates that the initiation of translation in mitochondria differs markedly from that in bacteria and chloroplasts (Zoschke and Bock, 2018). Indeed, a plant-specific mechanism of mitochondrial translation initiation has been identified recently in which protein–mRNA rather than an rRNA–mRNA interactions mediate ribosome binding (Tran et al., 2023). During translation the ribosomes ensure the correct positioning of the mRNA codons to base-pair with anticodons of the transfer RNAs (tRNAs), which carry the amino acids and are the real ‘translators’ of genetic information. Although chloroplast translation systems use a complete set of tRNAs that are encoded by chloroplast genes (Zoschke and Bock, 2018), the situation is more complex in mitochondria. tRNAs present in plant mitochondria have different genetic origins. The pool of mitochondrial tRNAs includes tRNAs transcribed from native mitochondrial genes, intracellularly transferred genes from chloroplasts, and horizontally transferred genes from other species, as well as tRNAs expressed from nuclear genes and imported from the cytosol (Warren and Sloan, 2020). The study of tRNA expression by sequencing methods is hampered by the secondary structure and post-transcriptional modifications of tRNAs. However, recently, a modified high-throughput tRNA sequencing (tRNA-seq) analysis in plants resulted in detecting higher abundance and diversity of tRNA reads, including organellar tRNAs (Warren et al., 2021).

In this paper, we present current methods for studying translation in plant organelles. These experimental approaches determine translational activity either directly, by measuring the accumulation of newly synthesized proteins, or indirectly, by examining the ribosome coverage of mRNA. We outline the principles and critical points of each approach as well as their benefits, limitations, and optimal applications, and present representative recent discoveries made using these techniques. Finally, we describe some newly developed methods potentially applicable to studying organellar translation in plants.

An overview of approaches used to study translation in plant mitochondria and chloroplasts

Approaches based on detection of newly synthesized proteins

The most direct way to study translation is the detection of its output—the newly synthesized proteins. In this approach metabolic labeling (e.g. radioisotope labeling) is used to incorporate detection tags into nascent proteins in the native translation system of interest.

Radioisotope labeling assays

Isolated organelles, cells, or intact plant tissues are incubated with radioactively labeled amino acids for a period of time, allowing their incorporation into newly synthesized proteins proportional to their synthesis rate (Barkan, 1998). The radiolabeled polypeptides are visualized by autoradiography or phosphorimaging and the relative intensity of signals reflects the rates of synthesis of individual polypeptides.

Such an assay on isolated organelles, commonly referred to as in organello translation analysis, takes advantage of the fact that both mitochondria and chloroplasts retain the ability to synthesize proteins after their isolation. The synthesis of radiolabeled proteins by isolated plant mitochondria was first studied by Christopher J. Leaver’s group (Forde et al., 1978; Forde and Leaver, 1980); they showed that mitochondria from three cytoplasmic male-sterile lines of maize synthesized specific variant polypeptides unique to each line and different from those from lines with wild-type cytoplasm. The in organello assay was later applied to study mitochondrial translation products in other plants, such as sorghum, tobacco, and artichoke (Forde et al., 1978; Dixon and Leaver, 1982; Håkansson et al., 1988). The radiolabeling assay in chloroplasts was performed for the first time by John Ellis and his coworkers to demonstrate that isolated pea and spinach chloroplasts synthesized discrete proteins under the influence of light (Blair and Ellis, 1973; Eaglesham and Ellis, 1974). Those ground-breaking studies laid the foundation for the currently used optimized assays in Arabidopsis organelles (Kwasniak et al., 2013; Karpinska et al., 2022; Kwasniak-Owczarek et al., 2022).

To preserve their protein synthesis potential the organelles have to be isolated quickly and gently. Since a prolonged storage of isolated organelles compromises their bioenergetic capacity, it is recommended to perform in organello translation directly after isolation. Protein synthesis in isolated mitochondria is carried out in the presence of ADP and an oxidizable substrate (succinate, malate, pyruvate) to allow ATP to be replenished by oxidative phosphorylation (Forde et al., 1978; Lind et al., 1991) or with a direct supplementation of ATP (Grohmann, 1995). In contrast, the protein synthesis in isolated chloroplasts requires no external chemical energy source since the triphosphonucleosides required can be regenerated via light-dependent processes (Gnanam et al., 1988). However, it has been shown that the chloroplast protein synthesis system is also functional in the dark if ATP is supplied, although with a lower efficiency compared with the light driven synthesis (Siddell and Ellis, 1975). Notably, when Mg2+ was supplied as well, the ATP-driven protein synthesis in the dark proceeded at an even greater rate than that driven by light (Fish et al., 1983).

Protein synthesis in isolated organelles is performed in a suitable medium containing a mixture of unlabeled amino acids, and protein labeling is started by the addition of a radioactive amino acid, such as [35S]methionine, which competes with the ‘cold’ methionine for incorporation into nascent polypeptides (Zhang et al., 2000; Kwasniak-Owczarek and Janska, 2014). It should be emphasized that the incorporation of amino acids is linear for a certain period of time and then slowly plateaus upon reaching a balance between protein synthesis and degradation. Thus, a relatively short translation period only should be allowed to minimize the influence of degradation of the newly synthesized proteins. A maximum of 60 min of pulse-labeling has been found to be optimal for intact mitochondria and chloroplasts (Karpinska et al., 2022; Kwasniak-Owczarek et al., 2022).

