The conceptual starting point is a fundamental limitation of cellular protein synthesis. In living cells, ribosomes and messenger RNAs meet largely by random collision, with translation initiation governed by base-pairing between the Shine-Dalgarno sequence at the start of an mRNA and the anti-Shine-Dalgarno tail of the 16S rRNA. This stochastic encounter system is remarkably versatile, but it makes it nearly impossible to optimize the entire translation apparatus for the production of one specific protein, a capability that would be enormously valuable in biotechnology, where engineered cells are often asked to churn out a single high-value polypeptide. The Illinois team asked whether the ultimate solution would be to eliminate the encounter problem altogether by fusing the gene to the machine itself.

To test this idea, the researchers constructed what they call a chimeric messenger-ribosomal RNA, or mrRNA. They appended a protein-coding sequence to the 3′ end of the 16S rRNA, creating a single RNA molecule that contains both the structural information needed to build a small ribosomal subunit and the genetic information needed to encode a protein. The design drew on decades of work on translation initiation in bacteria, including the classic discovery by Shine and Dalgarno in 1974 that the 3′-terminal sequence of 16S rRNA pairs with ribosome binding sites on mRNAs. By placing a coding region downstream of the rRNA proper, separated by a short linker, the team created a transcript in which the ribosome’s own RNA carries an open reading frame.

The critical question was whether this hybrid molecule would behave. Ribosomal RNA is heavily processed, folded, modified and assembled with dozens of proteins into functional subunits, and there was no guarantee that an rRNA carrying a bulky protein-coding appendage would survive the cell’s ribonucleases or assemble into a competent particle. The team built a library of 16S rRNA-mRNA fusions and screened for constructs in which the mrRNA was incorporated into 30S small subunits and, subsequently, into complete 70S ribosomes. Their experiments showed that the chimeric RNA was indeed processed at its ends, assembled with ribosomal proteins, and incorporated into small subunits that could bind the large subunit to form functional monosomes, which the researchers named Ribo-M.

Proof that Ribo-M actually translates its own RNA came from a series of stringent controls. The team showed that the mrRNA-encoded proteins, including variants of green fluorescent protein and the small HiBiT peptide tag used in luminescence assays, were produced only when the small subunit was functional. Mutations known to inactivate the decoding center of the 16S rRNA, such as the G530A mutation in helix 18, abolished production of the mrRNA-encoded protein, as did antibiotics that impair small subunit function, such as spectinomycin. Crucially, these same mutations did not prevent the mrRNA from being incorporated into ribosomes, ruling out the alternative explanation that the protein was being made by ordinary ribosomes reading a stray mRNA fragment released from the construct. The translation, in other words, was carried out in cis, by ribosomes assembled on the very RNA molecule that encodes the protein.

The team pushed the concept further by combining the mrRNA design with Ribo-T, a previously engineered ribosome in which the small and large subunits are tethered together by a linker RNA, allowing fully orthogonal translation systems to operate inside cells. When the protein-coding sequence was integrated into the rRNA of the tethered ribosome, the result was Ribo-TM, a single RNA scaffold that unites three functions never before combined in one molecule: encoding a protein, decoding genetic information, and catalyzing peptide bond formation. In Ribo-TM, the ribosome is simultaneously the factory, the foreman and the blueprint, a self-contained unit in which the message and the machine are physically inseparable.

Beyond its synthetic biology applications, the work speaks directly to one of the deepest puzzles in evolutionary biology. In RNA world models of life’s origin, the earliest translation systems would have faced a chicken-and-egg problem: proteins are needed to make the translation machinery efficient, but the machinery is needed to make proteins. Explaining how early protein synthesis could function reliably despite the scarcity and poor organization of its components has been a long-standing challenge. The demonstration that a ribosome can translate its own RNA shows that a single RNA molecule could, in principle, have served simultaneously as structural scaffold and genetic template, providing a plausible bridge between an RNA-only world and the protein-based biochemistry that followed. The finding resonates with recent theoretical proposals about the origin of the ribosome, including hypotheses about selfish ribosome lineages and symbiotic origins published in 2026.