The isolated organelles for a radiolabeling assay should be free of bacteria, which could also take up amino acids from the medium and synthesize their own proteins. To identify translation products derived from a possible bacterial contamination, a control reaction is recommended with the oxidizable substrate–ADP mix replaced with sodium acetate, a non-oxidizable substrate that can be metabolized by bacteria (Leaver et al., 1983; Bhadula and Shargool, 1991). Additionally, to identify translation products of cytosolic ribosomes that could potentially contaminate isolated organelles, it is recommended to perform the translation assay in the presence of a specific inhibitor of cytoplasmic translation such as cycloheximide (Mulo et al., 2003). To confirm conclusively the mitochondrial or chloroplastic origin of the labeled polypeptides, the assay should be run in the presence of prokaryotic-type translation inhibitors. Chloramphenicol blocks protein synthesis both in mitochondria and chloroplasts, while lincomycin and erythromycin are specific inhibitors of chloroplast translation (Mulo et al., 2003).

Radiolabeling of translation products can also be performed in intact plant tissues. Such in vivo radiolabeling has never been reported for mitochondrial proteins, but is frequently used to assess the rate of protein synthesis in chloroplasts (Pesaresi et al., 2001, 2006). Thanks to the high efficiency of in vivo labeling of chloroplast proteins, only a few leaf discs vacuum-infiltrated in a suitable radiolabeling medium are required to obtain a strong signal (Pesaresi, 2011). When cytosolic translation is prevented with cycloheximide, only proteins synthesized in organelles are labeled. Immediately, after the infiltration the discs are illuminated for different periods of time to monitor the progress of protein synthesis (at least 15 min is required to obtain visible labeling) (Pesaresi, 2011). Following the labeling, total chloroplast or stroma- and thylakoid-enriched protein fractions are obtained and analysed. The main disadvantage of this method is that for low-abundance proteins with a high turnover rate the 15 min labeling time could be too long as such proteins would become degraded during the labeling (Barkan, 1998). In such a case, another assay, such as in organello labeling, is recommended.

The radiolabeled polypeptides are resolved by SDS-PAGE electrophoresis and then detected by autoradiography or phosphorimaging. A global analysis of the labeled polypeptides offers little information, and a deeper insight into, for example, the effects of mutations or treatments requires individual protein species to be identified. To the best of our knowledge mass spectrometry has not yet been applied to in organello-synthesized polypeptides, and only indirect methods have been used for tentative identification of individual bands. In most cases a simple comparison of molecular masses determined from electrophoretic mobility of labeled proteins with those calculated from the amino acid composition of known organellar proteins has been used. Also patterns of synthesized polypeptides already known from the literature can aid the protein identification (Giegé et al., 2005; Pesaresi et al., 2006). In this way, several of Arabidopsis OXPHOS proteins have been putatively recognized, including ATP synthase subunits ATP1 and ATP9, NADH:ubiquinone oxidoreductase (NAD) subunits 7 and 9, cytochrome b (COB), and cytochrome c oxidase subunit II (COXII) (Giegé et al., 2005; Kwasniak et al., 2013; Karpinska et al., 2022). Among the soluble chloroplast proteins of Arabidopsis, the band representing the large subunit of ribulose-1,5-bisphosphate carboxylase/oxygenase (RBCL) has been identified, and among thylakoid membrane proteins those corresponding to the D1 and D2 core proteins from photosystem II (Pesaresi, 2011; Karpinska et al., 2022). Another approach based on immunological analysis with specific antibodies was used to identify the NAD9 subunit from potato mitochondria (Grohmann, 1995), as well as to identify the ATP1, COB, COXII, COXIII, and ATP9 subunits from Arabidopsis mitochondria (Kolli et al., 2019). We stress again that these methods offer a provisional identification only, and an unambiguous identification of a given polypeptide/band requires a direct experimental verification.

Assays based on interactions of mRNA with ribosome

A different set of strategies is currently used to study organellar translation target transcripts that are associated with ribosomes. Efficiently translated mRNAs are associated with several ribosomes, forming polysomes, whereas poorly translated mRNAs are associated with a single ribosome only (monosomes). The strategies in question comprise polysome profiling and ribosome-profiling (Ribo-seq). They both assume that all mRNAs bound by a ribosome undergo translation and, provided the elongation rate is constant across the translatome, the ribosome occupancy of a given mRNA species can serve as a proxy for its translation efficiency.

Polysome profiling assay

The assay is based on the separation of complexes of mRNA with ribosomes depending on the number of ribosomes bound using sucrose density gradient centrifugation. While the most intensive development of polysome isolation protocols in plant systems occurred in the 1960 and 1970s (Watts and Mathias, 1967; Davies et al., 1972; White and Murakishi, 1977), the first applications of polysome profiling to plant mitochondrial or chloroplast mRNA were only published in the late 1990s (Lu and Hanson, 1994; Barkan, 1998).

To be biologically relevant, polysome profiling requires that the polysomes be isolated with high yield and well-preserved integrity. A total plant lysate is obtained first by grinding tissues in a buffer ensuring stability of the ribosome–mRNA complexes. Polysomes are stable at high pH and at high concentration of Tris and KCl. Addition of EGTA to chelate divalent cations (Ca2+, Cu2+, Zn2+) accumulated in leaf tissue, and Mg2+ to prevent dissociation of ribosomal subunits, improves the yield of polysomes provided the concentration of Mg2+ exceeds that of EGTA (Jackson and Larkins, 1976; White and Murakishi, 1977). To extract membrane-bound polysomes, detergents are added to the extraction buffer (Kwasniak et al., 2013; Hameed et al., 2017). To minimize the mobility of ribosomes on cytosolic and organellar mRNA, suitable translation inhibitors are added, cycloheximide and chloramphenicol, respectively (Lecampion et al., 2016). Heparin and/or specific RNase inhibitors are used to preserve mRNA integrity and an anti-protease cocktail to prevent protein degradation. The clarified lysate is loaded onto a sucrose gradient and separated by ultracentrifugation, during which the complexes sediment down the gradient according to their sedimentation coefficients. Thus, large polysomes sediment the farthest followed by monosomes, free ribosome subunits, and finally free mRNA. After the ultracentrifugation, the gradient is collected, usually from bottom to top, in several fractions of equal volume. Monitoring the absorbance at 254 nm (in a continuous manner during the fractionation, using a flow-through recording spectrophotometer, or in collected fractions individually) gives an absorbance profile with typical peaks. While the unloading of the gradient is usually performed from bottom to top, it is conceptually more straightforward to describe the obtained profile pattern in the reverse order (i.e. from top to bottom). Thus, the first (topmost) peak corresponds to free mRNA unbound to ribosomes, the second and third peaks to ribosome SSU and LSU, and the fourth and subsequent peaks to monosomes and polysomes with increasing ribosome loading (Lecampion et al., 2016). By determining the area under individual peaks one can estimate the relative amounts of mRNA bound to monosomes and polysomes; the polysome/monosome ratio reflects the overall translational activity. Subsequently, mRNA and proteins are extracted from each fraction separately and analysed by RT–qPCR or northern blotting with mRNA-specific probes, and by western blotting.

For such analyses to be biologically relevant, one has to verify that the mRNAs present in the polysome fractions are indeed associated with translating ribosomes. Also mRNA not undergoing translation can be associated with RNA-binding proteins or with several stalled ribosomes, and such complexes can fortuitously co-sediment with polysomes (Chassé et al., 2017). Thus, to differentiate between translationally active polysomes and other mRNA complexes, additional assays should be performed including diverse agents leading to the destabilization of polysomes, for example treatment with EDTA, RNase, or puromycin (Uyttewaal et al., 2008; Hameed et al., 2017). EDTA and RNase destabilize ribosomes by chelating Mg2+ ions or degrading RNA necessary for their integrity, whereas puromycin specifically destabilizes ribosomes in the elongation step of translation and only affects active polysomes. Thus, puromycin-containing gradients are recommended for identification of fractions containing polysomes and devoid of free mRNA or ribosomes (Kahlau and Bock, 2008).

Polysome profiling allows two parameters related to mRNA translatability to be determined: ribosomal loading and ribosomal density (Kwasniak et al., 2013). Ribosomal loading (or ribosome occupancy) represents the proportion of ribosome-bound mRNA relative to total mRNA in polysomal and non-polysomal fractions and is used as a proxy of translational efficiency. The second indicator of mRNA translability is ribosomal density, which defines the average number of ribosomes present on a transcript reflected in the mRNA distribution across the polysome fraction. Such distribution can be expressed as a percentage of total RNA present in each fraction and compared between different samples/conditions. Shifts of mRNA abundance maxima to lighter or heavier fractions indicate that the number of ribosomes on the mRNA is lower or higher, and thus show whether the mRNA is poorly or highly translated.

Polysome profiling can be used for targeted analyses of specific organellar mRNAs, but it can also be employed to investigate mRNA–ribosome interactions at the whole transcriptome level using microarray (Hameed et al., 2017) or RNA-seq (Grimes et al., 2014) approaches, as well as to identify ribosomal and ribosome-associated proteins using mass spectrometry (Firmino et al., 2020). Notably, polysome profiling is applicable not only to the plant species with annotated genomes but also to those whose genome remains unannotated (Goldenkova-Pavlova et al., 2018).

Polysome profiling has been widely used to determine the translational status of organellar mRNA in various plant tissues and species (Lecampion et al., 2016). This assay has been utilized, among others, to compare the translational status of chloroplast mRNAs in the prpl11 mutant with a chloroplast ribosomal protein L11 knockout (Pesaresi et al., 2001) and of mitochondrial mRNAs in the rps10 mutant deficient in mitoribosomal protein S10 (Kwasniak et al., 2013) with those in wild-type Arabidopsis. Interestingly, the analysis of ribosomal occupancy and density in the rps10 mutant revealed marked differences of the translational status between two classes of mitochondrial mRNAs: those encoding OXPHOS subunits and mitoribosomal ones (Kwasniak et al., 2013). Polysome analysis was also used to analyse the binding of chloroplast ribosomes with the psbD mRNA to determine the impact of Arabidopsis transcription factors CIA2 and CIL on translation in chloroplasts (Gawronski et al., 2020). In plants lacking these factors, psbD mRNA was associated with fewer ribosomes than in the wild-type ones, indicating reduced chloroplast translation. More examples of the use of polysome profiling include comprehensive analyses of the chloroplast translatome changes during fruit development and the chloroplast-to-chromoplast conversion in tomato (Kahlau and Bock, 2008) and of the association of mitochondrial mRNAs with ribosomes in different tobacco tissues (Hameed et al., 2017).