The technical achievement required overcoming formidable obstacles in ribosome assembly and RNA processing. The researchers used engineered Escherichia coli strains lacking native ribosomal RNA operons, allowing them to control which rRNA variants populated the cell’s ribosomes. They employed primer extension analysis to distinguish mrRNA from native 16S rRNA, sucrose gradient fractionation to track assembly into subunits and polysomes, and chemical probing with dimethyl sulfate to verify that the chimeric RNA adopted a native-like fold within the ribosome. Mapping experiments showed that the mrRNA’s 5′ and 3′ ends were processed in ways that preserved both the mature rRNA structure and the integrity of the appended coding sequence, suggesting that the cell’s RNA processing machinery can accommodate these hybrid transcripts.

The practical implications are considerable. Because a Ribo-M or Ribo-TM ribosome is committed to translating a single, internally encoded protein, the system offers a versatile platform for orthogonal protein production, in which a dedicated subpopulation of ribosomes manufactures one product with maximal efficiency and minimal interference from the cell’s own translation program. This builds on a lineage of engineered translation systems, from the specialized ribosomes of Hui and de Boer in 1987 to the orthogonal ribosome-mRNA pairs developed by Rackham and Chin in 2005 and the tethered-subunit ribosomes introduced by Orelle and colleagues in 2015. Each generation has tightened the coupling between ribosome and message; the new work completes the arc by making them one and the same molecule. Such systems could prove especially valuable for incorporating non-canonical amino acids into proteins, expanding the genetic code for new chemistries, and for cell-free protein synthesis platforms where dedicated production lines are at a premium.

Cautions remain before the technology can be widely deployed. The fraction of cellular ribosomes carrying mrRNA is limited, translation levels from the chimeric RNA are modest compared with native mRNA-driven expression, and the researchers note that the work establishes mechanistic feasibility rather than a production-ready system. The authors have filed a pending patent application covering the Ribo-M and Ribo-TM designs, and the key plasmids and strains are being made available through Addgene, which should accelerate adoption across the synthetic biology community. Still, the conceptual breakthrough is unmistakable. A ribosome that reads its own RNA is no longer just a molecular machine but a self-referential one, and in demonstrating that such a construct can live, assemble and translate inside a bacterial cell, the Illinois team has both expanded the engineering toolkit for protein production and offered experimental support for a compelling scenario in which life’s first proteins were made by ribosomes translating themselves.

Subject of Research: Engineering ribosomes that translate proteins encoded within their own ribosomal RNA

Article Title: Rewiring the ribosome to translate proteins encoded in its own RNA

Article References: Alizadeh, K., Klepacki, D., Vázquez-Laslop, N., & Mankin, A. S. (2026). Rewiring the ribosome to translate proteins encoded in its own RNA. Nature. https://doi.org/10.1038/s41586-026-10962-1

Image Credits: AI Generated

DOI: 10.1038/s41586-026-10962-1

Keywords: ribosome, translation, ribosomal RNA, synthetic biology, mRNA, Ribo-M, Ribo-T, orthogonal translation, RNA world, origin of life, protein synthesis, molecular engineering

A new laboratory study, published in MicrobiologyOpen, suggests that a solution may lie in one of the most abundant molecules inside every living cell: ribosomal RNA. Previous work on prokaryotes demonstrated that when viruses lyse bacterial cells, substantial amounts of rRNA are released into the extracellular medium, where it can be detected and quantified. Building on that observation, researchers developed an approach called Mortality by Ribosomal Sequencing, or MoRS, which compares ribosomal RNA sequences in cellular and dissolved fractions of seawater to estimate taxon-specific cell lysis. What remained untested was whether the concept could be extended to eukaryotic microalgae, whose complex internal organization and diverse viral enemies make such an extrapolation far from trivial.