Despite its wide use to study plant organellar translation, polysome profiling has certain limitations, including a limited ability of the sucrose gradient to resolve higher-order polysomes (e.g. trisomes from tetrasomes) (Mazzoni-Putman and Stepanova, 2018). Consequently, the ribosome density cannot be determined exactly since the mRNA-ribosome complexes from a given gradient fraction may contain different numbers of ribosomes. Polysome profiling also cannot determine the actual ribosome distribution along the transcript, which makes it impossible to identify the mRNA regulatory elements important for efficient translation (Goldenkova-Pavlova et al., 2018). Moreover, it does not allow distinguishing mRNAs loaded with actively elongating ribosomes from those with paused or stalled ribosomes (they have similar molecular mass), and therefore detection of translational regulation is possible only at the level of initiation (Zoschke and Bock, 2018). Finally, the interpretation of polysome profiling experiments on plant organelles is often complicated since some of their genes, especially chloroplast ones, can be transcribed as mono-, di-, and polycistronic units, all capable of being translated (Zoschke and Bock, 2018; Best et al., 2020, Preprint). Since the reading frames located on a single transcript are very close to each other or may even overlap, their individual translation rates cannot be precisely defined.

Ribosome profiling assay

The ribosome profiling assay, also referred to as ribosome sequencing (Ribo-seq), is based on the fact that ribosome-bound mRNA regions are protected against ribonuclease action (Steitz, 1969). The ribosome-protected mRNA fragments, also called ribosome footprints (RFs), are converted to a cDNA library, which is then subjected to deep sequencing or analysed by microarray hybridization. In contrast to other translatome techniques, Ribo-seq allows in vivo determination not only of the relative number of translating ribosomes, but also of their exact location along the mRNA (Ingolia et al., 2009).

Ribosome profiling was developed for yeast by the team of Jonathan Weissman and Nicholas Ingolia (Ingolia et al., 2009) and first applied to study plant organellar translation in 2013, when Zoschke et al. (2013) investigated protein synthesis in maize chloroplasts. Zoschke et al (2013) used a modified assay, exchanging deep-sequencing analysis of RFs by microarray hybridization. They argued that the key benefits of using microarrays instead of deep sequencing were time and cost saving. Later the ribosome profiling method based on deep sequencing of RFs was used to characterize the chloroplast (Lukoszek et al., 2016; Chotewutmontri and Barkan, 2018; Gawronski et al., 2018) and mitochondrial (Planchard et al., 2018; Kwasniak-Owczarek et al., 2019; Waltz et al., 2019; Tran et al., 2023) translational landscapes in Arabidopsis. With a single exception, all those studies performed Ribo-seq on total plant extracts; only Kwasniak-Owczarek et al. (2019) used isolated mitochondria. It should be emphasized that a very high read depth is required in case of mitochondrial RFs obtained from total plant extracts since they are vastly outnumbered by cytosolic and chloroplast RFs. According to the data published by Planchard et al. (2018), only about 2% of 900 million reads obtained from Arabidopsis flowers were mapped to mitochondrial mRNAs. The use of mitochondrial fractions instead of total cell extract improves the mapping efficiency to mitochondrial mRNAs (Kwasniak-Owczarek et al., 2019). However, even then, a percentage of RFs map to nuclear mRNAs, probably due to the presence of cytosolic ribosomes on the outer surface of mitochondria (Gold et al., 2017).

Ribosome profiling comprises two stages: ribosome footprinting and cDNA library construction (Wu and Hsu, 2022). Lysate preparation, ribonuclease digestion, monosome isolation, RF size selection, and rRNA depletion are essential steps in the first stage. Individual protocols applied to plant organelles differ in details of each of these steps, including the tissue treatment prior to harvest, type of RNase used to generate monosomes, and the method used to purify monosomes. Also the sequencing libraries can be prepared following different protocols. The approach used is often optimized to a particular plant material or to the specific questions addressed by the study.