To test the idea, a research team led by investigators working with marine algal virus systems conducted controlled infection experiments with two contrasting host-virus pairs. The first was the diatom Chaetoceros tenuissimus, a small coastal phytoplankton species less than ten micrometers across that is susceptible to Chaetoceros tenuissimus RNA virus type II, a small single-stranded RNA virus with particles roughly 22 to 38 nanometers in diameter. The second was the raphidophyte Heterosigma akashiwo, a globally distributed, flagellated species notorious for forming harmful algal blooms, which was challenged with HaV strain 120, a large double-stranded DNA virus with strain-specific infectivity. Together, these systems span different host physiologies and viral genome types, providing a rigorous framework for testing whether cell-free rRNA behaves as a general lysis marker in eukaryotes.

The experimental design was carefully controlled. Host cultures were grown at 22 degrees Celsius under a 12-hour light and 12-hour dark cycle in sterile artificial seawater media and acclimated for multiple generations before infection. For each host-virus system, triplicate flasks received virus at a multiplicity of infection of ten, while triplicate control flasks received none. The cultures were incubated for six days and sampled daily. To track viral proliferation, the team used the most probable number method to determine infectious virus titers at the beginning and end of the incubation. Critically, they also added purified Escherichia coli ribosomes to every flask as spike-in controls, at a concentration of approximately 2.9 times ten to the seventh copies per milliliter, allowing them to measure how quickly free rRNA degrades in the medium.

Separating the cellular from the dissolved fraction was achieved by gentle filtration through 0.22-micrometer polycarbonate filters under low vacuum. RNA retained on the filter was operationally defined as the cellular fraction, while RNA passing through was defined as cell-free. Cellular RNA was extracted with a silica-column kit enhanced by bead-beating, whereas cell-free RNA was concentrated from twenty-milliliter filtrates using a column-based vacuum system optimized for large water volumes. After genomic DNA removal and complementary DNA synthesis, absolute quantification was performed with a digital PCR system, using newly designed primers targeting the V4 region of the algal 18S rRNA gene and primers specific to the major capsid protein genes of each virus. Extraction efficiencies of 21.9 percent for the cellular fraction and 83.1 percent for the cell-free fraction were used to correct the measured concentrations.

The growth dynamics of the two hosts diverged sharply. In the Chaetoceros experiment, cells in the virus-added treatment grew at rates comparable to controls until day three but then grew significantly more slowly, while cell density, chlorophyll a, and cellular rRNA all continued to rise until day three or four. In striking contrast, Heterosigma cultures declined immediately after viral addition, with most cells losing motility within 24 hours, echoing earlier reports on this host-virus system. Infectious virus particles increased dramatically in both systems, from 5.8 times ten to the fifth to 1.9 times ten to the eighth most probable number units per milliliter for the diatom RNA virus, and from 7.6 times ten to the fifth to 1.4 times ten to the seventh for the Heterosigma DNA virus, confirming active viral replication.

The central result concerned the redistribution of rRNA between particulate and dissolved pools. In the Chaetoceros experiment, cell-free 18S rRNA remained low in controls, below about 4 times ten to the eighth copies per milliliter, but rose sharply in infected flasks to a maximum of 3.7 times ten to the ninth copies per milliliter on day four. Per-cell rRNA content varied up to 2.8-fold within each condition but showed no significant treatment effect, and cellular rRNA correlated strongly with cell density in both species, with Pearson correlation coefficients of 0.72 and 0.94. These findings established that rRNA abundance tracks plankton biomass at the population scale, while its release into the dissolved phase signals cell rupture.