Because ribosome profiling maps the exact positions of ribosomes on a transcript, it is imperative to immobilize the ribosomes on mRNA first. The simplest way is to arrest translation using antibiotics. However, studies in yeast and bacteria have revealed that in fact cycloheximide and chloramphenicol, respectively, alter the ribosome distribution at both the transcript and codon levels (Gerashchenko and Gladyshev, 2014; Mohammad et al., 2019). To avoid such artefacts flash-freezing the material in liquid nitrogen is recommended instead. Following this recommendation we have immobilized ribosomes by flash-freezing isolated mitochondria and then lysing them in the presence of antibiotics (cycloheximide to stop cytosolic ribosomes and chloramphenicol to stop chloroplast and mitochondrial ribosomes) (Kwasniak-Owczarek et al., 2019) while others have performed the whole procedure in the presence of antibiotics only (Chotewutmontri and Barkan, 2018; Gawronski et al., 2018; Planchard et al., 2018). The obtained lysate is treated with RNase, which is critical to the nucleotide-level resolution of the whole procedure. An excessive hydrolysis is likely to cause rRNA fragmentation and monosome breakdown, and therefore the use of an appropriate RNase and a careful determination of its minimal concentration and time of sample digestion is of the utmost importance. The RNase most widely used in plant organellar Ribo-seq procedures is RNase I (Gerashchenko and Gladyshev, 2017) since it cleaves after any nucleotide and displays little to no cutting preference. As an alternative, micrococcal nuclease S7 is suggested. This nuclease cleaves after all four nucleotides, but apparently has some obscure cutting preference. After the digestion, the protected ribosome-bound mRNA fragments are collected by sucrose gradient centrifugation in the monosome fraction. The next step, size selection, involves electrophoresis of purified RNA in a denaturating urea polyacrylamide gel and extraction of fragments of the required size. Different authors prefer somewhat different RF lengths, and there is no consensus in the literature regarding the ‘correct’ value. Thus, some studies have used RFs within a narrow size range (e.g. 27 and 33 nt) (Planchard et al., 2018), while others have used a broader range (e.g. 25–35 nt (Kwasniak-Owczarek et al., 2019) or even 16–42 nt (Gawronski et al., 2018).

Interestingly, both mitochondrial and chloroplast RFs showed a bimodal size distribution, with the shorter ones more numerous than the longer ones in mitochondria (Chotewutmontri and Barkan, 2016; Planchard et al, 2018; Kwasniak-Owczarek et al., 2019), and a similar abundance of both populations in chloroplasts (Zoschke et al., 2013; Chotewutmontri and Barkan, 2016; Ting et al., 2023, Preprint). It has been speculated that the two populations could reflect distinct conformations of the ribosomes, depending on the actual stage of translation (Rooijers et al., 2013; Lareau et al., 2014).

One must bear in mind that the RNase treatment not only produces RFs from mRNA but also leads to the generation of rRNA fragments that can co-purify with the RFs and become incorporated into the sequencing libraries, sometimes dramatically reducing the number of informative reads (Ting et al., 2023, Preprint). To eliminate or at least to reduce this effect the rRNA depletion step is carried out before library construction. The most common approach relies on the hybridization of samples following size selection with rRNA specific oligonucleotide probes using commercial rRNA depletion kits dedicated to RNA-seq experiments. However, these kits are often not effective enough because they contain a limited number of probes that mostly target highly conserved rRNA sequences only and are unlikely to match all rRNA fragments generated by RNase action (Ting et al., 2023, Preprint). Therefore, the residual rRNA-derived reads must be filtered out at the stage of the bioinformatic analysis.

Construction of the Ribo-seq library is the second stage of ribosome profiling, which is often accomplished with commercial kits designed for small RNA molecules (Chotewutmontri et al., 2018; Gawronski et al., 2018; Planchard et al., 2018; Kwasniak-Owczarek et al., 2019). These kits utilize oligonucleotides called adapters that are ligated to ends of small RNAs to allow for PCR amplification and to accurately identify them during sequencing. After construction, the libraries are quality-tested with a fragment analyser (Wu and Hsu, 2022). Following that, a deep-sequencing of the library is conducted, and bioinformatic analyses are performed on the sequencing results. While the standard bioinformatic approaches assume the use of in-house scripts and individual tools, nowadays, a comprehensive R toolkit, ORFik, is available and can be used both for processing and analysing of Ribo-seq data (Tjeldnes et al., 2021).

The initial step of bioinformatic analysis is the removal of adapter sequences and low-quality reads with tool such as Cutadapt (Martin, 2011). Bowtie2 is then used to filter away rRNA and tRNA reads since their excess can interfere with further quantification (Langmead and Salzberg, 2012). The next step is read mapping, a crucial procedure that determines the precise placement of each read within the reference genome. Splice-aware aligner tools like STAR and TopHat2 are among the mapping tools (Engström et al., 2013). The alignment is set up to allow a defined number of mismatches. In case of plant organellar RFs, usually two mismatches are preferred (Chotewutmontri and Barkan, 2016; Gawronski et al., 2018; Planchard et al., 2018). Ribo-seq data can be visualized using the IGV (Integrative Genomics Viewer) tool dedicated for interactive exploration of large, integrated genomic datasets (Robinson et al., 2011).

Ribo-seq reads should map mainly to protein coding sequences (CDS). They are unique or map to different locations. To prevent the overinterpretation of data, a conservative approach would be to analyse only uniquely mapped reads (Bartholomäus et al., 2016). Ribo-seq reads should also display a strong triplet (3 nt) periodicity, which corresponds to ribosomes decoding 3 nt per codon (Ingolia, 2016). The most straightforward method to calculate 3 nt periodicity is to count reads in all three possible frames (expected 0 frame, +1 frame, +2 frame). The calculation requires determination of the 5ʹ-P-site offset for every RF length species, which is the number of bases located 5ʹ to the peptidyl-site (P-site) of a respective RFs (Gotsmann et al., 2024). This can be done by riboWaltz software (Lauria et al., 2018). Then, the 5ʹ-offsets are used to calculate the exact P-site position and reading frame of every RFs using RiboTaper software (Calviello et al., 2016). A clear preference for a single reading frame is expected for RFs with strong 3 nt periodicity.