To convert rRNA dynamics into lysis rates, the team built a rate-based model describing daily changes in cell-free rRNA concentration. The model rests on first-order degradation kinetics, an assumption validated by the spike-in ribosomes, which decayed exponentially with rate constants of roughly 1.78 to 2.16 per day in the Chaetoceros experiment, corresponding to half-lives of 7.7 to 9.3 hours. By rearranging a simple mass-balance equation, the researchers solved for the cell-free rRNA production rate, a direct proxy for cell lysis. Normalizing this production rate by host cell abundance yielded a metric they call cell-free rRNA production per cell. In the Chaetoceros system, this metric was 46.1-fold higher in infected treatments than controls between days two and three, and in the Heterosigma system it reached 302.1-fold higher between days three and four, with maximum per-cell release rates up to 46.1- and 302.1-fold above controls across the experiments.

The two host-virus systems also revealed fundamentally different infection strategies. In Chaetoceros, viral marker genes accumulated progressively in the cellular fraction over days one to four, and lysis peaked between days two and three even as the population was still growing, demonstrating that active viral lysis can occur during the apparent growth phase of a bloom. In Heterosigma, elevated per-cell rRNA release appeared both early, during days zero to one, and late, during days three to five. The early mortality could not be explained by canonical viral lysis, since viral capsid gene expression peaked only on day two. The authors suggest several possible mechanisms, including abortive infection, membrane destabilization during viral entry, a phenomenon analogous to lysis from without described in bacteriophage systems, or damage caused by bacteria introduced with the nonaxenic viral lysate, which flow cytometry confirmed had proliferated to high densities in infected treatments.

The implications reach well beyond the laboratory. Unlike dilution assays, which are labor-intensive and lack taxonomic resolution, cell-free rRNA quantification combined with high-throughput sequencing could simultaneously assess lysis across diverse taxa in natural communities, offering a taxon-resolved measure of lytic mortality that plankton ecologists have long sought. Because viral lysis is estimated to contribute up to roughly 45 percent of the labile dissolved organic carbon pool in marine systems, better quantification of who is being lysed, and when, could sharpen models of carbon cycling and microbial loop dynamics. The authors caution that the approach has limits: ribosome content per cell varies several-fold, infected bacteria can actively reduce their ribosome pools, degradation constants differ among seawater samples and must be measured for each one, and nonviral processes such as sloppy feeding, parasitism, and algicidal bacteria can also release intracellular rRNA. They also note that the degradation kinetics of E. coli 16S rRNA may not perfectly match those of eukaryotic 18S rRNA, calling for spike-in standards derived from representative plankton. Even with these caveats, the study provides a mechanistic foundation for reading cell death directly from the RNA dissolved in seawater, turning a ubiquitous molecule into a quantitative witness of the viral shunt at work.

Subject of Research: Quantifying viral lysis of eukaryotic microalgae using cell-free ribosomal RNA as a molecular marker

Article Title: Quantifying Viral Lysis in Microalgae Using Cell‐Free rRNA

Article References: Kikuya, S., Tomaru, Y., Nagasaki, K., Morimoto, D., Yamagishi, Y., Ogata, H., & Endo, H. (2026). Quantifying Viral Lysis in Microalgae Using Cell‐Free rRNA. MicrobiologyOpen, 15(5), Article e70402. https://doi.org/10.1002/mbo3.70402

Image Credits: AI Generated

DOI: 10.1002/mbo3.70402

Keywords: viral lysis, microalgae, ribosomal RNA, phytoplankton, viral shunt, marine viruses, Chaetoceros tenuissimus, Heterosigma akashiwo, MoRS, digital PCR, plankton mortality, biogeochemistry

A new study published in MicrobiologyOpen has now combined two cutting-edge technologies, CRISPR interference and nanopore native RNA sequencing, to systematically probe the epitranscriptome of Escherichia coli. The research, led by Miranda E. Pitt of the University of Technology Sydney and Lachlan J. M. Coin of the University of Melbourne, together with colleagues, demonstrates a workflow in which individual RNA modification enzymes are switched off in living bacteria, and the resulting loss of chemical marks is read out directly at the single-molecule level. The approach not only confirmed several well-known modification sites on ribosomal RNA but also revealed candidate modification sites scattered across bacterial messenger RNA, a frontier that has remained largely unmapped.