The number of Ribo-seq reads per CDS can be determined using a program such as featureCounts (Liao et al., 2014). The read count number should be normalized for sequencing depth and CDS length and expressed in one of the units RPKM (reads per kilobase million), FPKM (fragments per kilobase million), or TPM (transcript per million) (Conesa et al., 2016). Normalized read count can be used to estimate the relative translation efficiency of individual CDSs. The calculation of this parameter needs both Ribo-seq and RNA-seq data. The ratio of normalized RFs from Ribo-seq to mRNA counts from parallel RNA-seq within a CDS is used to compute the translation efficiency (Ingolia et al., 2009). Moreover, normalized read count can be applied to differential translation analyses using software designated for this task, such as Xtail (Xiao et al., 2016). So far, similar approaches have been used to determine the efficiency of translation of plant organellar CDSs in several species and conditions (Chotewutmontri and Barkan, 2016, 2018; Planchard et al., 2018; Kwasniak-Owczarek et al., 2019; Tran et al., 2023).

Notably, ribosome profiling can be used to obtain much more detailed information than the translational efficiency as it provides a single-nucleotide-resolution footprint of all the ribosomes associated with a given mRNA species. Moreover, the data obtained are of a quantitative character allowing the calculation of the statistics of the ribosome distribution along the mRNA. Thus, it could be revealed that mitoribosomes deficient in the S10 protein protect shorter mRNA fragments that also exhibit a weaker 3 nt periodicity compared with the wild-type (Kwasniak-Owczarek et al., 2019). The mapping of ribosomes on mRNA at a codon resolution allowed defining translation start and termination sites for mitochondrial mRNAs from which the translation initiation and termination codons are removed by post-transcriptional mRNA processing (Planchard et al., 2018). Also pause sites on Arabidopsis chloroplast mRNAs could be identified by comparing ribosome density on specific segments of the mRNA with the mean density across the coding region; this in turn indicated specific features of the mRNA and of the nascent polypeptide causing the ribosome pausing (Gawronski et al., 2018).

Yet another application of ribosome profiling involved a high resolution analysis of membrane-associated and soluble ribosome footprints to reveal the principles of co-translational targeting of proteins to the thylakoid membrane in maize (Zoschke and Barkan, 2015; Hristou et al., 2019). In this context it is worth mentioning that also unedited and unspliced mRNAs were found to be associated with ribosomes in plant mitochondria, suggesting that ribosomes do not discriminate between correctly processed and unprocessed mRNAs (Planchard, et al., 2018; Kwasniak-Owczarek et al., 2019). Interestingly, Rugen et al. (2023) found that proteins translated from non-edited transcripts can be incorporated into native mitoribosomes. It was shown that SSU ribosomal protein RPS3 is assembled into native plant mitoribosomes regardless of editing state, implicating the existence of heterogeneous mitoribosome. In turn, the study of Kwasniak-Owczarek et al. (2019) revealed that mitoribosomes lacking the S10 protein more frequently carried out translation of unspliced mRNAs than did wild-type mitoribosomes.

Despite the impressive refinement of the Ribo-seq technology since its introduction, which has substantially improved the ability to study plant organellar translation at a genomic level and with a single-nucleotide resolution, certain limitations remain. An incomplete fragmentation of polysomes to monosomes by RNase digestion can cause the transcript regions where ribosomes stack to be under-represented in the Ribo-seq library (Hou et al., 2016). Another major concern is that footprints on non-ribosomal RNA-binding proteins (RBPs) or RNA fragments refractive to digestion due to their secondary structure may be mistaken for true ribosomal footprints (Hsu et al., 2016). Indeed, putative RBP footprints in Ribo-seq data have been identified in non-coding regions of Arabidopsis mitochondrial mRNAs as short non-coding RNAs (sRNAs) (Kwasniak-Owczarek et al., 2019). Some of these mtRibo-seq-sRNAs resemble clustered organellar sRNAs (cosRNAs) that coincide with the 3ʹ termini of mature mitochondrial transcripts. With this in mind, to reconstruct the real translational status of the transcriptome, it is critical to select for analysis only RNA fragments that are bona fide ribosomal footprints. The true RFs display a 3 nt periodicity that is a hallmark of high-quality Ribo-seq data (Wu and Hsu, 2022). Furthermore, the technique cannot distinguish between terminating and initiating ribosomes in regions of adjacent reading frames in multicistronic transcripts, and consequently their individual translation rates cannot be determined with precision (Ting et al., 2023, Preprint).

A future methodological perspective for the study of plant organellar translation based on recently developed strategies

The benefits, limitations, and areas of applicability of the three methods used to monitor plant organellar translation covered in this review are summarized in Table 1 and presented schematically in Fig. 1. The choice of the most appropriate assay for a given application should be determined by the specific goals of the study.