The technical challenge the team faced is formidable. Traditional methods for detecting RNA modifications, such as immunoprecipitation or chemical treatments like bisulfite conversion, are laborious, require highly purified and concentrated RNA, and generally capture only one modification type at a time. Bacterial RNA adds further complications: rigorous lysis procedures and complex cell membranes accelerate RNA degradation, and bacterial transcripts have notoriously short half-lives, in many cases under one minute. Enzymes called RNases can rapidly destroy extracted RNA unless inhibited immediately. Direct RNA sequencing, released by Oxford Nanopore Technologies in 2017, offers a faster alternative because it reads native RNA molecules as they pass through protein nanopores, preserving the chemical modifications that conventional methods destroy when RNA is converted to complementary DNA. However, the chemistry was originally incompatible with bacterial transcripts, which lack the polyadenylated tails found in eukaryotic messenger RNA.

The researchers overcame this barrier by artificially adding poly(A) tails to bacterial transcripts before sequencing, a technique they had refined in previous work. To create the perturbations needed to detect modifications, they turned to CRISPR interference, or CRISPRi. Unlike standard CRISPR gene editing, which cuts DNA, CRISPRi uses a catalytically inactive form of the Cas9 protein, dCas9, that binds to a target gene guided by a matching guide RNA and simply blocks its transcription. This knocks down, rather than knocks out, gene expression, avoiding the toxicity and off-target DNA disruption associated with genome cutting while allowing inducible, tunable silencing. The team used a plasmid system optimized for E. coli, in which dCas9 expression is triggered by the small molecule anhydrotetracycline.

Five genes encoding known ribosomal RNA modification enzymes were targeted: rlmF and rlmJ, both responsible for N6-methyladenosine marks on 23S ribosomal RNA; rsmF, which deposits 5-methylcytosine on 16S ribosomal RNA; rsmG, which installs 7-methylguanosine on 16S ribosomal RNA; and rluD, a pseudouridine synthase that modifies three positions in a functionally critical stem-loop of the 23S subunit. A sixth gene, add, encoding adenosine deaminase, was included to explore whether it might carry previously unrecognized RNA editing activity. The knockdown system was tested across multiple E. coli reference strains, including the type strain ATCC 11775, ATCC 8739, ATCC 25922, and a multidrug-resistant isolate, ATCC BAA-2452. Quantitative reverse transcription PCR confirmed that most target genes were repressed by more than 80 percent, although rsmG proved harder to silence, reaching only about 54 percent reduction, and the resistant isolate could not be transformed with the plasmid without a severe fitness cost.

With knockdown strains in hand, the researchers sequenced both total RNA and ribosomal RNA-depleted, messenger RNA-enriched samples on nanopore flow cells, using computational tools to compare ionic current signals between knockdown and control conditions. The software nanocompore, which does not require prior training and is agnostic to modification type, compared each perturbed sample against two controls: the unmodified original isolate and a plasmid control expressing dCas9 without a guide RNA. For rluD, all three known pseudouridine sites in the 23S ribosomal RNA were detected with strong statistical significance, and the sequence motifs surrounding those sites matched published reports. The rsmG knockdown likewise produced a clear signal at the known 7-methylguanosine position in 16S ribosomal RNA. In contrast, the sites modified by rlmF, rlmJ, and rsmF yielded weak signals that were difficult to distinguish from noise, echoing the findings of earlier benchmarking studies showing that certain modification types, particularly 5-methylcytosine and some N6-methyladenosine sites, are inherently harder to detect with current nanopore basecalling and current-difference tools.