Table 1. Benefits, limitations and optimal applications of plant organellar translatome analysis techniques

Method	Benefits	Limitations	Optimal applications	Selected references	
Radioisotope labelling	✓ Direct evaluation of newly synthesized proteins in organello (mitochondria and chloroplasts) or in vivo (chloroplasts)

✓ Useful in assessing the synthesis of highly expressed proteins

	✗ Hazards of working with radioactive isotopes

✗ Possible bacterial and cytoplasmic contamination of organellar samples

✗ The amount of labeled proteins is affected by their degradation

✗ No unambiguous identification of proteins

✗ Problems with detection of proteins with low expression levels

	Bulk translation quantification of newly synthesized proteins	Mitochondria:
Giegé et al. (2005), Kwasniak et al. (2013)
Chloroplasts:
Pesaresi et al. (2001, 2006)	
Polysome profiling	✓ Visualization of mRNA distribution across polysome fractions

✓ Determination of the proportion of ribosome-bound mRNA by ribosome occupancy analysis

	✗ Labor intensive

✗ Scaling issues

✗ Time-consuming analyses

✗ Specialized equipment

✗ Contamination of polysomal fractions by non-polysomal complexes

✗ Problems with differentiation between active and stalled/paused ribosomes

✗ Imprecise determination of the number of ribosomes per mRNA

✗ No information about the actual ribosome distribution along mRNA

	Determination of the translation status of a given mRNA/particular set of mRNAs
Initial screen for translational changes	Mitochondria:
Kwasniak et al. (2013), Hameed et al. (2017)
Chloroplasts:
Pesaresi et al. (2001), Kahlau and Bock (2008)	
Ribosome profiling	Determination of the ribosome number on mRNA

Determination of the actual location of ribosomes along mRNA

Insight into mechanism and dynamics of translation (e.g. detection of translational initiation, pausing, and termination sites)

Possible study of connection between translation and other gene expression processes

	✗ Labor intensive

✗ Time-consuming analyses

✗ Specialized equipment

✗ Highly stochastic action of ribonuclease and incomplete conversion of polysomes to monosomes

✗ Contamination of RFs by rRNA fragments

✗ PseudoRFs contamination; misidentification of footprints of non-ribosomal proteins associated with mRNA as RFs

✗ Extensive bioinformatic analyses

	Determination of mRNA translation efficiency on global scale
Determination of molecular mechanisms of translational control	Mitochondria:
Planchard et al. (2018), Kwasniak-Owczarek et al. (2019)
Chloroplasts:
Chotewutmontri and Barkan (2018), Gawronski et al. (2018)	

Fig. 1. Experimental approaches to the investigation of plant organellar translation. Plant tissue or isolated organelles (mitochondria/chloroplasts) can be used as a starting material. Radioisotope labeling (left) utilizes the incorporation of 35S-labeled methionine into newly synthesized proteins. Following separation by SDS-PAGE radiolabeled proteins are visualized by autoradiography. Polysome profiling (middle) relies on separation of mRNA–ribosomal complexes according to their size using ultracentrifugation in a sucrose gradient. RNA and proteins are isolated from collected fractions and used for targeted or global analysis of mRNA and/or ribosomal proteins. Ribosome profiling (right) employs RNase digestion to degrade ribosome-free mRNA regions and reveal ribosome footprints (RFs). The RFs are isolated and deep sequenced or analysed by microarray hybridization.

The use of mass spectrometry (MS)-based analysis to detect directly and quantify newly synthesized proteins may be the direction of future technical advances in plant mitochondrial and chloroplast translation studies. Conceptually, this is the most straightforward way of quantifying translation. It should be noted, however, that mass spectrometric identification of organellar proteins could be impeded by their largely hydrophobic nature and relatively low abundance, particularly of mitochondrial proteins. Despite that, one of the MS-based methods, namely bio-orthogonal non-canonical amino acid tagging (BONCAT), has allowed the identification of some newly synthesized chloroplast-encoded and mitochondrially encoded polypeptides in Arabidopsis (Glenn et al., 2017). The assay relies on specific tagging of nascent polypeptides synthesized in native plant tissue by incorporation of a non-canonical amino acid (the methionine surrogate azidohomoalanine) followed by their isolation via affinity chromatography and MS analysis. A cautionary note is warranted here: among all the identified proteins, only 23 chloroplast-encoded and two mitochondrially encoded ones were found (Glenn et al., 2017). So far, this technique has been used in a single plant study only and has never been applied to investigate specifically the translatomes of plant mitochondria or chloroplasts. However, a similar approach—pulse stable isotope labeling by amino acids in cell culture (pSILAC)—was used by Imami et al. (2023) for a direct detection of proteins produced in human mitochondria. Imami et al. combined pSILAC with the isolation of the mitochondrial fraction and achieved a nearly 100% identification rate of mitochondrially encoded proteins (12 of the 13 encoded in the human mitochondrial genome). Paradoxically, SILAC-based approaches, widely used in yeast and mammalian systems, have been deemed inappropriate for plants due to poor metabolic labeling efficiency impairing the reliability of peptide ratio quantification (Gruhler et al., 2005). Nevertheless, it is worth mentioning that SILAC has been successfully utilized to determine the turnover rate of mitochondrial proteins in enriched mitochondrial fractions isolated from Arabidopsis cell cultures (Nelson et al., 2013).

Another approach for analysing translation by measuring the incorporation of labeled amino acids relies on in vitro transcription/translation systems. Since there is no available translation-competent lysate for mitochondria, an option to assess specific mitochondrial translation steps is to reconstitute an in vitro translation system from the single components (Kummer et al., 2021). By using such in vitro system the unique features of translation initiation in human mitochondria have been described (Remes et al., 2023). It has been shown that leaderless mRNAs (without the 5ʹ leader sequences), characteristic for human mitochondria, were loaded directly onto assembled mitoribosomes, but not onto the SSU, in a manner that requires initiator fMet–tRNAMet binding. So far, the in vitro translation system has not been reported for plant mitochondria. However, recently the initiation of translation in plant mitochondria was successfully investigated using two different RNA–protein interaction assays: RNA electrophoretic mobility shift assay (REMSA) and RNA immunoprecipitation sequencing (RIP-seq) (Tran et al., 2023). It has been demonstrated that mTRAN1 protein, a component of SSU, binds to A/U-rich motifs located in the 5ʹ untranslated regions of plant mitochondrial mRNAs, which act as ribosome binding sites. In this context mTRAN1 is a homing factor to guide the mitoribosome to mRNAs and initiate translation.

New and interesting findings about the mechanics of plant organellar translation may be also deduced from cryo-electron microscopy (cryo-EM) analyses of ribosomes. Structural data obtained for plant mitoribosome (Waltz et al., 2020) suggested that plant-specific rRNA segments and ribosomal proteins are involved in the mitochondrial membrane association and mRNA recruitment prior to translation initiation. In turn, cryo-EM study of chlororibosomes revealed that some of the chloroplast-specific ribosomal proteins that are located in the SSU have a specialized functions in translation (Ahmed et al., 2017). For example, the mRNA exit site is highly remodeled due to the unique localization of the cS23, suggesting a different mechanism of translation initiation in the chloroplast that may function in addition to the canonical bacterial pathway. Noteworthy, the cryo-EM reconstructions of human mitoribosomes help to explain how mitoribosomal proteins stabilize binding of mRNA and tRNAs during translation (Singh et al., 2023, Preprint). It has been demonstrated that mitochondria-specific proteins direct the mRNA to the decoding center, where it is aligned with a tRNA, while one of the SSU proteins, mS29, stabilizes the inter-subunit communication. Although such detailed mechanistic aspects of translation have not yet been reported in plant organelles, it is believed that the improvement of resolution of cryo-EM reconstructions of their ribosomes combined with biochemical analyses can contribute to filling this gap.

It should also be highlighted that all the assays currently used to study translation in plant organelles lack cell-type specificity. Thus, the ability to visualize organellar protein synthesis in a small number of cells or even a single cell is an obvious goal to be achieved in plant organellar translation research. Such methodology, called mitochondrial fluorescent non-canonical amino acid tagging combined with fluorescence-activated cell sorting (mito-FUNCAT-FACS) has been recently introduced to investigate mitochondrial translation in mammalian cells (Kimura et al., 2022; Saito et al., 2023). It is based on labeling of nascent peptides with a methionine analog, l-homopropargylglycine. In the presence of cytosolic translation inhibitors, l-homopropargylglycine is incorporated selectively into polypeptides generated by mitoribosomes. The labeled peptides are then conjugated to a fluorophore and the signal is detected in individual cells by flow cytometry. To date, this strategy has not been used to track organellar translation in plants.

Another type of experimental strategy for monitoring organellar translation relies on in vivo visualization of translating ribosomes. This approach, called ribo-bimolecular fluorescence complementation (Ribo-BiFC), was first employed to visualize ribosomes in Drosophila axons (Singh et al., 2020) and recently used to monitor mitoribosomes in live human cells (mtRibo-BiFC) (Lee et al., 2022). The mitoribosomal proteins MRPL2 and MRPS6, located at the intersubunit bridge of the ribosome, were tagged with complementary halves of a fluorescent protein (mVenus) and served as a BiFC pair (Lee et al., 2022). The two non-functional halves of mVenus could make a stable contact and thereby form a functional complex only when the SSU and LSU were assembled to form a complete ribosome at initiation, so the emission of fluorescence indicated that translation initiation had occurred (Al-Jubran et al., 2013). Thus, the mtRibo-BiFC signal represents actively translating mitoribosomes exclusively. A similar approach should be applicable to plant systems. However, such real-time visualization of plant organellar ribosomes has the obvious limitation of requiring the production of transgenic plants or a transient expression of tagged components of ribosomes.

Acknowledgements

We are grateful to Dr Jan Fronk (Faculty of Biology, University of Warsaw, retired) for critical reading of the manuscript.

Author contributions

MK-O wrote the manuscript with input from HJ. MK-O designed and revised the manuscript, and prepared the figure and table.

Conflict of interest

The authors declare no conflict of interest.

Funding

This work was supported by grant 2021/41/B/NZ3/00571 to HJ from the National Science Centre, Poland. Open access publication of this article was financed by a subsidy for the Faculty of Biotechnology, University of Wroclaw.
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