Extending the analysis to messenger RNA, the team identified significant modification-site changes on several bacterial transcripts shared across strains. Genes harboring candidate modification sites included ompC, which encodes an outer membrane porin; lpp1; salPB; cspC; dbhA and dbhB; and secY, which encodes a core component of the protein secretion machinery. For the rluD and rsmG knockdowns, which had produced the strongest ribosomal RNA signals, the researchers searched messenger RNA for sequence motifs matching those seen at the known ribosomal sites. Strikingly, a significant change was found on the well-characterized pseudouridine motif in the gene encoding elongation factor Tu, the workhorse protein of translation, as well as on transcripts of lpp1, gapA, and fusA. For rsmG, significant sites matching the known 7-methylguanosine motif appeared in hupA, lpp1, rplT, rpsA, and ompC. Because these messenger RNA sites were detected in more than one strain background, they represent plausible, though not yet chemically verified, targets of bacterial modification enzymes acting beyond the ribosome.

The study also examined the downstream consequences of losing these modifications, measuring bacterial growth and profiling the proteome by quantitative mass spectrometry. Growth effects varied by strain: activation of dCas9 itself imposed a measurable burden in ATCC 11775, which carries an extra plasmid, while ATCC 8739 tolerated the system well. Statistically significant growth delays were observed for knockdowns of rluD and rsmF in one strain and rlmF, rluD, and rsmF in another, consistent with prior reports that the phenotypic impact of ribosomal RNA modification loss depends heavily on genetic background. Proteomic analysis detected roughly 1,800 to 2,400 proteins per strain and revealed modest but reproducible changes, including alterations in citrate lyase, the trehalose-specific phosphotransferase component TreB, the xylose-binding protein XylF, the sodium/proline symporter PutP, formate hydrogenlyase subunit HycB, and the oligopeptide transporter OppC. The authors note these changes require further validation and may only become pronounced under stress conditions such as antibiotic exposure or temperature shifts, which previous work has shown can reshape the bacterial epitranscriptome.

The broader significance of the work lies in its implications for antimicrobial resistance and bacterial virulence. RNA modifications are already known to help pathogens evade entire antibiotic classes, including aminoglycosides, chloramphenicol, and macrolides, and to contribute to antibiotic tolerance in organisms such as Vibrio cholerae. Loss of tRNA modification enzymes can impair a bacterium’s ability to infect and cause disease in a host. A reliable, rapid, genome-wide method for mapping modification dynamics in bacteria could therefore illuminate how pathogens respond to clinical interventions at the level of their RNA. The authors caution that fine-tuning is still needed, particularly for messenger RNA modification detection across diverse bacterial species and under varying growth conditions, and newer nanopore chemistry promises higher accuracy and greater yield. Yet as a proof of principle, pairing CRISPR interference with native RNA sequencing offers a scalable blueprint for interrogating the bacterial epitranscriptome, one silenced enzyme and one molecule of RNA at a time.

Subject of Research: Mapping RNA modifications in the Escherichia coli epitranscriptome using CRISPR interference gene knockdown and nanopore direct RNA sequencing

Article Title: Interrogating the Escherichia coli Epitranscriptome Via CRISPR Interference and Nanopore Native RNA Sequencing

Article References: Pitt, M. E., Zhang, J., Nguyen, A. N. T., Hall, M. B., Jebeli, L., Featherstone, L. A., Myers, G. S. A., Scott, N. E., & Coin, L. J. M. (2026). Interrogating the Escherichia coli Epitranscriptome Via CRISPR Interference and Nanopore Native RNA Sequencing. MicrobiologyOpen, 15(5), Article e70413. https://doi.org/10.1002/mbo3.70413

Image Credits: AI Generated

DOI: 10.1002/mbo3.70413

Keywords: epitranscriptome, E. coli, CRISPR interference, nanopore sequencing, direct RNA sequencing, RNA modifications, ribosomal RNA, messenger RNA, pseudouridine, m6A, antimicrobial resistance, MicrobiologyOpen